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Volume 143

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Number 5

November 24, 2010

Muscle Aging Cues Systemic AgingMuscle Aging Cues Systemic Aging

Reviews: Posttranslational ModifiReviews: Posttranslational Modificationscations

Muscle Aging Cues Systemic Aging

Reviews: Posttranslational Modifications

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Leading EdgeCell Volume 143 Number 5, November 24, 2010

IN THIS ISSUE

SELECT

657 Cell Cycle

PREVIEWS

665 ER Sheets Get Roughed Up C. Barlowe

667 SIRT3 in Calorie Restriction:Can You Hear Me Now?

C. Sebastian and R. Mostoslavsky

669 ATP Consumption PromotesCancer Metabolism

W.J. Israelsen and M.G. Vander Heiden

ESSAYS

672 Glycomics Hits the Big Time G.W. Hart and R.J. Copeland

677 What Determines the Specificityand Outcomes of Ubiquitin Signaling?

F. Ikeda, N. Crosetto, and I. Dikic

MINIREVIEW

682 Ubiquitin: Same Molecule,Different Degradation Pathways

M.J. Clague and S. Urb�e

PERSPECTIVE

686 Will the Ubiquitin System Furnish asMany Drug Targets as Protein Kinases?

P. Cohen and M. Tcherpakov

REVIEWS

694 Pathogen-Mediated PosttranslationalModifications: A Re-emerging Field

D. Ribet and P. Cossart

703 Modifications of Small RNAsand Their Associated Proteins

Y.-K. Kim, I. Heo, and V.N. Kim

SNAPSHOT

848 The SUMO System S. Creton and S. Jentsch

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ono-clonal antibodies are produced under license (granting certain rights including those under U.S. Patents No. 5,675,063 and 7,429,487).

TOP IMAGE: To the right, the nuclear pore complex is located in the nuclear double bilayer. To the left, nuclear proteins are interspersed between DNA and nucleosomes in various levels of compact-ness. Red = histone H2A.X; blue = histones H2B, H3, H4; green = p53 (with nearby ATM below); purple = Rb (with a nearby E2F dimer on the right); orange = damage/repair MRN complex, loaded onto a DNA double strand break. Please visit www.cellsignal.com for the complete story.

Confocal IF analysis of HeLa (upper) and HT-29 cells (lower), untreated (left) or UV-treated (right), using Phospho-Histone H2A.X (Ser139) (20E3) Rabbit mAb #9718 (green, upper) or Phospho-p53 (Ser15) (16G8) Mouse mAb #9286 (green, lower). Actin filaments were labeled with DY-554 phalloidin (red).

untreated +UV

HT-2

9He

La

phospho-histone H2A.Xphospho-p53

ArticlesCell Volume 143 Number 5, November 24, 2010

711 The ER UDPase ENTPD5 Promotes ProteinN-Glycosylation, the Warburg Effect,and Proliferation in the PTEN Pathway

M. Fang, Z. Shen, S. Huang, L. Zhao, S. Chen,T.W. Mak, and X. Wang

725 Stepwise Histone Replacement by SWR1Requires Dual Activation with HistoneH2A.Z and Canonical Nucleosome

E. Luk, A. Ranjan, P.C. FitzGerald, G. Mizuguchi,Y. Huang, D. Wei, and C. Wu

737 Sororin Mediates Sister ChromatidCohesion by Antagonizing Wapl

T. Nishiyama, R. Ladurner, J. Schmitz, E. Kreidl,A. Schleiffer, V. Bhaskara, M. Bando, K. Shirahige,A.A. Hyman, K. Mechtler, and J.-M. Peters

750 Nonenzymatic Rapid Controlof GIRK Channel Functionby a G Protein-Coupled Receptor Kinase

A. Raveh, A. Cooper, L. Guy-David, and E. Reuveny

761 Sequence-Dependent Sortingof Recycling Proteins by Actin-StabilizedEndosomal Microdomains

M.A. Puthenveedu, B. Lauffer, P. Temkin, R. Vistein,P. Carlton, K. Thorn, J. Taunton, O.D. Weiner,R.G. Parton, and M. von Zastrow

774 Mechanisms Determining the Morphologyof the Peripheral ER

Y. Shibata, T. Shemesh, W.A. Prinz, A.F. Palazzo,M.M. Kozlov, and T.A. Rapoport

789 Abortive HIV Infection MediatesCD4 T Cell Depletion and Inflammationin Human Lymphoid Tissue

G. Doitsh, M. Cavrois, K.G. Lassen, O. Zepeda,Z. Yang, M.L. Santiago, A.M. Hebbeler,and W.C. Greene

802 Sirt3 Mediates Reduction of OxidativeDamage and Prevention of Age-RelatedHearing Loss under Caloric Restriction

S. Someya, W. Yu, W.C. Hallows, J. Xu,J.M. Vann, C. Leeuwenburgh, M. Tanokura,J.M. Denu, and T.A. Prolla

813 FOXO/4E-BP Signaling in DrosophilaMuscles Regulates Organism-wideProteostasis during Aging

F. Demontis and N. Perrimon

(continued)

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C

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AD7.pdf 7/24/2008 12:07:29 PM

826 Reelin and Stk25 Have OpposingRoles in Neuronal Polarizationand Dendritic Golgi Deployment

T. Matsuki, R.T. Matthews, J.A. Cooper,M.P. van der Brug, M.R. Cookson, J.A. Hardy,E.C. Olson, and B.W. Howell

RESOURCE

837 A Human Genome Structural VariationSequencing Resource Reveals Insightsinto Mutational Mechanisms

J.M. Kidd, T. Graves, T.L. Newman, R. Fulton,H.S. Hayden, M. Malig, J. Kallicki, R. Kaul,R.K. Wilson, and E.E. Eichler

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On the cover: The progressive decrease of muscle strength in the elderly is one of the first

signs of aging in many organisms. Here, Demontis and Perrimon (pp. 813–825) demonstrate

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stasis and muscle function and is beneficial for systemic aging by extending life span. The

image is a view of old Drosophila flight muscles with highlighted nuclei (white), myofibrils

(blue), protein aggregates (red), and mitochondria (green).

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Laser-based FluorescenceImaging: What you need to knowLasers are replacing conventional broadband light sources for fl uorescence imaging applications due to desirable laser properties like high brightness, stability, long lifetime, and narrow spectral bandwidth. These features enable higher sensitivity, better image fi delity, and superior axial resolution in a variety of imaging applications using laser-scanning and spinning-disk confocal microscopes and total-internal-refl ection fl uorescence (TIRF) microscopes. The narrow beam divergence, high spatial and temporal coherence, and well-defi ned polarization properties of lasers have enabled new fl uorescence imaging techniques, such as super-resolution.

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Leading Edge

In This Issue

An EnERgy Boost for CancerPAGE 711

Rapidly growing cancer cells increase their rate of aerobic glycolysis in a meta-bolic shift known as the Warburg effect. Their proliferation also demands highprotein folding capacity in the endoplasmic reticulum (ER). Fang et al. identifyan ER-localized enzyme, ENTPD5, that is responsible for both of these featuresof tumor cells. Inhibition of ENTPD5, which is commonly upregulated in humancancers, blocked tumor growth in mice. Thus, ENTPD5 inhibition could poten-tially become an anticancer therapy.

A Nudge and a Kick for Histone ReplacementPAGE 725

Most promoters in eukaryotes are marked with nucleosomes carrying a specialhistone H2A.Z, which is important for gene regulation. SWR1 incorporates

H2A.Z into nucleosomes in a histone replacement reaction. Luk et al. now report a mechanism that ensures that only nucle-osomes containing the canonical histone H2A are targeted for replacement. SWR1’s ATPase activity is sequentially stimu-lated by H2A-containing nucleosomes and free H2A.Z-H2B dimers, leading to eviction of nucleosomal H2A-H2B and depo-sition of H2A.Z-H2B. These stepwise events ensure the specificity of the nucleosome replacement reaction.

Locking Chromosome Cohesion during ReplicationPAGE 737

In eukaryotic cells, sister chromatids remain physically connected from the time of theirsynthesis during DNA replication until their separation during mitosis. Sister chromatidcohesion depends on the stable association of cohesin with DNA. Nishiyama et al.now show that Sororin binds cohesin during replication and stabilizes the cohesin-DNA complex by displacing the cohesin ‘‘unloading’’ protein Wapl. Distant orthologsof Sororin exist in many species, implying that this may be a widespread mechanismfor the maintenance of sister chromatid cohesion.

G Protein Lockdown for ChannelsPAGE 750

G protein-coupled potassium channels need to be turned off quickly, on a timescale faster than that afforded by either ligandclearance or receptor endocytosis. Raveh et al. now show that the GPCR kinase, GRK2, achieves rapid desensitization of theGIRK potassium channel by sequestering the G protein subunits required for GIRK activity. This kinase-independent functionof GRK2 thus allows rapid control of ligand-stimulated channel function.

Actin Cherry Picks Recycling ReceptorsPAGE 761

Signaling receptors recycle efficiently during endocytosis in a manner that differs from bulk membrane recycling. Puthen-veedu et al. use live cell imaging to show that distinct endosomal subdomains mediate active recycling of signaling receptors.The actin cytoskeleton binds in a sequence-dependent manner to the receptors, further concentrating and stabilizing thesedomains for recycling.

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Shapewear for the ERPAGE 774

The endoplasmic reticulum (ER) consists of the nuclear envelope and an exten-sive peripheral network of tubules and membrane sheets. Shibata et al. demon-strate that ER sheets are formed through stabilization of their highly curvededges by the reticulon/DP1/Yop1 p proteins. The membrane protein Climp63further shapes the sheets, acting as a spacer to regulate their area and luminalwidth.

HIV Pushes the T Cell Self-Destruct ButtonPAGE 789

The depletion of CD4 T cells during HIV infection is a hallmark of AIDS. Doitshet al. show that abortive infection of CD4 T cells elicits cell death. Incompletereverse transcripts of the virus accumulate in these cells and activate suicidalinnate antiviral and inflammatory responses. Thus, T cell death is not triggered

by new virus production but, rather, by a suicide mechanism, which likely evolved to protect the host but in fact contributes toimmunodeficiency.

Hungry but Still HearingPAGE 802

Caloric restriction (CR) extends the life span of many species and slows the progression of age-related hearing loss (AHL).Here, Someya et al. report that mitochondrial Sirt3 mediates the prevention of AHL and reduces oxidative damage incalorie-restricted mice. In response to CR, Sirt3 deacetylates and activates isocitrate dehydrogenase 2, leading to anenhanced glutathione antioxidant defense system in mitochondria. These results suggest that Sirt3-dependent mitochondrialadaptations may be a central mechanism to delay aging in mammals.

Outfoxing AgingPAGE 813

Loss of muscle strength is one of the most obvious changes that we experienceas we age, but how this connects with systemic aging is unclear. Demontis andPerrimon report that accumulation of protein aggregates in aging Drosophilamuscle is reduced by FOXO/4E-BP signaling, delaying muscle senescence.This pathway in muscle prevents overall aging and protein aggregation in othertissues. These results provide a framework to understand the coordination oftissue and organismal aging.

Golgi Decides, Axon or DendritePAGE 826

Neuronal cells polarize to develop an axon at one pole and dendrites at theother. Matsuki et al. identify two signaling pathways that influence Golgimorphogenesis to regulate this polarization. The Stk25 kinase acts throughthe Golgi protein GM130 to promote a condensed Golgi morphology and axon development. The Reelin-Dab1 signalingpathway, previously known to regulate other aspects of nervous system development, antagonizes the Stk25 pathway topromote Golgi extension and dendrite development. Thus, Golgi distribution is a central factor in neuronal development.

Structural Fingerprints of the Human GenomePAGE 837

Genomic structural variation—insertions, duplications, and deletions—are important contributors to human disease andgenetic diversity. The precise molecular characteristics of these variants have been difficult to ascertain by standard high-throughput genome sequencing. Kidd et al. now report a resource of fosmid clones obtained from the genomes of 17 indi-viduals. The authors characterize the breakpoints of more than a thousand structural variants, allowing inference of the molec-ular pathways that generated them and offering an in-depth view of the characteristics of human genomic variation.


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Leading Edge

Select: Cell Cycle

The phases of the cell cycle must be exquisitely timed and tightly regulated in order to ensure properchromosome replication and segregation and cell division. New findings described in this issue’s Selectaddress key regulatory events in the cell cycle and reveal potential clinical outcomes of errors in theseprocesses.

An Epigenetic License to ReplicateChromosome replication needs to occur once and only once during the cell cycleto produce daughter cells with accurate genetic content. Licensing of replicationorigins is one form of DNA synthesis regulation, in which origins are loaded withpre-replication complex (RC) proteins during the end of M phase and throughoutG1. Without this licensing event, replication origins cannot be activated. Newfindings from Tardat et al. identify the methyltransferase PR-Set7—and thehistone modification that it catalyzes, methylation of histone H4 lysine 20(H4K20me1)—as a key regulator of the onset of licensing in mammalian cells.The authors show that PR-Set7 and H4K20me1 levels are cell cycle regu-lated—both are high during M and G1 phases, dropping in S when synthesisbegins—and that proteasomal degradation of PR-Set7 is needed to preventDNA re-replication. The authors also show that silencing PR-Set7 leads todecreased chromatin loading of pre-RC proteins and reduced origin firing duringS phase, whereas targeting PR-Set7 to nonorigin sites on the chromatin is suffi-cient to induce H4K20me1 and the assembly of pre-RC proteins. Future studies

are needed to investigate how H4K20me1 facilitates chromatin loading of pre-RC proteins.M. Tardat et al. (2010). Nat. Cell Biol. Published online October 17, 2010. 10.1038/ncb2113.

Getting a Toehold on MicrotubulesThe ability of the kinetochore to maintain an attachment between chromo-somes and microtubules is necessary for proper chromosomal segregationduring anaphase. The Ndc80 complex is known to be a key regulatory sitefor microtubule attachment, but, given the highly dynamic process of micro-tubule assembly and disassembly occurring during segregation, it has beena challenge to identify how the Ndc80 complex physically holds on to sucha rapidly changing structure. Alushin et al. address this using cryo-electronmicroscopy to better reveal the metazoan Ndc80 complex bound to micro-tubules. The authors find that the Ndc80 complex binds both a- andb-tubulin monomers and identify a ‘‘toe’’—a short section of the NDC80protein that recognizes a site between two tubulin monomers, a hinge pointfor tubulin bending. The toe appears to prefer binding straight tubulin, sug-gesting that it could act as a sensor for tubulin conformation. At the sametime, the N terminus of NDC80 allows high-affinity microtubule binding andappears to mediate self-assembly of Ndc80 complexes in a manner that ismodulated via phosphorylation by Aurora B kinase. The authors proposea model in which phosphorylated Ndc80 complexes bind a microtubuleand spindle forces then pull the bound complex out of the Aurora B kinasephosphorylation zone. The resulting dephosphorylation of NDC80 results inhigh-affinity clusters forming in linear arrays along the microtubule. Thiscluster arrangement is consistent with a biased diffusion model of kineto-chore attachment and movement. On a shrinking microtubule, the Ndc80-microtubule interaction would be reduced due to conformational changesin tubulin at the disassembling or depolymerizing end, and the cluster would diffuse along the microtubules towardthe pole, thereby moving the chromosome in that direction.G.M. Alushin et al. (2010). Nature 467, 805–810.

Re-replicating G2 cells (cyclin B1, red;

EdU, green). Image courtesy of E. Julien.

Two Ndc80 molecules (blue and yellow;

N terminus, magenta) binding tubulin

(green; C terminus, red). Image courtesy of

E. Nogales.


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Mounting Tension in Lead-Up to Fateful DecisionAsymmetric cell division, which generates daughter cells with different developmental fates, is often achieved throughasymmetric positioning of the mitotic spindle. However, some dividing cells start out with a centered spindle thatbecomes displaced during anaphase. This progressive asymmetry has been postulated to arise from greater elonga-tion of microtubules on one side of the spindle. New findings from Ou et al. suggest that nonmuscle myosin II might alsoplay a role. The authors show that in the QR.a neuroblast of Caenorhabditis elegans, myosin II becomes asymmetricallydistributed during anaphase, concentrating at the anterior side of the cleavage furrow. Consequently, the anteriormembrane becomes less dynamic and shrinks inward, whereas the posterior membrane expands like a balloon, sug-gesting that cortical tension and contractile forces driven by myosin II could be a factor in developing asymmetry. Theauthors also used CALI (chromophore-assisted laser inactivation) to specifically inactivate myosin II at the anteriormembrane and find that this increases the size of the anterior daughter cell and can alter its fate from apoptosis todifferentiation into a neuron-like cell. Future work is needed to better understand the respective contributions of micro-tubule elongation, myosin polarization, and perhaps other unknown mechanisms to the regulation of asymmetric divi-sion and cell fate.G. Ou et al. (2010). Science. Published online September 30, 2010. 10.1126/science.1196112.

Spindle Position, a Neuronal Mover and MakerHuman microcephaly is a neurodevelopmental disorder characterized by a small brain,fewer surface ridges, and reduced cortical neuron numbers. Two recent papers usedlinkage analysis and genome capture in affected families to identify WDR62 asa common cause of genetic microcephaly and characterized the WDR62 protein asa spindle pole protein expressed in mitotic neural precursors. After sequencingaffected individuals to identify specific disease-causing mutations, Nicholas et al. ex-pressed mutant WDR62 in HeLa cells and showed that the normal accumulation of theprotein at the spindle poles during mitosis is disrupted. Given the phenotype ofreduced neuron numbers and small brain seen in microcephaly, one possibility theauthors suggest is that WDR62 could be involved in proper positioning of the mitoticspindle and cleavage furrow, such that mutant WDR62 results in insufficient symmetricdivisions—needed to produce neural precursors—early in cortical development. Inagreement, Yu et al. show that the brain of an affected individual has profound corticaldefects, with thin sparse cortical layers and aberrant repositioning of neurons tosubcortical regions, suggesting deficits in neurogenesis and migration. Further

description of the specific role of WDR62 at the spindle will clarify how it is involved in cerebral development andaid in our understanding of the etiology of microcephaly.A.K. Nicholas et al. (2010). Nat. Genet. Published online October 3, 2010. 10.1038/ng.682.T.W. Yu et al. (2010). Nat. Genet. Published online October 3, 2010. 10.1038/ng.683.

Rebecca Alvania

Photograph of human microce-

phalic brain. Image courtesy of

C. Walsh.


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Previews

ER Sheets Get Roughed UpCharles Barlowe1,*1Department of Biochemistry, Dartmouth Medical School, Hanover, NH 03755, USA

*Correspondence: [email protected] 10.1016/j.cell.2010.11.011

The molecular machinery that shapes the endoplasmic reticulum’s (ER’s) membrane into orderednetworks of ‘‘smooth’’ tubules and ‘‘rough’’ sheets is poorly defined. Shibata et al. (2010) now reportthat sheet-inducing proteins, such as Climp-63, are enriched in the ‘‘rough’’ ER by their associationwith membrane-bound ribosomes, whereas curvature-inducing proteins localize at highly bentedges of membrane sheets.

The elaborate morphologies of the endo-

plasmic reticulum have fascinated cell

biologists for years. Compartments of the

endoplasmic reticulum (ER) membrane

form the nuclear envelope and then

extend throughout the cell periphery in

an interconnected network of mem-

brane tubules and flattened discs called

cisternae. How do these ordered arrays

of membranes form, and how are their

structures connected to their cellular

function? In this issue of Cell, Shibata

and coworkers define a class of sheet-

inducing membrane proteins that are en-

riched in the ribosome-studded ‘‘rough’’

ER. These proteins cooperate with

membrane curvature-stabilizing factors

to govern the relative level of sheets

and tubules of the ER, providing a molec-

ular basis for the longstanding morpho-

logical descriptions of ‘‘rough’’ and

‘‘smooth’’ ER.

ER morphologies vary greatly across

different species and cell types. For

example, highly active secretory cells,

such as pancreatic exocrine cells and

plasma B cells, are packed full of flattened

cisternae of rough ER. Live cell imaging

also reveals that ER membranes are

highly dynamic networks, undergoing

constant remodeling often in response to

physiological conditions.

Previous studies focusing on the

smooth ER found that tubule formation

depends on a class of integral membrane

proteins belonging to the reticulon and

DP1 families (Voeltz et al., 2006). Reticu-

lon and DP1 proteins are highly enriched

in tubular ER elements, and they contain

transmembrane segments with a double

hairpin structure that induces positive

membrane curvature by inserting like

a wedge into ER membranes (Figure 1).

Indeed, reconstitution of purified reticulon

and DP1 proteins into synthetic lipo-

somes (i.e., artificial vesicles with a lipid

bilayer) was sufficient to generate mem-

brane tubules with a high degree of curva-

ture (Hu et al., 2008). Thus, intrinsic

properties of the reticulon and DP1 pro-

teins are sufficient to induce membrane

tubulation.

However, ER tubules also form

branched, reticular morphologies. Gener-

ation of these net-like structures requires

additional factors, specifically atlastin

GTPases, which drive fusion of ER

tubules into branched networks (Hu

et al., 2009; Orso et al., 2009). Of interest,

atlastin isoforms were detected in associ-

ation with the reticulon proteins, sug-

gesting that the formation of tubules and

branching are coordinated processes

(Hu et al., 2009).

In contrast to our understanding of

ER tubules, the molecular mechanisms

underlying the formation of ER sheets

have been elusive. Now, Shibata et al.

(2010) uncover an unexpected connec-

tion between the sheet-inducing factor

Climp-63 and the reticulon and DP1

proteins. Their discovery begins with

a key observation regarding the translo-

con complex, a large multisubunit chan-

nel that transports, or ‘‘translocates,’’

nascent polypeptides across ER mem-

brane into the interior of the ER.

Shibata and colleagues observe that

components of the translocon complex

are not only highly enriched in ER sheets,

but they also form a specialized subdo-

main within ER membranes. Moreover,

when the authors treat cells with the anti-

biotic puromycin, which disassembles

groups of ribosomes bound to the ER

membranes (i.e., polysomes), proteins

of the translocon complex redistribute

between ER sheets and tubules. This

finding suggests that actively translating

polysomes concentrate translocon com-

plexes into sheet subdomains of the ER.

To identify the structural components

of these ER sheet domains, Shibata and

colleagues then perform a proteomic

analysis of rough ER membranes from

pancreatic secretory cells. Indeed, the

most abundant protein constituents in

ER sheets are components of the translo-

con complex and Climp-63. Moreover,

microarray experiments reveal that

Climp-63 messenger RNA (mRNA) levels

are among the most highly induced

messages during proliferation of ER sheet

structures during the differentiation of

immature B cells into IgG secreting

plasma cells. Climp-63 is an ER trans-

membrane protein that contains an

extended coiled-coil domain in the interior

of the ER (i.e., the ER lumen). Previous

studies suggested that this coiled-coil

domain contributes to ER morphology

by forming a scaffold in the ER lumen

(Klopfenstein et al., 2001).

To test the functional role of Climp-63

in ER sheet formation, Shibata and

colleagues then overexpress Climp-63 in

cultured cells, which causes a dramatic

proliferation of ER sheets. Moreover, the

distance between the sheets is �50 nm,

the standard distance between ER sheets

in mammalian cells (Figure 1). In contrast,

decreasing the expression of Climp-63

does not deplete cells of ER sheets, but

instead, it causes a marked reduction in

the distance between cisternal sheets.

Further, these sheets are spread diffusely


throughout the cytoplasm,

a similar phenotype as the

authors observe when they

treat cells with puromycin.

Finally, Climp-63 and the

reticulon protein Rtn4 have

opposing effects on ER mor-

phology. Increased expres-

sion of Rtn4 reduces the

number of ER sheets,

whereas co-overexpression

with Climp-63 restores sheet

structures in these cells.

Importantly, reticulon pro-

teins strikingly localize to the

highly curved edges of ER

sheets, and this occurs

when reticulon genes are

expressed at endogenous

levels or when both Climp-63

and reticulon genes were

overexpressed together.

The authors then propose

the most basic mechanism

for sheet formation that is

also consistent with their find-

ings. In this model, reticulons

and DP1 proteins partition

into the edges of sheets, where they

induce a high degree of curvature at the

edges of closely apposed membrane bila-

yers (Figure 1). However, assembling the

ordered array of rough ER membranes in

active secretory cells also depends on

the coiled-coil domain of Climp-63, which

serves as a spacer between the sheets

in the ER lumen (Figure 1). Lastly, the

authors propose that Climp-63, together

with translocon complexes, partition into

sheet domains with membrane-bound

polysomes to generate the rough ER.

This model proposed by Shibata and

colleagues is also supported by previous

studies showing that the coiled-coil

domain of Climp-63 assembles into

a-helical rods that are required to restrict

the lateral mobility of Climp-63 (Klopfen-

stein et al., 2001; Nikonov et al., 2007)

(Figure 1). Moreover, Climp-63 is known

to bind microtubules (Klopfenstein et al.,

1998), suggesting an additional level of

ER organization that is connected to the

cell’s overall structure.

Although reticulon and DP1 proteins

partition into sheet edges in vivo and ex-

pressing Climp-63 drives ER sheet prolif-

eration, it is still unknown whether these

factors are sufficient for sheet formation

or whether other factors contribute to

this process. A minimally reconstituted

liposome system successfully demon-

strated that reticulon and DP1 proteins

drive tubule formation in vitro (Hu et al.,

2008). This system should provide a

powerful tool for determining whether

adding purified Climp-63 is sufficient

for sheet formation. Furthermore, varying

the ratio of curvature- and sheet- inducing

proteins in liposomes of defined lipid

compositions could provide insights into

the role that specific lipids play in gener-

ating observed ER morphology.

Finally, sheets and tubules

are not the only morphologies

of ER membranes. For

example, specialized struc-

tural domains of the ER are

involved in metabolism of

hydrophobic compounds,

formation of ER-mitochon-

drial junctions, transport of

Ca2+, formation of lipid drop-

lets, and protein export from

ER subdomains called transi-

tional ER sites. The molecular

machinery that generates

these ER structures awaits

elucidation. Although the

components that sculpt ER

sheets and tubules might

also contribute to the mor-

phology of these other struc-

tures, it seems likely that

novel mechanisms will also

be discovered.

REFERENCES

Hu, J., Shibata, Y., Voss, C., She-

mesh, T., Li, Z., Coughlin, M.,

Kozlov, M.M., Rapoport, T.A., and Prinz, W.A.

(2008). Science 319, 1247–1250.

Hu, J., Shibata, Y., Zhu, P.-P., Voss, C., Rismanchi,

N., Prinz, W.A., Rapoport, T.A., and Blackstone, C.

(2009). Cell 138, 549–561.

Klopfenstein, D.R., Kappeler, F., and Hauri, H.P.

(1998). EMBO J. 17, 6168–6177.

Klopfenstein, D.R., Klumperman, J., Lustig, A.,

Kammerer, R.A., Oorschot, V., and Hauri, H.P.

(2001). J. Cell Biol. 153, 1287–1300.

Nikonov, A.V., Hauri, H.P., Lauring, B., and Krei-

bich, G. (2007). J. Cell Sci. 120, 2248–2258.

Orso, G., Pendin, D., Liu, S., Tosetto, J., Moss,

T.J., Faust, J.E., Micaroni, M., Egorova, A., Marti-

nuzzi, A., McNew, J.A., and Daga, A. (2009). Nature

460, 978–983.

Shibata, Y., Shemesh, T., Prinz, W.A., Palazzo,

A.F., Kozlov, M.M., and Rapoport, T.A. (2010).

Cell 143, this issue, 774–788.

Voeltz, G.K., Prinz, W.A., Shibata, Y., Rist, J.M.,

and Rapoport, T.A. (2006). Cell 124,

573–586.

Figure 1. Molecular Model for the Generation of ER Membrane

Sheets and TubulesCross-section of an endoplasmic reticulum (ER) cisterna showing the curva-ture-inducing proteins reticulons and DP1 (purple) enriched in highly bentmembrane tubules and edges of the sheet. In contrast, the sheet-inducingprotein Climp-63 (blue) is excluded from tubules and, instead, partitions intosheet domains with translocon complexes. Climp-63 could assemble intoparallel coiled-coil arrangements to flatten membranes and to serve as luminalER spacers that keep individual sheets a specific distance apart (�50 nm inmammalian cells).

666 Cell 143, November 24, 2010 ª2010 Elsevier Inc.

Leading Edge

Previews

SIRT3 in Calorie Restriction:Can You Hear Me Now?Carlos Sebastian1 and Raul Mostoslavsky1,*1The Massachusetts General Hospital Cancer Center, Harvard Medical School, Boston, MA 02114, USA*Correspondence: [email protected]

DOI 10.1016/j.cell.2010.11.009

Caloric restriction decreases oxidative damage and extends life span in many organisms. Someyaet al. (2010) show that the sirtuin SIRT3mediates the protective effects of caloric restriction on age-related hearing loss by promoting the mitochondrial antioxidant system through the regulation ofisocitrate dehydrogenase 2 (Idh2).

Despite two decades of effort, caloric

restriction remains the only treatment

demonstrated to extend life span and to

delay the progression of several diseases

normally associated with aging, such as

cancer, diabetes, and neurological disor-

ders. Early experiments in yeast showed

that the life span extension mediated by

caloric restriction involves Sir2, the found-

ing member of the sirtuin family of histone

deacetylases. However, later experi-

ments have questioned this association

(Longo and Kennedy, 2006), and the role

of mammalian sirtuins in life span exten-

sion by caloric restriction is still under

study. In this context, although SIRT1

appears to be the major mammalian sir-

tuin involved in the metabolic effects of

caloric restriction (Haigis and Guarente,

2006), the precise role of sirtuins in the

longevity response remains unclear. In

this issue of Cell, Someya et al. (2010)

bring some light to the field by describing

a new function for the mitochondrial

SIRT3 protein in the prevention of hearing

loss mediated by caloric restriction during

aging. These tantalizing results suggest

that SIRT3 might play an important role

in slowing the aging process in mammals.

Age-related hearing loss is a hallmark of

mammalian aging and the most common

sensory disorder in the elderly (Liu and

Yan, 2007). It is characterized by a gradual

loss of spiral ganglion neurons and

sensory hair cells in the cochlea of the

inner ear (Liu and Yan, 2007). Given that

the affected cells are postmitotic and do

not regenerate, their loss leads to an

age-associated decline in hearing func-

tion. Several groups have studied hearing

loss as an example of age-related degen-

eration in mouse models. Remarkably,

early work demonstrated that caloric

restriction slows age-related hearing loss

in animal models (Sweet et al., 1988).

Moreover, in their previous work, Prolla

and colleagues demonstrated that caloric

restriction induces expression of the

SIRT3 gene in the cochlea (Someya et al.,

2007). They now elegantly follow up on

these results, proving a role for this sirtuin

in the delay in hearing loss due to caloric

restriction and elucidating the molecular

mechanisms underlying this effect.

Someya et al. use wild-type and SIRT3-

deficient mice fed a diet in which caloric

intake is reduced to 75% and compare

them to control mice fed with a regular

diet. The authors first look at the hearing

response of the animals and find that, as

expected, aging leads to hearing loss in

both wild-type and SIRT3-deficient mice.

However, whereas caloric restriction

delays the progression of hearing loss in

wild-type mice, this effect is completely

abolished in SIRT3-deficient animals.

These results are consistent with the

effects of caloric restriction on spiral

ganglion neurons and hair cells in these

mice. In wild-type animals, a calorie

restricted diet reduces the age-related

loss of neurons and hair cells, whereas

this effect is abrogated in SIRT3-deficient

mice. Together, these results clearly pin-

point SIRT3 as a critical molecular determi-

nant regulating the response to caloric

restriction in age-related hearing loss.

The authors next study the metabolic

effects induced by caloric restriction in

SIRT3-deficient mice. With a normal diet,

SIRT3-deficient animals appear pheno-

typically normal, in accordance with

previous studies (Schwer et al., 2009).

However, whereas wild-type mice display

lower levels of serum insulin and triglycer-

ides when fed a calorie-restricted diet,

SIRT3-deficient mice do not show this

response. Based on these results, the

authors argue that SIRT3 plays a role in

metabolic adaptations to caloric restric-

tion. It remains unclear, however, whether

SIRT3 can also mediate the effects of

calorie restriction in other tissues or

whether it does so specifically in the

context of hearing loss.

The authors then investigate the molec-

ular mechanisms involved in this process.

Given that caloric restriction reduces age-

associated oxidative damage to macro-

molecules (Sohal and Weindruch, 1996),

Someya et al. analyze levels of oxidative

damage to DNA in several tissues. They

find that a calorie-restricted diet reduces

this type of damage in wild-type mice,

but not in SIRT3-deficient animals. Impor-

tantly, this is the first evidence that

a mammalian sirtuin regulates levels of

oxidative stress in response to caloric

restriction.

But how does SIRT3 regulate oxidative

damage during caloric restriction? Given

that SIRT3 localizes to the mitochondria,

the authors hypothesize that SIRT3 could

regulate the antioxidant systems present

in this organelle. Using a combination of

cellular and biochemical experiments,

they discover that SIRT3 regulates the

mitochondrial glutathione antioxidant

defense system. Glutathione is the main

small molecule antioxidant in cells and is

generated by glutathione reductase in

a reaction dependent on NADPH. The

authors show that SIRT3 modulates the


conversion of oxidized gluta-

thione to reduced glutathione

in response to caloric restric-

tion. They find that, under

these conditions, SIRT3

binds and deacetylates the

mitochondrial isocitrate de-

hydrogenase 2 enzyme

(Idh2), the enzyme that gener-

ates NADPH, increasing the

enzyme’s activity. In agree-

ment with these results, Idh2

deacetylation and activity, as

well as NADPH levels, in-

crease during caloric restric-

tion in all wild-type tissues

tested, whereas SIRT3 defi-

ciency impairs this response.

Finally, overexpressing SIRT3

and Idh2 promotes cell

viability upon oxidative dam-

age. Together, these data

lead the authors to propose

a model in which caloric

restriction promotes SIRT3

expression, leading to the de-

acetylation and activation of

Idh2, thus providing resis-

tance to oxidative stress and

inhibiting the age-related

loss of spiral ganglion neu-

rons and hair cells (Figure 1).

Although Someya et al.

provide enough data to

prove that the effects of

caloric restriction on age-

related hearing loss are

dependent on SIRT3, key

questions remain. First, does SIRT3

mediate the effects of caloric restriction

in other tissues? And if so, what are its

substrates? Multiple mitochondrial

proteins are deacetylated upon caloric

restriction in a SIRT3-dependent manner

(Schwer et al., 2009). It is therefore

important to determine whether Idh2 is

the main SIRT3 target in preventing

oxidative stress or whether other SIRT3

substrates contribute as well. Second,

what is the relationship between the

effect of SIRT3 on Idh2 and the recently

described role for SIRT3 in fatty acid

oxidation during nutrient stress (Hirschey

et al., 2010)? Are these functions coordi-

nated? If they are not, how is specificity

achieved? Third, can we mimic the

effects of caloric restriction using SIRT3

activators? If so, such reagents would

have significant therapeutic potential.

Finally, because other sirtuins also have

prominent roles in metabolic regulation

(Finkel et al., 2009), can we extend

some of these findings to other sirtuins?

SIRT1, for example, has been linked to

the response of mammals to caloric

restriction (Haigis and Guarente, 2006),

and it is therefore possible that the

activity of this and other sirtuins may be

regulated in a coordinated fashion

following nutrient starvation.

Regardless of the utopian dream of life

span extension, answering some of these

questions may provide a step forward for

treating age-related pathologies, bringing

us closer to a healthier life

span. In the words of Francois

Jacob, ‘‘In a world of unlimited

imagination, we are continu-

ally inventing a possible world

or a piece of a world, and then

comparing it with the real

world.’’ In the context of sir-

tuins, it seems we are starting

to put some of these pieces

together.

ACKNOWLEDGMENTS

We would like to thank all of the

members of the Mostoslavsky lab

for helpful comments. C.S. is the

recipient of a Beatriu de Pinos Post-

doctoral Fellowship (Generalitat de

Catalunya). R.M. is a Sidney Kimmel

Scholar, a Massachusetts Life

Science Center New Investigator

Scholar, and the recipient of an

AFAR Award. Work in the Mosto-

slavsky lab is funded, in part, by

National Institutes of Health.

REFERENCES

Finkel, T., Deng, C.X., and Mosto-

slavsky, R. (2009). Nature 460,

587–591.

Haigis, M.C., and Guarente, L.P.

(2006). Genes Dev. 20, 2913–2921.

Hirschey, M.D., Shimazu, T., Goetz-

man, E., Jing, E., Schwer, B.,

Lombard, D.B., Grueter, C.A., Har-

ris, C., Biddinger, S., Ilkayeva, O.R.,

et al. (2010). Nature 464, 121–125.

Liu, X.Z., and Yan, D. (2007). J. Pathol. 211,

188–197.

Longo, V.D., and Kennedy, B.K. (2006). Cell 126,

257–268.

Schwer, B., Eckersdorff, M., Li, Y., Silva, J.C., Fer-

min, D., Kurtev, M.V., Giallourakis, C., Comb, M.J.,

Alt, F.W., and Lombard, D.B. (2009). Aging Cell 8,

604–606.

Sohal, R.S., and Weindruch, R. (1996). Science

273, 59–63.

Someya, S., Yamasoba, T., Weindruch, R., Prolla,

T.A., and Tanokura, M. (2007). Neurobiol. Aging

28, 1613–1622.

Someya, S., Yu, W., Hallows, W.C., Xu, J., Vann,

J.M., Leeuwenburg, C., Tanokura, M., Denu,

J.M., and Prolla, T.A. (2010). Cell 143, this issue,

802–812.

Sweet, R.J., Price, J.M., and Henry, K.R. (1988).

Audiology 27, 305–312.

Figure 1. Caloric Restriction, SIRT3, and Age-Related Hearing LossDuring aging (left), oxidative damage (ROS, reactive oxygen species) leads tothe loss of spiral ganglion neurons and sensory hair cells in the ear, leading toage-related hearing loss. Caloric restriction (right) prevents the age-associ-ated loss of spiral ganglion neurons and sensory hair cells. Someya et al.(2010) show that caloric restriction leads to an increase in SIRT3 levels inthe mitochondria. By promoting the deacetylation of isocitrate dehydrogenase2 (Idh2), SIRT3 promotes the accumulation of NADPH, hence activating gluta-thione reductase (GSR), which generates reduced glutathione (GSH), a cellularantioxidant.


Leading Edge

Previews

ATP Consumption PromotesCancer MetabolismWilliam J. Israelsen1 and Matthew G. Vander Heiden1,*1Koch Institute for Integrative Cancer Research, Massachusetts Institute of Technology, Cambridge, MA 02139, USA*Correspondence: [email protected]

DOI 10.1016/j.cell.2010.11.010

Cancer cells metabolize glucose by aerobic glycolysis, a phenomenon known as the Warburgeffect. Fang et al. (2010) show that the endoplasmic reticulum enzyme ENTPD5 promotes ATPconsumption and favors aerobic glycolysis. The findings suggest that nutrient uptake in cancercells is limited by ATP and satisfies energy requirements other than ATP production.

Mounting evidence suggests that cancer

cells engage in a unique metabolic pro-

gram that allows for rapid cell prolifera-

tion. Nonproliferating cells can use glycol-

ysis products to generate ATP for their

energy needs. Such cells generally have

low rates of glycolysis followed by

oxidation of pyruvate in the mitochondria,

leading to efficient generation of ATP.

Dividing cells, in contrast, also use glycol-

ysis intermediates for the synthesis of

macromolecules and must therefore

balance their ATP requirements and

biosynthetic needs (Vander Heiden et al.,

2009). Metabolism of glucose by aerobic

glycolysis, a phenomenon known as the

Warburg effect, may help dividing cells

strike this balance.

The phosphoinositide 3-kinase (PI3K)

signaling pathway, which is activated in

many cancers, regulates cell growth and

survival. PI3K signaling has been impli-

cated in the altered glucose metabolism

of cancer cells, and the serine/threonine

kinase AKT, a major PI3K effector,

promotes glucose uptake and increases

the activity of glycolytic enzymes (DeBer-

ardinis et al., 2008). In this issue of Cell,

Fang et al. (2010) report an important

mechanism by which AKT signaling leads

to increased aerobic glycolysis. They

showthat AKT activation promotesprotein

glycosylation in the endoplasmic retic-

ulum, which elevates ATP consumption

and derepresses a rate-limiting enzyme

in glycolysis that is otherwise inhibited by

an elevated ratio of ATP to AMP. This

work suggests how proliferating cells

may integrate growth signals with energy

status to enable increased glucose uptake

to support cell proliferation.

Activation of the PI3K pathway in

cancer can occur via genetic alterations

that allow growth factor-independent

kinase activation or via the loss of PTEN,

a lipid phosphatase that attenuates PI3K

signaling. Fang et al. now find that cell

extracts from PTEN-deficient cells have

an enhanced ability to generate AMP

from ATP. Subsequent purification and

biochemical characterization of this

activity led to the identification of ectonu-

cleoside triphosphate diphosphohydro-

lase 5 (ENTPD5) as the enzyme associ-

ated with the ATP hydrolysis activity.

PI3K signaling leads to upregulation of

ENTPD5, a UDPase that promotes the

N-glycosylation and folding of glycopro-

teins in the ER by hydrolyzing UDP to

UMP (Trombetta and Helenius, 1999)

(Figure 1). UDP hydrolysis in the ER is

a reaction necessary to promote protein

folding via the calnexin/calreticulin

pathway. It is linked to ATP hydrolysis in

the cytosol by a cycle of glucose and

phosphate transfer reactions. As part

of this cycle, the UDP-glucose/UMP anti-

porter exports UMP out of the ER in

exchange for importing UDP-glucose

into the ER (Hirschberg et al., 1998). The

UGGT enzyme then uses UDP-glucose

to transfer glucose to proteins in the ER

(Vembar and Brodsky, 2008). This

glucose addition to nascent glycoproteins

is necessary for their calnexin/calreticulin-

mediated protein folding. Thus, disruption

of ENTPD5 in PTEN-deficient cells results

in decreased protein N-glycosylation and

causes ER stress.

Cell surface proteins, including many

growth factor receptors, are N-glycosy-

lated. Fang et al. show that disruption of

ENTPD5 leads to decreased levels of

several growth factor receptors, including

epidermal growth factor receptor (EGFR),

insulin-like growth factor receptor

b (IGFR-b), and Her2/ErbB2. Given that

growth factor signaling plays an important

role in increasing nutrient metabolism in

rapidly proliferating cells (DeBerardinis

et al., 2008), these new findings suggest

that cellular ATP levels can influence the

folding and expression of growth factor

receptors, perhaps ensuring that cells do

not attempt to grow when ATP is limiting.

Furthermore, because glucose metabo-

lism by the hexosamine biosynthesis

pathway provides the carbon for these

receptor glycosylation events, the avail-

ability of glucose may provide a means

to couple nutrient levels with growth

factor receptor expression. These feed-

backs may exist to prevent a metabolic

catastrophe caused by activation of the

cell growth machinery when the supply

of nutrients or ATP is limiting.

How does ENTPD5 regulate ATP

levels? Fang et al. find that reconstitution

of the ATP consuming activity also

requires the presence of UMP/CMP

kinase-1 and adenylate kinase-1. UMP/

CMP kinase-1 catalyzes the rephosphor-

ylation of the UMP generated by ENTPD5

into UDP (Figure 1), in the process con-

verting ATP to ADP. Adenylate kinase-1

then converts ADP molecules into ATP

and AMP, thus allowing the cycle to

continue. Surprisingly, this cycle involving

ENTPD5 is a major source of ATP

consumption in PTEN-deficient cells.

Furthermore, these reactions directly

affect the cell’s glycolytic rate. Whereas

increased ENTPD5 expression has no


effect on cellular respiration, it increases

lactate production, suggesting a link

between ATP consumption and in-

creased glycolytic flux. In a series of

experiments to determine how ENTPD5

increases glucose entry into glycolysis,

Fang et al. find that the ratio of fructose-

6-phosphate to fructose-1-6-bisphos-

phate increases in cells following ENTPD5

knockdown, consistent with inhibition of

this step in glycolysis. Phosphofructoki-

nase (PFK), the enzyme that catalyzes

this reaction, is the major enzyme control-

ling glucose commitment to the glycolytic

pathway (Dunaway, 1983). A high ATP/

AMP ratio in the cell inhibits both PFK

activity and glucose metabolism by

glycolysis. In fact, the authors conclude

that increased ATP consumption by

ENTPD5 increases glycolysis by lowering

the ATP/AMP ratio and relieving allosteric

inhibition of PFK.

ATP is likely not the growth-limiting

resource for most cells (Vander Heiden

et al., 2009). The concept that glucose

utilization by tumor cells may be limited

by ATP consumption to prevent feedback

inhibition of PFK has been suggested

previously (Scholnick et al., 1973). This

study finally provides a mechanism by

which cells can increase ATP consump-

tion to drive glucose uptake. An additional

mechanism has also recently been

described in which glucose incorporation

into biosynthetic pathways occurs

without producing excess ATP (Vander

Heiden et al., 2010). Together, these

studies support the notion that altered

metabolism in cancer is not adapted to

support ATP production.

Fang et al. show that ENTPD5 expres-

sion correlates with PI3K activation in

human prostate cancer cell lines and

tumor tissue samples. Not all cancer cells

are dependent on activated PI3K, sug-

gesting that increased ENTPD5 activity

may not be a universal mechanism for

lowering ATP levels in tumors. However,

other enzymes involved in regulating

nucleotide pools in cells have also been

linked to cancer (Hartsough and Steeg,

2000), and there are additional homologs

of ENTPD5 whose functions are not well

understood. These enzymes may be

involved in analogous cycles of ATP con-

sumption, leading to enhanced glucose

metabolism in other genetic contexts.

Fang et al. also show that decreased

ENTPD5 expression inhibits tumor

growth, possibly via pleiotropic effects

involving induction of ER stress and

altered glucose metabolism. Consider-

ation of ENTPD5 as a potential thera-

peutic target in PI3K-driven cancer is

interesting given that pharmacological

inhibition of ENTPD5 is predicted to

decrease tumor ATP consumption.

Although counterintuitive, the resulting

increase in ATP/AMP ratio might reduce

the entry of glucose into glycolysis and

starve the cells of precursors necessary

for biosynthesis. Successful efforts to

target cancer metabolism will likely arise

from understanding the feedbacks and

complex regulation that are required for

anabolic metabolism. The study by Fang

et al. provides new insight by demon-

strating that ATP consumption serves to

increase glucose flux to satisfy the ener-

getic and biosynthetic demands of

a rapidly proliferating cell.

ACKNOWLEDGMENTS

We thank Brooke Bevis for her help preparing the

figure and editing the manuscript. M.G.V.H. is

a consultant to Agios Pharmaceuticals regarding

development of compounds targeting cancer

metabolism and is a member of its Scientific Advi-

sory Board.

REFERENCES

DeBerardinis, R.J., Lum, J.J., Hatzivassiliou, G.,

and Thompson, C.B. (2008). Cell Metab. 7, 11–20.

Dunaway, G.A. (1983). Mol. Cell. Biochem. 52,

75–91.

Fang, M., Shen, Z., Huang, S., Zhao, L., Chen, S.,

Mak, T.M., and Wang, X. (2010). Cell 143, this

issue, 711–724.

Hartsough, M.T., and Steeg, P.S. (2000). J. Bioen-

erg. Biomembr. 32, 301–308.

Hirschberg, C.B., Robbins, P.W., and Abeijon, C.

(1998). Annu. Rev. Biochem. 67, 49–69.

Figure 1. ENTPD5 Promotes Glycolysis in Proliferating CellsFang et al. (2010) show that the endoplasmic reticulum (ER) UDPase ectonucleoside triphosphatediphosphohydrolase 5 (ENTPD5) is expressed in response to phosphoinositide 3-kinase (PI3K) signaling.Activation of PI3K results in FOXO phosphorylation by AKT and loss of ENTPD5 transcriptional repres-sion. This leads to increased ENTPD5 enzyme activity in the ER, promoting protein folding. ENTPD5activity promotes the import of UDP-glucose into the ER, where UGGT transfers glucose to an unfoldedN-glycoprotein and produces UDP. Properly folded N-glycoproteins, such as growth factor receptors,exit the cycle, whereas unfolded proteins undergo further folding attempts or are degraded. ENTPD5activity enables this process by hydrolyzing UDP to provide the UMP necessary for exchange withUDP-glucose in the cytosol. The activities of UMP/CMP kinase-1 and adenylate kinase-1 couple theenergetic requirements of this cycle to the net conversion of ATP to AMP. Thus, increased ENTPD5activity leads to a decrease in the cellular ATP/AMP ratio. Because this ratio is the major determinantof glucose flux through the phosphofructokinase (PFK) step in glycolysis, a lowered ATP/AMP ratioincreases glycolysis, elevates lactate production, and provides glycolytic intermediates for biomassproduction.


Scholnick, P., Lang, D., and Racker, E. (1973).

J. Biol. Chem. 248, 5175.

Trombetta, E.S., and Helenius, A. (1999). EMBO J.

18, 3282–3292.

Vander Heiden, M.G., Cantley, L.C., and Thomp-

son, C.B. (2009). Science 324, 1029–1033.

Vander Heiden, M.G., Locasale, J.W., Swanson,

K.D., Sharfi, H., Heffron, G.J., Amador-Noguez,

D., Christofk, H.R., Wagner, G., Rabinowitz, J.D.,

Asara, J.M., and Cantley, L.C. (2010). Science

329, 1492–1499.

Vembar, S.S., and Brodsky, J.L. (2008). Nat. Rev.

Mol. Cell Biol. 9, 944–957.


Leading Edge

Essay

Glycomics Hits the Big TimeGerald W. Hart1,* and Ronald J. Copeland1

1Department of Biological Chemistry, School of Medicine, Johns Hopkins University, 725 North Wolfe Street, Baltimore, MD 21205-2185, USA


Cells run on carbohydrates. Glycans, sequences of carbohydrates conjugated to proteins andlipids, are arguably themost abundant and structurally diverse class of molecules in nature. Recentadvances in glycomics reveal the scope and scale of their functional roles and their impact onhuman disease.

By analogy to the genome, transcriptome,

or proteome, the ‘‘glycome’’ is the

complete set of glycans and glycoconju-

gates that are made by a cell or organism

under specific conditions. Therefore,

‘‘glycomics’’ refers to studies that attempt

to define or quantify the glycome of a cell,

tissue, or organism (Bertozzi and Sasise-

kharan, 2009). In eukaryotes, protein

glycosylation generally involves the cova-

lent attachment of glycans to serine,

threonine, or asparagine residues. Glyco-

proteins occur in all cellular compart-

ments. Glycans are also attached to

lipids, often ceramide, which is comprised

of sphingosine, a hydrocarbon amino

alcohol and a fatty acid. Complex glycans

are mainly attached to secreted or cell

surface proteins, and they do not cycle

on and off of the polypeptide. In contrast,

the monosaccharide O-linked N-acetyl-

glucosamine (O-GlcNAc) cycles rapidly

on serine or threonine residues of many

nuclear and cytoplasmic proteins. Identi-

fying the number, structure, and function

of glycans in cellular biology is a daunting

task but one that has been made easier in

recent years by advances in technology

and by our growing appreciation of how

integral glycans are to biology (Varki

et al., 2009).

The scope of the glycomics challenge is

immense. The covalent addition of

glycans to proteins and lipids represents

not only the most abundant posttransla-

tional modification (PTM), but also by far

the most structurally diverse. Although it

is commonly stated that more than 50%

of all polypeptides are covalently modified

by glycans (Apweiler et al., 1999), even

this estimate is far too low because it fails

to include that myriad nuclear and

cytoplasmic proteins are modified by

O-GlcNAc (Hart et al., 2007). Even though

the generic term ‘‘glycosylation’’ is often

used to categorize and lump all glycan

modifications of proteins into one bin,

side by side with other posttranslational

modifications such as phosphorylation,

acetylation, ubiquitination, or methylation,

such a view is not only inaccurate, but

also is completely misleading. If one only

considers the linkage of the first glycan

to the polypeptide in both prokaryotic

and eukaryotic organisms, there are at

least 13 different monosaccharides and

8 different amino acids involved in glyco-

protein linkages, with a total of at least

41 different chemical bonds known to be

linking the glycan to the protein (Spiro,

2002). Importantly, each one of these

unique glycan:protein linkages is surely

as different in both structure and function

as protein methylation is from acetylation.

Of course, this modification is not only

about a single linkage. When structural

diversity of the additional oligosaccharide

branches of glycans and the added diver-

sity of complex terminal saccharides on

glycans, such as fucose or sialic acids

(about 50 different sialic acids are known

[Schauer, 2009]), are taken into account,

the molecular diversity and varied func-

tions of protein-bound glycans rapidly

increase exponentially. Just the ‘‘sia-

lome’’ (Cohen and Varki, 2010) rivals or

exceeds many other posttranslational

modifications in abundance and struc-

tural/functional diversity. In addition,

chemical modifications, such as phos-

phorylation, sulfation, and acetylation,

increase the glycan structural/functional

diversity even more. Thus, categorizing

glycosylation as a single type of post-

translational modification is neither useful

nor at all reflective of reality.

Dynamic Structural ComplexityUnderlies Glycan FunctionsGlycoconjugates provide dynamic struc-

tural diversity to proteins and lipids that

is responsive to cellular phenotype, to

metabolic state, and to the developmental

stage of cells. Complex glycans play crit-

ical roles in intercellular and intracellular

processes, which are fundamentally

important to the development of multicel-

lularity (Figure 1). Unlike nucleic acids and

proteins, glycan structures are not hard-

wired into the genome, depending upon

a template for their synthesis. Rather,

the glycan structures that end up on

a polypeptide or lipid result from the

concerted actions of highly specific gly-

cosyltransferases (Lairson et al., 2008),

which in turn are dependent upon the

concentrations and localization of high-

energy nucleotide sugar donors, such as

UDP-N-acetylglucosamine, the endpoint

of the hexosamine biosynthetic pathway.

Therefore, the glycoforms of a glycopro-

tein depend upon many factors directly

tied to both gene expression and cellular

metabolism.

There are at-least 250 glycosyltrans-

ferases in the human genome, and it has

been estimated that about 2% of the

human genome encodes proteins

involved in glycan biosynthesis, degrada-

tion, or transport (Schachter and Freeze,

2009). Biosynthesis of the nucleotide

sugar donors is directly regulated by nu-

cleic acid, glucose, and energy metabo-

lism, and the compartmentalization of

these nucleotide sugar donors is highly

regulated by specific transporters. Protein

glycosylation is therefore controlled by

rates of polypeptide translation and

protein folding, localization of and compe-

tition between glycosyltransferases,


cellular concentration and localization of

nucleotide sugars, the localization of

glycosidases, and membrane trafficking.

Thus, individual glycosylation sites on the

same polypeptide can contain different

glycan structures that reflect both the

type and status of the cell in which they

are synthesized. For example, the glyco-

forms of the membrane protein Thy-1 are

very different in lymphocytes than they

are in brain, despite having the same poly-

peptide sequence (Rudd and Dwek,

1997). Conversely, even small changes in

polypeptide sequence or structure will

alter the types of glycan structures

attached to a polypeptide. For example,

histocompatibility antigen polypeptides

with more than 90% sequence homology

contain different N-linked glycan profiles

at individual sites, reflective of their

allelic type, even when they are synthe-

sized within the same cells (Swiedler

et al., 1985). Thus, site-specific protein

glycosylation is highly regulated by

gene expression of glycan-processing

enzymes, by polypeptide structure at all

levels, and by cellular metabolism.

Technology of GlycomicsA detailed understanding of cellular

processes will require a detailed appreci-

ation of the glycans modulating proteins

and pathways. Although this ultimate

goal of glycomics is laudable, we are

a very long way from having the tech-

nology to completely characterize the gly-

come of even a simple cell or tissue. Not

only is the glycome much more complex

than the genome, transcriptome, or pro-

teome, as noted above, it is also much

more dynamic, varying considerably not

only with cell type, but also with the

developmental stage and metabolic state

of a cell. Even very conservative esti-

mates indicate that there are well over

a million different glycan structures in

a mammalian cell’s glycome. However,

upon considering ‘‘functional glycomics,’’

it is estimated that the binding sites of

glycan-binding proteins (GBPs), such as

antibodies, lectins, receptors, toxins, mi-

crobial adhesions, or enzymes (Figure 1),

can accommodate only up to two to six

monosaccharides within a glycan struc-

ture (Cummings, 2009). Therefore, the

number of specific glycan substructures

that bind to biologically important GBPs

in a cell may be fewer than 10,000,

a number that is within the realm of

current analytical and, if targeted, chemi-

cal or enzymatic synthetic capabilities.

Until recently, the lack of tools and the

inherent complexity of glycans have

been major barriers preventing most biol-

ogists from embracing the importance of

glycans in biology. Recent technological

advances have significantly lowered these

barriers. Indeed, the tools of glycomics

and the subfields of glycoproteomics, gly-

colipidomics, and proteoglycomics have

all progressed substantially in recent

years (Krishnamoorthy and Mahal, 2009;

Laremore et al., 2010). Major technolog-

ical advances, many of which are shared

with proteomics, have recently allowed

semiquantitative profiling of glycans and

glycoproteins (Krishnamoorthy and

Mahal, 2009; Vanderschaeghe et al.,

2010). Some of these advances are the

result of the National Institute of General

Medical Science’s (NIGMS) support of

the Consortium for Functional Glycomics

(CFG), which has served to focus and

assist more than 500 researchers on

issues related to glycomics (Paulson

et al., 2006; Raman et al., 2006).

Kobata and colleagues were among the

first to profile N-glycans, well before the

current concepts of glycomics were

conceived. Despite the lack of many

modern methods, their pioneering work

was characterized by a high level of rigor

in defining the arrays of N-glycan struc-

tures present in cells and tissues and on

specific proteins (Endo, 2010). Currently,

a wide variety of high-resolution and

highly sensitive methods are available,

including capillary electrophoresis (CE),

high-performance liquid chromatography

(HPLC), and lectin microarrays.

Glycans are often profiled after their

release from polypeptides, which results in

the loss of any information about proteins

and sites to which they were attached.

Even though it is much more difficult, it is

also much preferable to perform

Figure 1. Glycans Permeate Cellular BiologyComplex glycans at the cell surface are targets of microbes and viruses, regulate cell adhesion and devel-opment, influence metastasis of cancer cells, and regulate myriad receptor:ligand interactions. Glycanswithin the secretory pathway regulate protein quality control, turnover, and trafficking of molecules toorganelles. Nucleocytoplasmic O-linked N-acetylglucosamine (O-GlcNAc) has extensive crosstalk withphosphorylation to regulate signaling, cytoskeletal functions, and gene expression in response to nutrientsand stress.


glycopeptide profiling (glycoproteomics) to

first identify attachment sites prior to

detailed profiling or structural analysis of

the glycans present on a polypeptide. The

ultimate goal of glycoproteomics, which is

todefine all of the molecular species (glyco-

forms) of glycoproteins in a cell or tissue,

has not yet been realized for any glycopro-

tein with more than one glycan attachment

site. N-glycans are generally released from

proteins by peptide-N-glycosidase F

(PNGase F), which cleaves most, but not

all, N-glycans. Unfortunately, no such

broadly specific enzyme exists for

O-glycans, which are generally released

by chemical methods, such as alkali-

induced b elimination, or by hydrazinolysis.

However, for relatively pure glycoproteins,

so called ‘‘top-down’’ mass spectrometric

methods, which do not involve prior release

of the glycans, may eventually prove useful,

as instrumentation and methods improve

(Reid et al., 2002).

Due to the small sample sizes involved,

most CE or HPLC separation methods

require chemical modification of released

glycans with fluorescent compounds. CE

and HPLC methods provide high-resolu-

tion separation of glycans, and when

combined with laser-induced fluorescent

detection (LIF), tagged glycans can be de-

tected in the low femtomole range. High

pH anion-exchange chromatography

(HPAEC) with pulsed-amperometric

detection separates glycans with high

resolution and detects them with high

sensitivity without chemical modification,

but the high alkalinity employed can be

problematic for some labile structures.

Lectins, which are defined as carbohy-

drate-binding proteins that are neither

antibodies nor enzymes, have a wide

range of glycan binding specificities, suit-

able for partial characterization of a gly-

come. Lectin microarrays use methods

and equipment similar to that employed

for nucleic acid arrays. Given the large

number of different lectins available, lectin

microarrays can provide information

about the glycome in a high-throughput

fashion, which is particularly useful in

profiling glycans produced by infectious

organisms (Hsu et al., 2006). In the future,

it is highly likely that glycomics will play

a central role in combating infectious

disease. However, many technical issues

remain to be resolved, such as standard-

ization required for clinical use, the

development of purified recombinant lec-

tins, and better definition of the specific-

ities of many lectins (Gupta et al., 2010).

Both matrix-assisted laser desorption

ionization (MALDI) and electrospray

mass spectrometry have played a key

role in glycan profiling and in glycoproteo-

mics (An et al., 2009; North et al., 2010;

Zaia, 2010). For biomarker discovery,

affinity enrichment approaches, based

upon chemical modification and solid-

phase extraction of N-linked glycopro-

teins, have proven useful in profiling

N-linked glycoprotein sites from serum-

or even from paraffin-embedded tissues

(Tian et al., 2009). Recently, using lectin

binding combined with advanced mass

spectrometric methods, thousands of

N-glycan attachment sites have been

mapped, a prerequisite for understanding

their functions (Zielinska et al., 2010).

Given the structural diversity of

glycans, all of these glycomic approaches

generate vast amounts of data. Glycan bi-

oinformatics has made great strides

within recent years with major efforts

from several laboratories (Aoki-Kinoshita,

2008). At least four major publicly

available carbohydrate databases (Glyco-

sciences.de, KEGG GLYCAN, Euro-

carbDB, and CFG) are now maintained,

and efforts to structure them in a uniform

format have been in progress for quite

some time. In addition, the Carbohy-

drate-Active EnZyme database (CAZy)

has played a key role in providing a global

understanding of carbohydrate active

enzymes, documenting their evolutionary

relationships, providing a framework for

elucidating common mechanisms, and

establishing the relationship between gly-

cogenomics and glycomes expressed by

cells (Cantarel et al., 2009). Moreover,

recent advances in bioinformatic analysis

tools for complex glycomic mass spec-

trometry data sets have allowed complex

data to be presented in formats useful to

nonexperts in all fields of biology (Ceroni

et al., 2008; Goldberg et al., 2005).

Perhaps one of the most important

contributions to the field of functional

glycomics has been the development of

well-defined glycan microarrays, which

currently display more than 500 different

glycan structures (Smith et al., 2010).

The NIGMS-supported Consortium for

Functional Glycomics (CFG) has gener-

ated and made publicly available

custom-made DNA microarrays that

represent glycosyltransferases and

glycan-binding proteins. The CFG also

has developed databases that present

phenotypic and biochemical data on gly-

cosyltransferase knockout mice. Even

though knocking out a single glycosyl-

transferase gene often affects hundreds

of glycoconjugates and myriad biological

processes, these mutant mice have

proven valuable in revealing the funda-

mental biological importance of glycans.

The microarrays and the databases

produced by the CFG member community

at large are publically available on the CFG

website (http://www.functionalglycomics.

org) and have resulted in a profound

increase in our understanding of the

binding specificities of GBPs, including

lectins key to inflammation and immunity,

and on infectious microbes or viruses.

However, a major barrier preventing

glycan biology from being incorporated

more into the mainstream is the continued

failure by the community to adopt a univer-

sally standard glycan structural format

and database that are easily accessed

worldwide. Most importantly, glycan data-

bases must eventually be incorporated

into standard interactive databases that

are supported by public agencies (such

as NCBI or EMBL) before glycan biology

can be fully integrated into the wider

research community.

From Glycomics to BiologyGlycans are directly involved in almost

every biological process and certainly

play a major role in nearly every human

disease (Figure 1). Genetic studies in

tissue culture cells indicate that specific

complex glycan structures are generally

not essential to a cell growing in culture,

indicating that most of the functions of

complex glycans are at the multicellular

level. In contrast, the cycling monosac-

charide, O-GlcNAc, on nuclear and cyto-

plasmic proteins, is essential even at the

single cell level in mammals (Hart et al.,

2007).

The critical roles of glycans in mammals

are now well established not only by the

dearth of mutations in glycan biosynthetic

enzymes that survive development, but

also by the severe phenotypes generated

when such mutations are not lethal.

These severe phenotypes are clearly illus-

trated by the congenital disorders of


glycosylation (CDGs) (Schachter and

Freeze, 2009), which are associated with

severe mental and developmental abnor-

malities. Also, the severe muscular

dystrophy that results from defective

O-glycosylation of a-dystroglycan (Yosh-

ida-Moriguchi et al., 2010) further

illustrates how a mutation in a glycan

biosynthetic enzyme results in a devas-

tating disease. The interplay between

O-GlcNAcylation and phosphorylation on

nuclear and cytoplasmic proteins plays

a key role in the etiology of diabetes,

neurodegenerative disease, and cancer

(Hart et al., 2007; Zeidan and Hart, 2010).

It has long been appreciated that alter-

ations in cell surface glycans contribute to

the metastatic and neoplastic properties

of tumor cells (Taniguchi, 2008). The func-

tions of many receptors are modulated by

their glycans, such as modulation of

Notch receptors by the action of specific

glycosyltransferases (Moloney et al.,

2000), which regulate Notch’s activation

by its ligands, affecting many develop-

mental events. Selectins, which specifi-

cally bind to a subset of fucosylated and

sialylated glycans, play a critical role in

leukocyte homing to sites of inflamma-

tion. Indeed, a selectin inhibitor is

currently in phase two clinical trials for

vaso-occlusive sickle cell disease (Chang

et al., 2010). Siglecs, which are a family of

cell surface sialic acid-binding lectins,

play a fundamental role in regulating

lymphocyte functions and activation.

Recent studies on galectins, a family of

b-galactoside-binding lectins, have

shown that they play a critical role in the

organization of receptors on the cell

surface and play important roles in immu-

nity, infections, development, and inflam-

mation (Lajoie et al., 2009). Proteoglycans

and glycosaminoglycans play a key role in

the regulation of growth factors, in micro-

bial binding, in tissue morphogenesis, and

in the etiology of cardiovascular disease.

Proteoglycans are perhaps the most

complicated and information-rich mole-

cules in biology, and progress in proteo-

glycomics has begun to accelerate

(Ly et al., 2010). Nearly all microbes and

viruses that infect humans bind to cells

by attaching to specific cell surface

glycans. Glycomics and glycan arrays

will have a substantial impact upon future

research toward both diagnosing and

preventing infectious disease.

Some of the most important drugs on the

market are already the result of glycomics.

The anti-flu virus drugs Relenza and Tami-

flu are structural analogs of sialic acids that

inhibit the flu virus neuraminidase and the

transmission of the virus. Natural heparin,

a sulfated glycosaminoglycan, and chemi-

cally defined synthetic heparin oligosac-

charides have long been widely used in

the clinic as anticoagulants and for many

other clinical uses. Hyaluronic acid, a non-

sulfated glycosaminoglycan, is used in the

treatment of arthritis. Many recombinant

pharmaceuticals, including therapeutic

monoclonal antibodies, are glycoproteins,

and their specific glycoforms are key to

their bioactivity and half lives in circulation

and to their possible induction of delete-

rious immune responses when they do

not contain the correct glycans. Given this

landscape, the pharmaceutical industry

and the US Food and Drug Administration

are rapidly realizing the critical importance,

in terms of both bioactivity and safety, of

carefully defining the glycoforms of any

therapeutics derived from glycoconju-

gates.

Glycoproteomics, Glycolipidomics,and BiomarkersClinical cancer diagnostic markers are

often glycoproteins, but most current

diagnostic tests only measure the expres-

sion of the polypeptide. Clearly, given the

long known alterations in glycans associ-

ated with cancer, it is highly likely that

cancer markers that detect specific glyco-

forms of a protein will have much higher

sensitivity and specificity for early

detection of cancer (Packer et al., 2008;

Taniguchi, 2008). Thus, the convergence

of glycomics and glycoproteomics is key

to the discovery of biomarkers for the early

detection of cancer (Taylor et al., 2009).

Recently, the Food and Drug Administra-

tion has approved fucosylated a-fetopro-

tein as a diagnostic marker of primary

hepatocarcinoma. In addition, fucosy-

lated haptoglobin may be a much better

marker of pancreatic cancer than simply

monitoring the expression of the hapto-

globin polypeptide. Indeed, The National

Cancer Institute has begun an initiative to

discover, develop, and clinically validate

glycan biomarkers for cancer (http://

glycomics.cancer.gov/). System biology

analyses of the glycome to identify

biomarkers of human disease will, by

necessity, also employ many of the same

methods used by genomics, proteomics,

metabolomics, and lipidomics (Figure 2)

(Packer et al., 2008). Due to the critical

roles of glycans in cardiovascular disease

and lung disease and in the functions of

blood cells, the National Heart Lung and

Blood Institute (NHLBI) has recognized

an acute need to train more researchers

in the area of glycosciences by creating

a ‘‘Program of Excellence in Glycoscien-

ces,’’ which will not only support collabo-

rative research, but will also provide

hands-on laboratory training in the

methods of glycosciences to fellows.

Thus, though our knowledge about the

biology of glycans and glycomics

continues to lag behind more mainstream

fields of genomics and proteomics, tech-

nological advances in glycomics in the

Figure 2. Glycomic Complexity Reflects Cellular ComplexityGiven that glycan structures are regulated by metabolism and glyco-enzyme expression and glycansmodify both proteins and lipids, functional glycomics also requires the tools of genomics, proteomics, lip-idomics, and metabolomics (modified after Packer et al., 2008).


last 5 years have begun to accelerate the

integration of glycobiology into the other

major fields of biomedical research. A

complete mechanistic understanding of

the etiology of almost any disease will

depend upon the elucidation of the func-

tions of all posttranslational modifications

but will especially depend upon our

understanding the many roles of glycans,

the most abundant and structurally

diverse type of posttranslational modifi-

cation.

ACKNOWLEDGMENTS

We thank Dr. Chad Slawson for helpful sugges-

tions. Original research in the author’s laboratory

was supported by NIH grants R01CA42486, R01

DK61671, and R24 DK084949.

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Leading Edge

Essay

What Determines the Specificityand Outcomes of Ubiquitin Signaling?Fumiyo Ikeda,1 Nicola Crosetto,1 and Ivan Dikic1,*1Frankfurt Institute for Molecular Life Sciences and Institute of Biochemistry II, Goethe University School of Medicine, Theodor-Stern-Kai 7,

D-60590 Frankfurt (Main), Germany*Correspondence: [email protected]

DOI 10.1016/j.cell.2010.10.026

Ubiquitin signals and ubiquitin-binding domains are implicated in almost every cellular process, buthow is their functionality achieved in cells? We assess recent advances in monitoring the dynamicsand specificity of ubiquitin networks in vivo and discuss challenges ahead.

IntroductionA small protein modifier, ubiquitin, is the

building block of a repertoire of molecular

signals spanning from single ubiquitin

to ubiquitin chains of different linkage

used for posttranslational modification of

dozens of cellular proteins (Hershko and

Ciechanover, 1998). The seven lysines

(K) of ubiquitin (K6, K11, K27, K29, K33,

K48, and K63) and the amino-terminal

methionine (M1) are connected to the

C-terminal glycine for chain assembly,

generating polymers (Ikeda and Dikic,

2008; Iwai and Tokunaga, 2009). Ubiquitin

signals are recognized and processed

by specialized ubiquitin-binding domains

(UBDs) that form transient, noncovalent

interactions either with ubiquitin moieties

or with the linkage region in their chains.

So far, roughly 200 intracellular proteins

have been recognized to contain one

or more UBDs (Dikic et al., 2009). Ubiqui-

tin-UBD interactions regulate almost

every aspect of cellular physiology,

including protein degradation, receptor

trafficking, DNA repair, cell-cycle pro-

gression, gene transcription, autophagy,

and apoptosis (recently reviewed in

Deshaies and Joazeiro, 2009; Kirkin

et al., 2009; Raiborg and Stenmark,

2009; Ulrich and Walden, 2010; Wickliffe

et al., 2009; Winget and Mayor, 2010).

Yet, very little is known about the nature

of ubiquitin signals and the dynamics

of their interpretation by UBDs in the

highly crowded molecular environment

of the cell. In particular, it remains unclear

how a relatively limited pool of signals

(ubiquitin chains and UBDs) with partially

overlapping biochemical properties can

orchestrate the localization and function

of thousands of proteins involved in very

different cellular processes. Here we

summarize the most recent advances in

understanding specificity determinants

in ubiquitin signaling and discuss future

challenges in the development of sensi-

tive and reliable methods for monitoring

spatial and temporal patterns of ubiquiti-

nation in vivo.

Diversity of Ubiquitin SignalsDespite its relatively rigid globular struc-

ture, ubiquitin is one of the most versatile

signaling molecules in the cell. Although

the surface of ubiquitin is mostly com-

posed of polar residues, it is through its

few hydrophobic patches that it interacts

with most UBDs (Dikic et al., 2009; Winget

and Mayor, 2010). Moreover, the pres-

ence of seven lysine residues and the

N-terminal methionine within ubiquitin

that can be fused to the C-terminal di-

glycine motif of another ubiquitin allows

the formation of polymeric chains en-

dowed with flexibility, as in the case of

K63-linked or M1-linked chains (often

referred to as linear) (Ikeda and Dikic,

2008; Iwai and Tokunaga, 2009). K48-

linked and K11-linked chains adopt

a more rigid conformation, in which ubiq-

uitin monomers are tightly packed against

each other. This creates unique modules

composed of aligned ubiquitin moieties

in which the hydrophobic patch contain-

ing isoleucine 44 is either embedded or

facing out toward the surface (Pickart

and Fushman, 2004; Bremm et al., 2010;

Matsumoto et al., 2010). Conversely,

K6-linked chains form an asymmetric

compact conformation distinct from any

other known type of ubiquitin chain

(Virdee et al., 2010). The possibility of

heterotypic ubiquitin chains (that is, with

mixed linkages) has been shown in vitro,

but their presence and biological func-

tions in vivo remain to be confirmed. Alto-

gether, monoubiquitin and homotypic

polyubiquitin chains comprise no more

than ten signal types. However, the ability

to synthesize homotypic chains of various

lengths indicates that the repertoire of

ubiquitin signals in the cell may be larger

than expected.

Signals Decoders:Ubiquitin-Binding DomainsUbiquitin signals are read and processed

by UBDs that bind noncovalently to

mono- or polyubiquitin chains. To date,

five structural folds are known with

more than 20 UBDs identified overall

(Dikic et al., 2009). UBDs are commonly

a-helical structures, zinc fingers, pleck-

strin homology (PH) folds, or similar to

the ubiquitin-conjugating (Ubc) domain

present in E2 enzymes (Dikic et al.,

2009). In the majority of cases, isolated

UBDs preferentially bind to monoubiquitin

via a conserved hydrophobic patch sur-

rounding isoleucine 44. The measured

affinity of isolated UBDs for monoubiqui-

tin typically falls in the micromolar range

(Dikic et al., 2009; Winget and Mayor,

2010). In certain cases, monoubiquitin-

UBD interactions have also been demon-

strated in the context of endogenous full-

size proteins. For example, UBDs present

in Y family polymerases performing DNA

translesion synthesis bind the monoubi-

quitinated sliding clamp PCNA (Bienko

et al., 2005), and monoubiquitinated

transmembrane receptors are recognized


by endocytic sorting proteins containing

diverse UBDs (Hicke and Dunn, 2003).

The affinity of UBD-containing proteins

for their monoubiquitinated targets in the

cellular environment, however, may be

different from that inferred from in vitro

studies. In fact, the way ubiquitin signals

are decoded in cells may be influenced

by multiple factors, including the pres-

ence of tandem copies of one UBD in

the same protein, oligomerization, and

protein compartmentalization (reviewed

in Dikic et al., 2009; Winget and Mayor,

2010).

In addition to monoubiquitin, many

UBDs display either relative or absolute

selectivity for certain types of chains

(Ikeda and Dikic, 2008; Dikic et al., 2009;

Winget and Mayor, 2010). For instance,

the Pru (Plextrin receptor for ubiquitin)

domain in the proteasome receptor

Rpn13 preferentially interacts with K48-

linked diubiquitin (Husnjak et al., 2008),

and the NZF (Npl4 zinc finger) domain in

TAK1-binding protein 2 (TAB2) binds

specifically to K63-linked ubiquitin (Kula-

thu et al., 2009; Sato et al., 2009). In con-

trast, UBDs in NEMO and ABIN proteins

(UBAN) bind linear diubiquitin chains

with approximately 100-fold higher affinity

than K63 or K48 chains, and binding to

monoubiqutitin could not be detected

(Rahighi et al., 2009; Lo et al., 2009). The

selectivity of UBAN for linear chains has

been explained by the observation that

a NEMO dimer binds symmetrically to

linear diubiquitin, involving direct interac-

tions with residues exposed in the

glycine-methionine linkages (Rahighi

et al., 2009). In addition, the crystal struc-

tures of the NZF domain of TAB2 and

TAB3 in complex with K63-linked diubi-

quitin have shown a two-sided ubiquitin-

binding surface thanks to a flexible

K-linkage positioned away from the

interaction surface (Kulathu et al., 2009;

Sato et al., 2009). Linkage selectivity can

also result from multivalent interaction

between tandem UBD arrays in a given

protein and ubiquitin monomers or link-

ages in a polyubiquitin chain. Tandem

ubiquitin-interacting motifs (UIMs) in the

DNA double-strand break response pro-

tein Rap80 are positioned to cross two

K63-linked monomers, whereas Ataxin-3

UIMs display K48 avidity (Sims and

Cohen, 2009). The proteasome receptor

S5a has two UIMs separated by linker

regions and shows a 10-fold higher

affinity for diubiquitin over monoubiquitin

(Zhang et al., 2009). These observations

suggest that the function of tandem UBD

arrays is to increase the affinity for a given

ubiquitinated substrate rather than simul-

taneously binding multiple substrates.

Specificity and Plasticityof Ubiquitin SignalingHistorically, distinct ubiquitin signals have

been linked to specific cellular functions.

For example, K48-linked chains, also

known as ‘‘classical’’ ubiquitin chains,

were originally described as the signal

that targets substrates for proteasomal

degradation (Hershko and Ciechanover,

1998). Nonclassical linkage types, such

as K63-, K11-, or M1-linked chains are

instead associated with DNA repair

regulation, cell-cycle progression, innate

immunity, and inflammation (Ikeda and

Dikic, 2008; Iwai and Tokunaga, 2009;

Matsumoto et al., 2010; Wickliffe et al.,

2009). Recent reports, however, have

challenged the notion that distinct chain

types exclusively regulate specific pro-

cesses in the cell. For instance, nonclas-

sical ubiquitin signals, such as K11

chains generated by the anaphase-

promoting complex (APC/C), can also

target selected substrates for proteaso-

mal degradation (Jin et al., 2008). In yeast,

cyclin B1 is modified by a mix of K48-,

K63-, and K11-linked chains rather than

by K48 chains alone (Kirkpatrick et al.,

2006). This heterogeneous pool is suffi-

cient to bind to proteasomal ubiquitin

receptors and drive cyclin B1 degradation

(Kirkpatrick et al., 2006). Furthermore,

linear chains, initially discovered as acti-

vators of the NF-kB pathway (Tokunaga

et al., 2009), can also trigger proteasomal

degradation when fused to artificial sub-

strates (Zhao and Ulrich, 2010).

So, how is functional specificity of ubiq-

uitin signaling achieved in vivo? Even

though evidence indicates that specific

chain types control distinct molecular

processes, as clearly exemplified by

NF-kB signaling, we speculate that addi-

tional signals (monoubiquitin and chains

with different linkage and length) can

control the same molecular process with

different kinetics and spatial constraints.

It has also been speculated that unan-

chored ubiquitin chains can regulate

NF-kB activation (Xia et al., 2009).

However, the importance of this regula-

tory mechanism in vivo remains to be

further investigated. Therefore, the de-

coding of ubiquitin signals might be per-

formed in vivo by different UBDs (not

necessarily endowed with absolute selec-

tivity toward monoubiquitin or a particular

chain type) embedded in key proteins

controlling a particular process. Although

this scenario could allow a certain degree

of plasticity in ubiquitin signaling, speci-

ficity might be determined by the localiza-

tion and assembly of UBD-containing

proteins and enzymes catalyzing ubiquiti-

nation reactions.

Catching Ubiquitin Signalingin the ActThe huge discrepancy between our

current understanding of the ubiquitin

system from in vitro studies compared to

in vivo models stems from the fact that

ubiquitination and its recognition and

cleavage occur in milliseconds (Pierce

et al., 2009), therefore making it chal-

lenging to analyze these events in living

systems. The first attempts to study ubiq-

uitin signaling in vivo have used anti-

bodies against monoubiquitin, polyubi-

quitin chains, or, more recently, selective

linkages, including K11, K48, K63, and

linear chains (Matsumoto et al., 2010;

Newton et al., 2008; Wang et al., 2008;

Tokunaga et al., 2009) (Figure 1A). Raising

linkage-selective antibodies is not easy,

despite being urgently needed to provide

tools to discriminate between different

chain types in the cell. These antibodies

were produced either by synthesizing

peptides resembling specific linkage

bonds (Wang et al., 2008; Tokunaga

et al., 2009) or by using the phage-display

method (Matsumoto et al., 2010; Newton

et al., 2008). Although chain-selective

antibodies have been used to demon-

strate specific chain formation in several

biological settings (such as the NF-kB

pathway and cell-cycle progression), their

ability to monitor substrates with low

abundance and the dynamics of chain

(de)conjugation as well as their distribu-

tion in vivo are still very limited.

Monoclonal antibodies recognizing di-

glycine-modified lysines have been used

in combination with mass spectrometry

in efforts to increase the sensitivity of

immune-based techniques (Xu et al.,

2010) (Figure 1B). These antibodies enrich


for the C-terminal di-glycine

motif of ubiquitin attached to

the acceptor lysine following

proteolysis of ubiquitinated

proteins by trypsin (Fig-

ure 1B). This method revealed

more than 200 ubiquitinated

proteins from human embry-

onic kidney 293 cells, the

majority of which were previ-

ously unknown targets (Xu

et al., 2010). This strategy

can be coupled to stable

isotope labeling with amino

acids in cell culture (SILAC)

to quantitatively explore pro-

tein ubiquitination in diverse

biological settings. However,

it needs to be noted that

this approach can neither

detect short-lived proteins

nor distinguish lysine modifi-

cation by NEDD8.

The AQUA (absolute quan-

tification) method developed

in the Gygi laboratory is

another promising approach

to measure the dynamics of

ubiquitin signaling in cells

(Kirkpatrick et al., 2005).

AQUA relies on the use of

stable isotope-labeled inter-

nal standard peptides that

mimic those produced during

tryptic digestion of ubiquiti-

nated proteins of interest.

In case of mono- or polyubi-

quitinated proteins, tryptic

digestion produces a series

of unbranched and di-glycine-

branched peptides. Initial analysis of

such mixtures allows identification of

ubiquitination sites in the substrate and

the type of ubiquitin chain linkage (such

as monoubiquitination or K63- or K48-

ubiquitin chains). Based on this informa-

tion, substrate-, site-, and linkage-specific

reference peptides are synthesized and

used as quantitative internal standards,

allowing for precise quantification of

monoubiquitin and polyubiquitin chains

by targeted proteomics approaches such

as selective reaction monitoring. With

this methodology, the stoichiometry of

ubiquitin moieties on a protein of interest

can be determined (Figure 2A). Its

simplicity and sensitivity, coupled with

the current widespread availability of

tandem mass spectrometers, makes

AQUA the tool of choice for quantitatively

measuring ubiquitin modifications directly

in cell lysates (Kirkpatrick et al., 2006).

What Is Known about UbiquitinChain Length In Vivo?The methods described above are pre-

dicted to provide quantitative information

on the repertoire of ubiquitin signals and

ubiquitinated proteins generated in dif-

ferent biological settings. However, these

methods cannot monitor the length of

ubiquitin chains in vivo. At present, all

our knowledge on their length in vivo

relies on nonquantitative analysis of

immunoblots. Several procedures have

been designed to cause ubiquitin chains

and polyubiquitinated substrates to accu-

mulate in the cell to facilitate

their detection, including the

use of inhibitors of the pro-

teasome and of deubiquiti-

nating enzymes (DUBs). This

has often led to the conclu-

sion that high-mobility ubiqui-

tin-positive smears observed

on immunoblots represent

the natural modification of

substrates by very long ubiq-

uitin chains. This, however,

can be misleading because

the combination of different

ubiquitin signals (monoubi-

quitin or ubiquitin chains) on

the same type of substrate

can also yield high-mobility

smears (Haglund et al.,

2003; Huang et al., 2006),

and inhibition of DUBs and

the proteasome may cause

an overrepresentation of

long ubiquitin chains that

might not naturally occur in

the cell.

The question of chain

length is important given that

chains with different topology

and length may regulate dif-

ferent cellular functions. For

instance, the length of K48-

linked tetraubiquitin chains

is optimized for interaction

with proteasomal receptors

(Pickart and Fushman, 2004),

as a ternary complex can

be formed between the ubiq-

uitin chains and proteasomal

receptors Rpn13 and S5a (Zhang et al.,

2009). Moreover, given that trimming

of ubiquitinated substrates occurs from

the distal end of the chains, it seems

that the length of K48-linked chains

dictates the duration of proteasomal

degradation (Lee et al., 2010).

Monoubiquitination can also drive pro-

teins to proteasomal degradation (Shabek

et al., 2009). These observations collec-

tively suggest that the ubiquitin chain

length required for proteasomal degrada-

tion is determined by the substrate’s

affinity for the proteasome and must be

just high enough to allow processivity of

the proteolytic process. This kind of

adjustment is most likely controlled by

a proteasome-associated complex,

which is equipped with both ubiquitin

Figure 1. Antibodies for Ubiquitin Signals(A) Linkage-specific antibodies, such as a-lysine 11(K11)-, a-K48-, a-K63-linked ubiquitin chains and a-linear ubiquitin chains, can be applied for thedetection of endogenous ubiquitination of a specific linkage type.(B) After trypsin digestion of total cell extracts, immunoprecipitation of thesamples by a specific antibody against glycine-glycine-lysine peptides(a-GGK Ab) can enrich fragments with ubiquitinated K residues from bothsubstrates and ubiquitin chains. Analysis by mass spectrometry enables theidentification of new target proteins as well as sites of ubiquitination.


ligase (HUL5) and deubiquiti-

nating (UBP6) activities (Cro-

sas et al., 2006).

In the case of the NF-kB

pathway, distinct activation

steps involve K63, linear,

and K48 chains (Bianchi and

Meier, 2009), which are

further edited (in length and

topology) by ligases and

DUBs (Wertz et al., 2004;

Newton et al., 2008). An initial

mechanism proposed for NF-

kB activation implicated long

K63-linked chains in the

recruitment of TAK1 and IKK

kinases via their respective

adaptor proteins TAB2 and

NEMO (reviewed in Bianchi

and Meier, 2009). This model

has been challenged by the

demonstration that cells ex-

pressing ubiquitin lacking

K63 have intact NF-kB

signaling via tumor necrosis

factor-a receptors (Xu et al.,

2009). Interestingly, based

on available structures it

appears that chain-selective

UBDs in TAB2 and NEMO

interact with K63-linked or

linear diubiquitin chains,

respectively (Kulathu et al.,

2009; Rahighi et al., 2009;

Sato et al., 2009). Given that

no data are available on the

precise length of ubiquitin

chains in the NF-kB pathway,

it is tempting to speculate

that diubiquitin chains are

the fundamental units recog-

nized by selective UBDs.

However, UBDs also show

promiscuous binding with

lower affinities to other types

of chains. Examples include

the NZF domain of TAB2,

which also binds K48 chains

in solution (Kulathu et al.,

2009), and the UBAN domain

in NEMO, which interacts

with K63- and K48-linked

chains longer than diubiquitin

(Rahighi et al., 2009). We

speculate that diubiquitin

units in longer chains may

amplify signals that can be

recognized by nonselective

UBDs. In such a scenario,

short ubiquitin chains added

to substrates will be preferen-

tially decoded by linkage-

selective UBDs, whereas

long chains may be promis-

cuously read by different

UBDs, possibly placing chain

length next to chain linkage

type in determining ubiquitin-

UBD selectivity.

Development of SensorsUsing Selective UBDsIn order to measure the

dynamics of ubiquitin chain

formation/disassembly and

their length in vivo, functional

ubiquitin sensors are needed

(Figure 2B). A recently engi-

neered sensor (TUBE, tan-

dem repeated ubiquitin enti-

ties) possesses four tandem

UBA domains of either HR23

or ubiquitin 1 (Hjerpe et al.,

2009). The ubiquitin-binding

capacity of TUBE is markedly

higher for ubiquitin tetramers

in comparison to monoubi-

quitin. In addition, the affinity

of the interaction of TUBE

with either K63- or K48-tet-

raubiquitin chains is much

greater than that of a single

UBA domain (Hjerpe et al.,

2009). An intriguing feature

of TUBE is its ability to pro-

tect ubiquitin chains from

cleavage by blocking acces-

sibility to DUBs.

The design principle of

TUBE could be easily adap-

ted to other UBDs: for ex-

ample, a K63 chain-specific

sensor could be created by

fusing multiple NZF domains

of TAB2 in tandem, a K48-

specific sensor by merging

multiple Pru domains of

Rpn13, and a linear-specific

sensor by arraying several

copies of the UBAN domain

of NEMO or ABINs. These

UBD-derived ubiquitin sen-

sors could be used to protect

and purify substrates deco-

rated with endogenous ubiq-

uitin chains. They could also

Figure 2. Quantification and Detection of Ubiquitin Chains In Vivo(A) The workflow for the AQUA (absolute quantification) method of quantitativemass spectrometry is depicted. First, a representative tryptic peptide is selectedbased on initial proteomic sequencing experiments and then synthesized witha stable isotope at one residue for identification. The tryptic peptide sequencefor lysine48 (K48)-linkedubiquitinchains is indicated (upper panel).AQUApeptidestandards are added to the sample (cell lysates or immunocomplexes) prior totrypsin digestion and targeted proteomic analysis is performed using selectivereaction monitoring. The amount of total protein and the extent of ubiquitinationat that particular site can be determined by comparing the precise amounts ofthe unmodified and ubiquitinated versions of the peptide (lower panel).(B) Schematic models of ubiquitin sensors are shown. By using different ubiq-uitin-binding domains (UBDs), the sensor can be applied for specific linkagetype of ubiquitin chains (left), such as K48, K63, and linear chains. TandemUBDs may be used to determine the chain length (right). One UBD recognizes1 unit of diubiquitin. The tag chosen depends on the experimental purposes.


be used to determine the predominant

linkage type within these chains by

competition experiments and for

measuring the length of ubiquitin poly-

mers in their natural environment.

A further critical challenge will be to

evaluate chain-specific ubiquitin sensors

using advanced (high-throughput) single-

cell or -molecule microscopy. This might

permit the qualitative and quantitative

assessment of ubiquitin chain formation

and the interplay between different chain

types in vivo. Analyzing additional proper-

ties, such as the spatial and temporal

regulation of conjugation and deconjuga-

tion of ubiquitin chains as well as their

length in vivo, could enable a high-

resolution, systems-level analysis of the

‘‘ubiquitinome.’’

PerspectiveEven though we have attained a sophisti-

cated mechanistic understanding of the

ubiquitin system, it has been more difficult

to analyze the orchestration of its func-

tions in vivo. Within the cellular environ-

ment, ubiquitin signals must select the

correct binding partner at the right place

and time, ensuring accurate signaling.

To understand the specificity and

dynamics of the ubiquitin system in its

biological context, it is critical that highly

sensitive methods, such as mass spec-

trometry and advanced microscopy, are

deployed to measure key parameters,

such as the amount of different ubiquitin

signals, the kinetics of UBD-ubiquitin

recognition, and the type and length of

ubiquitin chains attached onto substrates

in vivo. By shedding light onto these prop-

erties, we will gain a deeper under-

standing of one of the most important

and widely used regulatory systems of

cell physiology.

ACKNOWLEDGMENTS

We are grateful to C. Behrends, A. Ciechanover, K.

Rittinger, and S. van Wijk for comments and

discussions. Research in the I.D. laboratory is sup-

ported by the Deutsche Forschungsgemeinschaft,

the Cluster of Excellence ‘‘Macromolecular

Complexes’’ of the Goethe University Frankfurt

(EXC115), and the European Research Council

under the European Union’s Seventh Framework

Programme (FP7/2007-2013)/ERC grant agree-

ment n� [250241-LineUb].

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Leading Edge

Minireview

Ubiquitin: Same Molecule,Different Degradation PathwaysMichael J. Clague1,* and Sylvie Urbe1,*1Cellular and Molecular Physiology, Institute of Translational Medicine, University of Liverpool, Crown Street, Liverpool L69 3BX, UK

*Correspondence: [email protected] (M.J.C.), [email protected] (S.U.)DOI 10.1016/j.cell.2010.11.012

Ubiquitin is a common demoninator in the targeting of substrates to all three major protein degra-dation pathways in mammalian cells: the proteasome, the lysosome, and the autophagosome. Thefactors that direct a substrate toward a particular route of degradation likely include ubiquitin chainlength and linkage type, which may favor interaction with particular receptors or confer differentialsusceptibility to deubiquitinase activities associated with each pathway.

The dynamic state of bodily proteins was established by

analyzing the fate of stable isotope-labeled amino acids that

had been fed to mice. These classic experiments, conducted

by Rudolf Schoenheimer in the late 1930s, presage modern

stable isotope labeling techniques (such as SILAC), which allow

determination of the turnover rate of hundreds to thousands of

individual proteins in a single mass spectrometry experiment

(Kristensen et al., 2008). After its discovery, the lysosomal

compartment was considered the principal site of degradation

of cellular proteins, through the action of resident acid-depen-

dent proteases. However, this view perished with the demon-

stration that the half-lives of most cellular proteins are insensitive

to alkalinization of the lysosomes. The subsequent discovery of

the ubiquitin-proteasome degradation system as the major route

to protein degradation generated a new orthodoxy. Central to

this model is the idea that covalent modification of substrate

proteins with a polypeptide ubiquitin tag targets them to the large

(26S) proteolytic complex known as the proteasome.

It came then as a surprise to discover that ubiquitin tagging also

provides a signal to route endocytosed receptors to the lyso-

somal degradation pathway and more recently to mark organ-

elles for disposal by the third major cellular degradative pathway

of autophagocytosis. The role of ubiquitin in protein degradation

is more ubiquitous than once thought (Figure 1). In this Minire-

view, we consider how a ubiquitin tag selects for specific degra-

dation pathways and also highlight the interplay between these

pathways that a shared dependence on ubiquitin engenders.

General ConsiderationsSubstrate proteins are selected for modification of lysine resi-

dues by ubiquitin through interaction with an E3 ligase protein

that recruits an E2-enzyme charged with ubiquitin. This can

result in transfer of a single ubiquitin molecule (monoubiquitina-

tion) or coupling of further ubiquitin molecules, through integral

lysine residues, to form a chain. The seven lysines of ubiquitin

provide for the formation of different isopeptide chain linkages,

which adopt different three-dimensional structures, and all of

which are represented in eukaryotic cells (Xu et al., 2009). The

specific combination of E2 and E3 enzymes recruited to

a substrate dictates the chain linkage type. The human genome

encodes more than 20 different types of ubiquitin-binding

domains, and proof of principle for linkage specificity of binding

has been established (see Essay by F. Ikeda, N. Crosetto, and

I. Dikic on page 677 of this issue). One means to achieve this is

through the spatial arrangement of tandem ubiquitin-binding

domains (UBDs) either encoded in a single protein or by

combining domains within a multimolecular complex, such that

simultaneous occupancy of two binding sites is restricted to

particular chain configurations.

Proteasomal DegradationEarly work suggested that proteasomal targeting requires

a lysine 48 (K48)-linked ubiquitin chain consisting of at least

four conjoined ubiquitin molecules. This was based first upon

the biochemical analysis of chains formed on a model substrate,

b-galactosidase, in a reticulocyte lysate system and second

upon studies showing that unique among lysine mutant versions

of ubiquitin, K48R cannot serve as the sole source of ubiquitin in

yeast (Finley, 2009; Xu et al., 2009). The affinity of unanchored

K48 polyubiquitin chains for the proteasome increases more

than 100-fold from di- to tetraubiquitin (�170 nM) and less

steeply thereafter (Thrower et al., 2000).

A body of work now suggests that in fact the proteasome

happily accepts other ubiquitin chain types. Indirect evidence

for this comes from the observation that acute proteasome inhi-

bition does not lead to the selective accumulation of K48 chains.

Rather, all chain types with the exception of K63 are increased

(Jacobson et al., 2009; Xu et al., 2009). During cell division, the

human anaphase-promoting complex (APC/C) recruits two E2

ligases (UbcH10 and Ube2S), which combine to exclusively

generate K11-linked chains on substrates. Loss of this unit leads

to strong defects in mitotic progression due to failure of the

necessary degradation processes (Song and Rape, 2010).

In vitro studies have even shown that K63-modified dihydrofo-

late reductase provides an efficient proteasome substrate

(Hofmann and Pickart, 1999).

The proteasome is composed of a core (20S) particle contain-

ing multiple proteolytic sites and a 19S regulatory particle that


governs access to the core. To enter the core, substrates must

be amenable to unfolding by a hexamer of ATPases associated

with the base of the regulatory particle. Other constituents of the

regulatory particle are implicated in the recruitment of substrates

(Finley, 2009). Rpn10 and Rpn13 interact with ubiquitinated

substrates through UIM (ubiquitin-interacting motif) domains

and a Pru (pleckstrin-like receptor for ubiquitin) domain, respec-

tively. The UBL/UBA family of proteins are substoichiometric

components of purified proteasomes that bind ubiquitin via their

UBA (ubiquitin-associated) domain and the proteasome regula-

tory particle through its UBL (ubiquitin-like) domain. They are

proposed to remotely scavenge ubiquitinated substrates and

present them to the proteasome (Figure 2). Particular protea-

some-associated ubiquitin receptors have been linked with the

degradation of specific substrates (reviewed in Finley, 2009).

The mammalian regulatory particle has three associated deu-

biquitinating enzymes (DUBs), POH1/PSMD14, USP14, and

UCH37 (Rpn11 and Ubp6 in budding yeast), which have distinct

specificities for different chain linkages (Finley, 2009). What is the

function of these DUB activities? One important function is to

salvage ubiquitin in order to maintain the cellular ubiquitin pool.

The JAMM/MPN+ metalloprotease POH1 is thought to specifi-

cally disassemble K63-linked chains, as well as cleave the

isopeptide bond that links the substrate and the proximal ubiqui-

tin, allowing for en bloc removal of an ubiquitin chain. It also

governs entry into the central proteolytic chamber, thereby

coupling substrate degradation to recycling of ubiquitin (Yao

and Cohen, 2002). Ubiquitin-aldehyde-sensitive cysteine

protease activities (that is, USP14 and UCH37) account for all

activity directed toward K48-linked chains and also contribute

to K63-linked chain disassembly (Jacobson et al., 2009). One

attractive notion is that the integration of these DUB activities

may provide for a proof-reading mechanism, facilitating release

from the proteasome if commitment to degradation is not

accomplished within a given time window. For example, prefer-

ential proteasomal DUB activity against K63-linked chains has

been proposed to select against these substrates for degrada-

tion (Jacobson et al., 2009). Also in line with this principal,

Figure 1. Ubiquitin Is a Common Denominator of Protein Degrada-

tion PathwaysSpecific ubiquitin receptors are associated with each degradation pathway.Autophagosomal and multivesicular body (MVB) pathways merge at the lyso-some and share a dependence on v-ATPase activity (inhibited by bafilomycin).Both pathways also share sensitivity to inhibitors of phosphoinositide 3-kinaseactivity, such as wortmannin or 3-methyladenine, as the family memberhVPS34 is required both for recruitment of ESCRT (endosomal sortingcomplex required for transport) components to MVBs and for expansion ofthe double-membrane preautophagosomal structure. Proteasomal inhibitorsinclude lactacystin and epoxomicin.

Figure 2. Ubiquitin Recognition by the Major Degradative PathwaysDepiction of the ‘‘ubiquitin receptors’’ associated with each degradativepathway. The domain structures shown are for the human representatives ofeach protein family, except for yeast Ddi1, the human ortholog of whichdoes not contain a UBA domain. CB: clathrin-binding motif; CC: coiled coil;ESCRT: endosomal sorting complex required for transport; GGA: golgi-asso-ciated, gamma adaptin ear containing, ARF-binding protein; GAE: gammaadaptin ear; GAT: GGA and TOM1; GLUE: GRAM-like ubiquitin-binding inEap45; HRS: HGF receptor tyrosine kinase substrate; LIR: LC3-interactingregion; PB1: Phox and Bem1; PRU: Pleckstrin-like receptor for ubiquitin;SH3: Src homology domain 3; STAM: signal transducing adaptor molecule;TOM1: target of myb1; TSG101: tumor susceptibility gene 101; UBA: ubiqui-tin-associated domain; UBL: ubiquitin-like domain; UEV: ubiquitin E2 variantdomain; UIM: ubiquitin-interacting motif; VHS: Vps27, HRS, and STAM;VPS36: vacuolar protein sorting 36; vWFA: von Willebrand Factor type A;ZZ: zinc finger. Note the following gene names and commonly used alternativenames also apply: p62; SQSTM1 (sequestosome), NDP52; CALCOCO2,UBQLN1; PLIC1; DSK2. Domain annotation based on PFAM and UNIPROT.


a specific chemical inhibitor of USP14 has recently been shown

to enhance the rate of protein degradation (Lee et al., 2010).

In yeast, a ubiquitin ligase, Hul5 (mammalian ortholog is

KIAA10/E3a), that is associated with proteasomes can oppose

Ubp6 activity through chain elongation (E4) (Crosas et al.,

2006). Thus a balance between proteasome-associated ubiqui-

tin ligase and DUB activity may determine receptor fate.

Endolysosomal DegradationThe lysosomal degradation pathway is the principle means by

which a cell turns over plasma membrane proteins, such as recep-

tors or channels. Its defining characteristic is a requirement for

organelle acidification, mediated by the v-ATPase. Endocytosed

proteins are either recycled to the plasma membrane or captured

into lumenal vesicles of the multivesicular body (MVB) as it

matures from the sorting endosome, before fusing directly with

lysosomes. Some receptors use ubiquitin as an internalization

signal, but for other ubiquitinated receptors, such as epidermal

growth factor receptor, this is secondary to, or redundant with,

other adaptor-binding motifs. Ubiquitination directs internalized

proteins toward lysosomal degradation by engagement with en-

dosomal sorting complexes required for transport (ESCRTs) (re-

viewed in Clague and Urbe, 2006). Monoubiquitination, in the

form of an irreversible linear fusion appended to various receptors,

is a sufficient signal for this sorting step. However, evidence

suggests K63 as the primary ubiquitin chain type involved in endo-

somal sorting. Early studies in yeast cells, which suggested that

appendage of K63-linked diubiquitin enhances vacuolar sorting,

have been recently elaborated on with a detailed analysis of the

downregulation of the Gap1 permease. These studies conclude

that monoubiquitination is sufficient for initial internalization (at

least so long as it is an irreversible linear fusion) but that efficient

sorting at the endosome by the ESCRT machinery requires K63-

linked polyubiquitin (Lauwers et al., 2009). Concordantly, studies

of the mammalian TrkA and MHC class I proteins reveal their utili-

zation of K63-linked polyubiquitination for routing to the lysosome

(Duncan et al., 2006; Geetha and Wooten, 2008).

The first point of engagement of ubiquitinated cargo with the

MVB sorting machinery is proposed to be the ESCRT-0 complex,

comprising HRS and STAM, both of which possess UIM and VHS

(Vps27, HRS, and STAM) domains, which can bind ubiquitin

(Figure 2). Intact ESCRT-0 binds 50 times more tightly to K63-

linked tetraubiquitin than to monoubiquitin, but only 2-fold more

tightly than to K48-tetraubiquitin (Ren and Hurley, 2010).

ESCRT-0 is recruited to endosomes through binding to phospha-

tidylinositol 3-phosphate but also binds to clathrin and the down-

stream ESCRT-I complex. An alternative ESCRT-0 complex

comprising TOM1, Tollip, and Endofin possesses all these salient

features of the HRS-STAM complex. It is currently unclear

whether these two complexes are redundant or used to receive

different cargoes. In a further striking parallel to the proteasomal

system, the ESCRT machinery has known associations with at

least two DUB activities, AMSH and USP8 (UBPY), drawn from

the JAMM/MPN+ and USP families, respectively. In yeast, the

dominant endocytic E3 ligase activity Rsp5 can also associate

with the STAM ortholog Hse1, providing a counterbalance to

Ubp2 and Ubp7 (Ren et al., 2007), while a third ESCRT-associ-

ated DUB Doa4 is required for ubiquitin recycling of receptors

that are committed to degradation. Although deubiquitination is

not an obligate step for MVB sorting, proof-reading and ubiquitin

recycling roles akin to those suggested for proteasomal DUBs

are consistent with available data (Clague and Urbe, 2006).

AutophagyThe signature of autophagy is the capture of cytosol and organ-

elles through envelopment within a double-membrane compart-

ment derived from the preautophagosomal structure. In

common with the MVB, the autophagosome can then directly

fuse with late endosomes or lysosomes to form the autolyso-

some, wherein the double-membrane structure is digested. It

is well suited for the digestion of cytosolic entities, which are

incompatible with unfolding by the proteasome, such as organ-

elles or protein aggregates.

Identification of autophagy (Atg) genes and subsequent

biochemical characterization revealed two essential posttransla-

tional modification pathways, which resemble ubiquitination.

In one case, Atg12 is stably conjugated to Atg5 in a constitutive

fashion. In the second case, Atg8 is conjugated to the lipid phos-

phatidylethanolamine by transfer from an E2 enzyme following the

onset of autophagy (for example, as induced by amino acid depra-

vation). This is a prerequisite for the expansion of the preautopha-

gosomal structure, perhaps by facilitating fusion between

membranes. Inmammalian cells,Atg8 is known as LC3and its lipi-

dated form as LC3-II. In fact, there are six Atg8 homologs in the

human genome collectively known as the LC3/GABARAP family.

Whereas autophagy is generally thought of as a nonselective

degradation process, certain structures and organelles are selec-

tively removed by this pathway. For example, mitochondria are

lost during reticulocyte maturation and as a consequence of un-

coupling (disconnecting the electron transport chain from ATP

production) in cultured cells. Ribosomes, peroxisomes, and path-

ophysiological protein aggregates can also be degraded by

autophagy. Recent studies have led to the proposal of a common

principle involved in ‘‘selective autophagies’’ and once againubiq-

uitin plays a critical role (Kirkin et al., 2009). In general if the body to

be cleared is ubiquitinated, then an adaptor molecule is required

to couple this to the preautophagosomal membrane rich in

Atg8/LC3. The prototypical adaptor of this class is p62/sequesto-

some 1, which contains both a ubiquitin-interacting domain (UBA)

and a LIR motif (LC3-interacting region), a domain structure

shared with Neighbor of BRCA1 gene 1 (NBR1) (Figure 2) (Pankiv

et al., 2007). p62 has been previously implicated in the clearing of

protein aggregates, which are known to be concentrated in ubiq-

uitin. Recent data have indicated an essential role for ubiquitin

(K63 and K27 polyubiquitin chain linkages have been implicated)

in the selective autophagy of depolarized mitochondria, which

become ubiquitinated following recruitment of the E3 ubiquitin

ligase Parkin (Geisler et al., 2010). Using a lysine-less mutant of

ubiquitin fused with red fluorescent protein, Kim et al. established

that irreversible monoubiquitination is sufficient to concentrate

a soluble protein within autophagosomal structures in a p62-

dependent manner (Kim et al., 2008).

A selective pathway requiring the Ubp3:Bre5 DUB complex in

Saccharomycescerevisiaeoperates in the removal of mature ribo-

somes (Kraft and Peter, 2008). In cells deficient in Ubp3, ribosomal

fractions are enriched with ubiquitin. Although an intimate


connection has been established, the exact role of ubiquitin in ri-

bophagy is unclear. One model posits that ubiquitin may be pro-

tecting ribosomes from autophagy, which is then promoted by

Ubp3 activity. Alternatively, a dynamic modification with ubiquitin

may be required, perhaps as an engulfment signal similar to that of

mitochondria. In support of this notion, a temperature-sensitive

defect in the E3 ligase Rsp5 shows a synthetic ribophagy defect

with loss of Ubp3 ascompared withcells lacking Ubp3alone (Kraft

and Peter, 2008). If correct, then the principle of ensuring ubiquitin

homeostasis through deubiquitination may be conserved by each

of the selective degradation pathways we have discussed.

The Interdependence of Degradation PathwaysThe relative contribution of degradation pathways may vary

greatly between cell types. In most cases of cells cultured under

stress-free conditions, proteasomal degradation predominates,

but in muscle cells, lysosomal pathways (principally autophagy)

can account for 40% of degradation of long-lived proteins. In

atrophying muscle cells, both pathways are proposed to be

co-ordinately upregulated under the transcriptional control of

FOXO3 (Zhao et al., 2007). However, the proteasome is itself

degraded by starvation-induced bulk autophagy (Kristensen

et al., 2008).

The reliance of three major cellular degradation pathways

upon ubiquitination suggests that specific inhibition of any one

pathway may perturb the ubiquitin economy of the cell and

hence indirectly affect other degradation events (Figure 1).

A clear example of this is the activated Met receptor, for which

its lysosomal degradation is exquisitely sensitive to the depletion

in free ubiquitin caused by proteasomal inhibition (Carter et al.,

2004). Proteasome inhibition may also induce autophagy as

a compensatory response. The autophagy adaptor protein p62

has also been implicated in proteasomal degradation, whereas

the E3 ligase Parkin generates an autophagy tag on mitochon-

dria but elsewhere can target proteins to the proteasome.

VCP/p97 co-ordinates a number of ubiquitin-dependent

processes that include the proteasome-dependent ERAD (endo-

plasmic reticulum-associated degradation) pathway but inter-

estingly has recently been identified as a necessary factor for

autophagosome maturation under basal conditions and

following proteasome inhibition (Tresse et al., 2010).

The MVB and autophagy pathways merge at the late endo-

some/lysosome and are both sensitive to proton pump and

phosphoinositide 3-kinase inhibitors. Autophagosome formation

is inherently sensitive to perturbations earlier in the endocytic

pathway, which change the character of later endosomal

compartments (such as the composition of SNARE proteins).

Occasionally, teleological distinctions between these systems

blur, such that some ubiquitinated cytosolic proteins may be

degraded in the lysosome and cytoplasm-exposed domains of

receptors may be nibbled by the proteasome. Mounting

evidence suggests that there is a proteasome pool associated

with endosomes that influences receptor sorting (Geetha and

Wooten, 2008; Gorbea et al., 2010).

Concluding RemarksUbiquitin tagging is common to the three major cellular pathways

for protein degradation. Herein lies a conundrum: how is a given

substrate targeted to a particular pathway? Variable parameters

include location, chain length, and linkage type. A clear bias of

the endosomal pathway toward K63-linked chains has emerged.

This may simply reflect the subcellular localization of specific

E3 ligases in combination with a high local concentration of ubiq-

uitin-binding proteins, which couple to the ESCRT-machinery

rather than the proteasome. New techniques allow for the deter-

mination of individual protein turnover on a global scale (Kristen-

sen et al., 2008). This will enable the generation of a comprehen-

sive annotation of turnover rates as a function of experimental

perturbations or disease states, opening the door to

a systems-level understanding of protein degradation.

ACKNOWLEDGMENTS

S.U. is a Cancer Research UK Senior Research Fellow.

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Leading Edge

Perspective

Will the Ubiquitin System Furnish asMany Drug Targets as Protein Kinases?Philip Cohen1,2,* and Marianna Tcherpakov31MRC Protein Phosphorylation Unit2Scottish Institute for Cell SignallingSir James Black Centre, Dow Street, Dundee DD1 5EH, Scotland, UK3BCC Research, 40 Washington Street, Suite 110, Wellesley, MA 02481, USA

*Correspondence: [email protected]

DOI 10.1016/j.cell.2010.11.016

Protein phosphorylation and protein ubiquitination regulate most aspects of cell life, and defects inthese control mechanisms cause cancer and many other diseases. In the past decade, proteinkinases have become one of the most important classes of drug targets for the pharmaceuticalindustry. In contrast, drug discovery programs that target components of the ubiquitin systemhave lagged behind. In this Perspective, we discuss the reasons for the delay in this pipeline, thedrugs targeting the ubiquitin system that have been developed, and new approaches that maypopularize this area of drug discovery in the future.

Protein Phosphorylation Drug DiscoveryIt can take years, even decades, before a field of research rea-

ches the stage of maturity at which its discoveries can obviously

be exploited for the improvement of health. An excellent example

of this paradigm is the regulation of protein function by reversible

phosphorylation. Phosphorylation was identified in the mid

1950s as a mechanism for controlling glycogenolysis. Twenty-

five years later, it was still largely thought of simply as a control

switch for metabolism. Indeed, researchers finally realized that

protein phosphorylation regulates most aspects of cell life only

after many advances made throughout the 1980s and early

1990s (Cohen, 2002a).

Surprisingly, the idea that it would be possible to treat diseases

with drugs targeting protein kinases was even slower to take

root. Indeed, as late as 1998, the Head of Research and Develop-

ment at one major pharmaceutical company (which no longer

exists) told one of the authors that ‘‘there was absolutely no

future in kinase drug discovery.’’ Later that same year,

researchers revealed the remarkable clinical efficacy of a tyrosine

kinase inhibitor, called Gleevec, for treating chronic myeloge-

nous leukemia. Quite quickly, protein kinases then became one

of the most popular classes of drug targets for the pharmaceu-

tical industry, especially in the field of cancer treatment.

Over the past decade, 16 drugs targeting one or more protein

kinases have been approved for clinical use in cancer, 12 taken

orally as pills and 4 that are injected. As of 2009, 153 other

protein kinase inhibitors were undergoing clinical trials, and 23

of these drugs were in the most advanced stage of development,

termed Phase III (Table 1) (Lawler, 2009). The current global

market for kinase therapies is about US$15 billion per annum,

and this value is forecasted to double by 2020. Research on

protein kinases currently accounts for �30% of the drug

discovery programs in the pharmaceutical industry and over

50% of cancer research and development. The kinase inhibitors

undergoing Phase III clinical trials include Pfizer’s JAK3 inhibitor

for rheumatoid arthritis (CP-690550) and Incyte Pharmaceuti-

cal’s JAK1/JAK2 inhibitor (INCB18424) for treating inflammatory

diseases. If these drugs are approved, it will likely spark a new

wave of interest in developing kinase inhibitors for the treatment

of diseases other than cancer.

Even by the late 1970s and early 1980s researchers had shown

that oncogenes, such as Src (sarcoma), are protein kinases;

phorbol esters, which promote tumors, are kinase activators;

and, growth factor receptors, which have kinase domains, are

overexpressed or mutated in human cancer (reviewed in Cohen,

2002b). So why did it take so long for most pharmaceutical

companies to capitalize on the therapeutic potential of kinase

inhibitors? In retrospect, one realizes that many researchers

believed that kinase inhibitors were bad drug targets because

they thought that it would be difficult to achieve the requisite

specificity and potency. Most protein kinase inhibitors target

the ATP-binding pockets of these enzymes, and the structural

similarities of this site among many different kinases raised the

suspicion that it would be impossible to develop drugs that in-

hibited only one type of protein kinase. Furthermore, the concen-

tration of ATP in the cell is extremely high (i.e., millimolar), leading

researchers to doubt whether compounds could be developed

with the potency needed to compete successfully with intracel-

lular ATP. These were, and indeed still are, challenging problems

for many developing kinase inhibitors, but they have proven to be

quite surmountable.

Indeed, considerable potency and specificity have been

achieved by developing compounds that target not only the

ATP-binding site but also small hydrophobic pockets located

proximal to the ATP-binding site. Moreover, researchers are

identifying an increasing number of ‘‘allosteric’’ inhibitors that

bind to other regions of a kinase. These compounds induce

conformational changes in the kinase, which either suppress


the enzyme’s activity directly or block its activation by another

kinase in the same signaling cascade.

Furthermore, far from being a disadvantage, lack of specificity

can actually be an advantage. For example, Gleevec was devel-

oped as an Abelson kinase inhibitor for the treatment of a specific

type of leukemia. However, it is also an effective treatment for

gastrointestinal stromal cancers because it inhibits the c-Kit

receptor and the platelet-derived growth factor (PDGF) receptor

tyrosine kinases, which are overexpressed or mutated in gastro-

intestinal cancers (Demetri et al., 2006). In addition, the efficacy

of several anticancer drugs depends on their combined inhibition

of several different kinases, and these drugs may be less prone

to the development of drug resistance than ones that act on only

one specific kinase. Thus, some of the original prejudices against

protein kinases as drug targets have subsequently turned out to

have little substance.

The beauty of targeting protein kinases for therapeutics and

the basis for their popularity is that the same technologies and

small-molecule libraries can be used to develop inhibitors of

many types of protein kinases in almost every therapeutic

area. However, the vast amount of medicinal chemistry that

has been carried out in recent years has meant that novel patent

space is becoming quite difficult to find. Plus, there is a growing,

but probably unfounded, concern that the most important drug

targets in this area have been fully exploited. Therefore, the phar-

maceutical industry has begun to wonder where they may find

the next large set of drug targets that can be tackled in a manner

analogous to protein kinases. In this Perspective, we discuss the

premise that components of the ubiquitin system are prime

candidates for these new targets.

Ubiquitination More Versatile than Phosphorylation?Ubiquitination is the covalent attachment of a small protein,

ubiquitin (�8.5 kDa), to other proteins. In the first step, a thioester

bond is formed between the C-terminal carboxylate group of

ubiquitin and the thiol or sulfhydryl group of a cysteine residue

on an E1-activating enzyme. Next, the ubiquitin is transferred

to a cysteine on an E2-conjugating enzyme. In the third step,

the E2 interacts with an E3 ligase, and the ubiquitin is then trans-

ferred from the E2 enzyme to substrates, which also interact with

the E3 ligase. This last step can occur directly, as in the RING E3

ligases, or it can occur indirectly with the ubiquitin first trans-

ferred to a cysteine residue on the E3 ligase before being linked

to the substrate, as in the HECT family of E3 ligases. Chains of

ubiquitin are created by the same enzymatic process.

Similar to phosphorylation, ubiquitin can be linked covalently

to only one or several amino acid residues on the same protein

(Figure 1). However, in contrast to protein phosphorylation, ubiq-

uitin can also form polyubiquitin chains. Ubiquitin has seven

lysine residues and an a-amino group; thus eight different types

of polyubiquitin chains can form (and probably more because

chains with ‘‘mixed’’ linkages are also present in cells).

Even greater versatility is provided by ubiquitin-like proteins,

such as Nedd8, SUMO (1, 2, and 3), FAT10, and ISG15, which

are also attached covalently to proteins in processes called ned-

dylation, SUMOylation, tenylation, and ISGylation, respectively.

The formation of polyubiquitin chains and the existence of these

‘‘ubiquitin-like modifiers’’ make the ubiquitin system a more

complex and potentially more versatile control mechanism

than phosphorylation.

Like phosphorylation, ubiquitination is reversible. Isopepti-

dases, called deubiquitinases or DUBs, catalyze the cleavage

of the ubiquitin from proteins or ‘‘deubiquitination’’ (Figure 1).

Interestingly, the number of deubiquitinases is comparable to

the number of protein phosphatases, but taken together, the

number of E1-activating enzymes, E2-conjugating enzymes,

and E3 ligases encoded by the human genome exceeds the

number of protein kinases.

Ubiquitination and Phosphorylation: Analogous ControlMechanismsFor many years, the sole function of the ubiquitin system was

thought to be the regulation of protein turnover inside the cell. At-

taching a chain of ubiquitins linked at lysine 48 (K48-linked poly-

ubiquitination) to a protein directs it to the 26S proteasome for

destruction, and indeed, this is one of the key functions of the

Table 1. Phosphorylation, Ubiquitination, and Drug Discovery

Phosphorylation Ubiquitination

First publication 1955a First publication 1978b

>500 protein kinasesc 10 E1sf, �40 E2sf, >600 E3 ligasesf

140 protein phosphatasesc �90 deubiquitinasesc

Nobel Prize awarded 1992d Nobel Prize awarded 2004e

First drug approved in 2001 (Gleevec) First drug approved in 2003 (Bortezomib)

16 drugs approved, over 150 undergoing clinical trials One drug approved, 16 undergoing clinical trials

Current sales �US$15 billion per year Current sales �US$1.4 billion per year

�30% of pharmaceutical research and development <1% of pharmaceutical research and developmenta Fischer and Krebs, 1955.b Ciechanover et al., 1978.c Encoded by the human genome.d Nobel Prize for Physiology or Medicine awarded to Edmond Fischer and Edwin Krebs.e Nobel Prize for Chemistry awarded to Aaron Ciechanover, Avram Hershko, and Irwin Rose.f Includes the E1s and E2s for ubiquitin-related modifiers such as Nedd8, SUMO, FAT10, and ISG15.


ubiquitin system. However, other types of ubiquitination play

distinct roles in the cell and regulate diverse areas of biology,

as discussed in another article in this issue (Ikeda et al., 2010).

For example, K63-linked polyubiquitination (Bhoj and Chen,

2009; Zeng et al., 2010) and linear polyubiquitin chains

(Tokunaga et al., 2009) regulate innate immunity; K11-linked pol-

yubiquitin chains, which are formed by the anaphase-promoting

complex (APC/C) and the E2-conjugating enzyme UbcH10,

are critical for the regulation of mitosis (Garnett et al., 2009;

Jin et al., 2008); and K29/33-linked polyubiquitination inhibits

certain members of a protein kinase subfamily (Al-Hakim et al.,

2008).

Like phosphorylation, ubiquitination can also induce confor-

mational changes that alter biological function. For example,

the response to the proinflammatory cytokine interleukin-1 (IL-

1) generates K63-linked polyubiquitin chains that interact with

a component of the TAK1 complex, inducing a conformational

change that allows this protein kinase to autoactivate (Xia

et al., 2009). Similarly, monoubiquitination of the deubiquitinase

Ataxin 3 (Todi et al., 2009) and dihydrofolate reductase (Maguire

et al., 2008) enhances and suppresses their enzymatic activities,

respectively. In contrast, monoubiquitination of the tumor

suppressor p53 induces a conformational change that exposes

a nuclear export signal. This leads to the translocation of p53

to the cytosol where it may promote apoptotic events (Carter

et al., 2007). Neddylation of the Cullin RING E3 ligases (CRLs)

also induces conformational changes that bring the E2 active

site adjacent to the substrate, permitting the efficient ubiquitina-

tion of the substrate by CRLs (Saha and Deshaies, 2008).

Like phosphorylation, many effects of ubiquitination are medi-

ated by interactions with ubiquitin-binding proteins. Different

polyubiquitin chains adopt distinct three-dimensional structures

and hence interact with different polyubiquitin-binding proteins

to regulate distinct processes. For example, proteins tagged

with K48-linked polyubiquitin chains are targeted for destruction

because these ubiquitin chains bind to particular components of

the 26S proteasome. More than 20 different families of polyubi-

quitin-binding proteins have been identified, and this area has

become a large topic of research in its own right.

Interactions through ubiquitin are also critical for DNA-damage

signaling and for certain DNA-repair pathways. For example, the

monoubiquitinated form of FANCD2, a component of the Fanconi

Anemia Complex, interacts with the UBZ domain of the DNA

nuclease FAN1, and this interaction through ubiquitin is essential

for repair of DNA interstrand crosslinks (MacKay et al., 2010).

K63-linked polyubiquitin chains attached to histone 2A and

histone 2AX by the E3 ligase RNF8 and the E2 -conjugating

enzyme Ubc13 (Kolas et al., 2007) recruit and assemble factors

that are essential for DNA repair, such as BRCA1 (breast cancer

1), RAP80, and other proteins (Bennett and Harper, 2008).

It is important to emphasize that protein phosphorylation and

protein ubiquitination are not distinct and separate control mech-

anisms because the interplay between them is critical for the

regulation of many cellular processes. For example, phosphoryla-

tion regulates a number of E3 ubiquitin ligases and deubiquiti-

nases. Further, the E3 ligase Skp1-Cullin-F box (SCF) and some

other E3 ligases contain an additional component bTRCP

(b-transducin repeat-containing protein), which recognizes partic-

ular phosphorylated sequence motifs that direct the SCFbTRCP

complex to ubiquitinate these substrates. Finally, a number of

kinases can be activated or inhibited by interactions with polyubi-

quitin chains or by polyubiquitination. Given the omnipresence of

protein phosphorylation and ubiquitination inside the cell, under-

standing the interplay between these two systems is likely to

become increasingly more important over the next decade.

Developing Drugs that Target the Ubiquitin SystemThe Proteasome Inhibitor Bortezomib

The protease inhibitor Bortezomib, originally called PS341 and

then Velcade (Adams, 2002), was the first drug that targets

a component of the ubiquitin system to be approved for clinical

use in the United States. Developed by ProScript Inc in 1995,

Bortezomib entered clinical trials in 1997 and was approved by

the Federal Drug Administration in 2003. In 1999 ProScript was

acquired by Leukosite, which in turn was acquired by Millenium

Pharmaceuticals later that same year. Bortezomib has been

quite successful, with worldwide sales in 2009 of US$1.4 billion,

and this achievement led Takeda to acquire Millenium in 2008.

Bortezomib was approved as a front-line treatment for B cell

lymphoma found primarily in the bone marrow. It is also used

for the treatment of mantle cell lymphoma in patients who have

already received other treatments. It is in Phase III clinical trials

for follicular non-Hodgkin’s lymphoma, Phase II trials for diffuse

large B cell lymphoma, and a great many other clinical trials (re-

viewed in Tcherpakov, 2010).

Bortezomib, which is given by intravenous injection, has

remarkable efficacy against multiple myeloma, but the molecular

mechanism underlying its effect is still unclear. Nevertheless, the

multiple myeloma cells that are particularly sensitive to protea-

some inhibitors express lower levels of proteasome particles

Figure 1. Phosphorylation and Ubiquitina-

tion Regulate Most Aspects of Cell LifePhosphorylation involves the covalent attachmentof phosphate to proteins, mainly to serine, threo-nine, and tyrosine residues. Phosphorylation iscatalyzed by protein kinases and reversed byprotein phosphatases. Protein ubiquitinationinvolves the covalent attachment of ubiquitin,a small protein with 76 amino acids, to otherproteins, predominantly to lysine residues. Thisreaction is mediated by an E1-activating enzyme,an E2-conjugating enzyme, and an E3 ligase; thisreaction is reversed by deubiquitinases.


and have a higher proteasome workload than multiple myeloma

cells that are relatively resistant to these drugs. Thus, the balance

between proteasome workload and degradative capacity may be

an important determinant of the sensitivity of a cancer cell to

Bortezomib and other proteasome inhibitors (Bianchi et al.,

2009).

A dipeptidyl boronic acid, Bortezomib binds noncovalently to

the 20S proteasome and primarily inhibits its chymotrypsin-like

activity (Kisselev et al., 2006). Its success has led to considerable

interest in developing improved ‘‘second generation’’ inhibitors,

and Millenium/Takeda has another proteasome inhibitor,

MLN9708, which can be taken orally, in Phase 1 clinical trials.

Onyx Pharmaceuticals also has several orally active proteasome

inhibitors in clinical trials, which they obtained through the acqui-

sition of Proteolix. These inhibitors include Carfilzomib, which

has recently entered Phase III trials according to the website

http://clinicaltrials.gov. Other proteasome inhibitors that are

currently undergoing clinical development are listed in Table 2.

An Inhibitor of the E1 Enzyme for Neddylation

The Nedd8 protein shares �60% sequence identity with ubiqui-

tin, and it is conjugated to its target proteins in a similar manner

to ubiquitin, with a specific E1-activating enzyme (NAE-E1) and

the E2-conjugating enzymes Ube2M and/or Ube2F. The primary

target for neddylation appears to be the Cullin components of

Cullin RING E3 ubiquitin ligases. The Cullin RING ligases are

the largest family of E3 ligases in the human genome with more

than 100 members (Rabut and Peter, 2008). Neddylation permits

efficient ubiquitination by Cullin RING ligases; neddylation

induces a conformational change in the Cullin component to

bring the E2 active site adjacent to the lysine residue of its protein

target substrates (Duda et al., 2008; Saha and Deshaies, 2008).

Millenium/Takeda has developed a relatively specific inhibitor

of the NAE-E1 enzyme (Table 3). This compound, MLN4924,

showed promise in mouse models of cancer and has entered

Phase I clinical trials for the treatment of multiple myeloma and

non-Hodgkin’s lymphoma. MLN4924 seems to exert its effect

on these cancers by deregulating DNA synthesis during the S

phase of the cell division cycle. MLN4924 appears to stabilize

Cdt1, a DNA replication licensing factor normally ubiquitinated

by a Cullin RING E3 ligase and then degraded by the proteasome

(Soucy et al., 2009).

Inhibitors of Deubiquitinases

Deubiquitinases comprise five separate gene families. Four

families are cysteine proteinases (the USP, OTU, UCH, and

MJD deubiquitinases), and the other one consists of metallo-

proteinases (the JAMM/MPN domain family). The E3 ligase

HDM2 targets the tumor suppressor p53 for degradation. One

of the cysteine protease deubiquitinases, USP7 (ubiquitin-spe-

cific protease 7), deubiquitinates HDM2, leading to increased

levels of HDM2 and decreased levels of p53. Therefore, two

companies, Progenra and Hybrigenics, have developed inhibi-

tors of USP7 (i.e., P5091 and HBX 41108, respectively) (Colland

et al., 2009), with the hope of promoting the proteasomal degra-

dation of HDM2 by enhancing its polyubiquitination. Reduced

expression of HDM2 would then be expected to increase the

level of p53.

Progenra is also developing inhibitors targeting USP20, and

they are showing interest in agents for USP2a, USP33, and

Table 2. Proteasome Inhibitors Approved or in Clinical Trials

Company Inhibitor Development Stage Disease

Millenium/Takeda Bortezomib/Velcade Approved Multiple myeloma and mantle cell lymphoma

Millenium/Takeda MLN9708 Phase I Multiple myeloma and other cancers

ONYX (Proteolix) Carfilzomib/PR171 Phase III Multiple myeloma and other cancers

ONYX (Proteolix) Onx 0912/PR047 Phase I Multiple myeloma and other cancers

Cephalon CEP18770 Phase I Multiple myeloma and other cancers

Nereus Pharmaceuticals Salinosporamid A/NPI0052 Phase I Multiple myeloma and leukemia

Table 3. Inhibitors of E1-Activating Enzymes and E3 Ubiquitin Ligases Undergoing Clinical Trials

Company Inhibitor Target Stage Disease

Millenium/Takeda MLN4924 NAE-E1b Phase II Multiple myeloma and Hodgkin’s lymphoma

Roche Nutlin/R7112 E3-Hdm2 Phase I Blood cancers

and solid tumors

Johnson & Johnson JNJ26854165 E3-Hdm2 Phase I Multiple myeloma and solid tumors

Genentech/Roche GDC-0152 E3-IAP Phase I Metastatic malignancies

Novartis LCL161 E3-IAP Phase I Solid tumors

Ascenta Therapeutics AT-406 E3-IAP Phase I Solid tumors and lymphoma

Aegera Therapeutics AEG 35156a E3-IAP Phase II AML and liver cancer

Aegera Therapeutics AEG 40826 E3-IAP Phase I Lymphoid tumors

Tetralogics Pharma TL 32711 E3-IAP Phase I Solid tumors and lymphoma

Astellas Pharma YM155 E3-IAP Phase II Lung cancera Antisense oligonucleotide.b The E1-activating enzyme for neddylation.


AMSH (associated molecule with the SH3 domain of STAM)

(http://www.progenra.com/scientist.html, 2009). USP20, also

called VDU2 (von Hippel-Lindau deubiquitinating enzyme 2),

deubiquitinates and stabilizes hypoxia-inducible factor 1a (HIF-

1a) (Li et al., 2005). HIF-1a is expressed at high levels in many

human cancers because it is stabilized at the low concentration

of dissolved oxygen inside the tumor by high cytokine levels and

by specific genetic alterations. For example, in von Hippel-Lin-

dau disease, in which individuals develop a variety of tumors,

mutations in the VHL gene compromise the ubiquitination and

degradation of HIF-1a, leading to the accumulation and overex-

pression of HIF-1a and its target genes. Therefore, inhibitors of

USP20 (VDU2) and/or USP33 (VDU1) may reduce levels of HIF-

1a by enhancing its polyubiquitination.

Novartis has patented compounds that inhibit the deubiquiti-

nases USP2 and UCH-L3 (ubiquitin C-terminal hydrolase).

USP2 is another deubiquitinase reported to target MDM2, the

mouse ortholog of HDM2 (Stevenson et al., 2007), whereas

UCH-L3 probably plays a role in neurodegenerative disorders,

such as Parkinson’s disease. Recently, researchers identified

a small-molecule inhibitor of USP14, called IU1, which did not

inhibit eight other deubiquitylases tested, demonstrating the

feasibility of developing relatively specific inhibitors of these

enzymes (Lee et al., 2010). USP14 is associated with the protea-

some, and treating cells with IU1 enhanced the degradation of

several proteasomal substrates that have been implicated in

neurodegenerative diseases, such as Tau. Drugs that target

USP14 could, therefore, have a potential use in reducing or elim-

inating misfolded and aggregated proteins that accumulate in

neurodegenerative and other diseases.

Developing pharmaceutical agents that target deubiquitinases

is still in its infancy, and to our knowledge, no deubiquitinase

inhibitor has yet entered clinical trials. However, as this field

progresses, it is clearly going to be essential to assess the

specificities of these inhibitors. Therefore, assembling compre-

hensive panels of deubiquitinases for testing specificity will be

critical, similar to how large panels of protein kinases have

been of immense value in assessing the selectivity of kinase

inhibitors.

As with kinases, there are certainly going to be deubiquitinases

for which inhibition needs to be avoided. For example, mutating

or deleting the A20 deubiquitinase causes or predisposes indi-

viduals to inflammatory and autoimmune diseases (Musone

et al., 2008; Turer et al., 2008). Similarly, inactivating mutations

in the deubiquitinase CYLD cause cylindromatosis, a type of

skin cancer (Kovalenko et al., 2003; Trompouki et al., 2003).

Targeting E3 Ubiquitin Ligases

The human genome encodes more E3 ubiquitin ligases than

protein kinases (Table 1). Furthermore, the E3 ligase confers

specificity to ubiquitination when it transfers ubiquitin from an

E2 to a particular substrate. For these reasons, E3 ubiquitin

ligases are attractive candidates as drug targets. In some cases,

identifying compounds that disrupt the interaction of an E3 ligase

with its substrates has proven a frustrating experience for

several companies, and a number of programs have been

unsuccessful. For example, we understand that several compa-

nies have tried and failed to develop inhibitors of MuRF1, an E3

ligase involved in degrading myosin as a therapy for preventing

muscle wasting. Nevertheless, several programs have made

good progress and a number of E3 ligase inhibitors have

advanced to clinical trials (Table 3) (reviewed in Tcherpakov,

2010). Moreover, several recent and unexpected developments

in this area are likely to enhance future pharmaceutical interest in

developing E3 ligase inhibitors.

Several companies have discovered compounds that disrupt

the interaction of the E3 ligase HDM2 and its substrate, the tumor

suppressor p53, with the aim of elevating p53 expression. One

such compound, Nutlin 3/R7112, has entered clinical trials

(Table 3). A second class of E3 ligases actively targeted by

a number of companies is the Inhibitors of Apoptosis Proteins

(IAPs), and seven antagonists of IAPs have even entered clinical

trials (Table 3). These drugs are small-molecule mimetics of

Smac (also known as Diablo), a protein that antagonizes IAPs

by interacting with their BIR domains. Smac mimetics appear

to induce the autoubiquitination and degradation of the IAPs,

which then leads to the death of cancer cells by stimulating the

TNF-a pathway (Wu et al., 2007). Destruction of IAPs through

the Smac mimetics also suppresses the production of proinflam-

matory cytokines by Toll-like receptor agonists, suggesting that

these drugs may be worth exploring as possible treatments for

chronic inflammatory diseases (Tseng et al., 2010).

Recently, Ito et al. (2010) surprisingly discovered that the drug

thalidomide binds to cereblon (CRBN), a component of the Cullin

RING E3 ligase that is important for limb outgrowth and the

expression of a fibroblast growth factor (FGF8) during embryonic

development (Ito et al., 2010). This finding explained why thalid-

omide, originally prescribed as a sedative, caused multiple birth

defects in pregnant women. Thalidomide is still used for the

treatment of numerous conditions, including leprosy, skin sores,

and myelofibrosis. Therefore, pinpointing the molecular mecha-

nism of the drug’s devastating side effects may facilitate the

development of new thalidomide derivatives that are free from

this problem.

Arsenic is another drug that unexpectedly regulates an E3

ligase. Arsenic is an effective and specific treatment for acute pro-

myelocytic leukemia. In this cancer, the promyelocytic leukemia

(PML) protein becomes fused to the retinoic acid receptor (RAR).

Arsenic triggers the degradation of the PML-RAR fusion protein

by inducing the SUMOylation of PML. This modified version of

PML recruits the SUMO-binding E3 ubiquitin ligase RNF4, which

catalyzes the polyubiquitination (K48-linked) and proteasomal

degradation of the PML-RAR complex (Tatham et al., 2008).

Small-molecule inhibitors of several Cullin RING E3 ligases

have also been identified. SCFskp2 is a Cullin RING E3 ligase

that is highly expressed in some human cancers. Decreased

levels of p27kip1 are a poor prognosis factor in many malignan-

cies, and SCFskp2 ubiquitinates p27kip1, targeting it for protea-

somal destruction (Cardozo and Pagano, 2007; Merlet et al.,

2009). Researchers have identified one compound that prevents

the incorporation of Skp2 into the SCFskp2 complex, which trig-

gers cell death (i.e., autophagy) by stabilizing p27kip1 and

inducing G1/S cell-cycle arrest. This inhibitor synergizes with

Bortezomib and overcomes resistance to Bortezomib in models

of multiple myeloma. Moreover, the compound was active

against aggressive leukemia cells (i.e., leukemia blasts) and

plasma cells derived from patients (Chen et al., 2008).


SCFbTrCP1 isa CullinRINGE3 ligase that triggers the degradation

of IkBa, the inhibitory component of the proinflammatory tran-

scription factor NF-kB. Therefore, drugs that target SCFbTrCP1

may have potential as anti-inflammatory agents, and it is of great

interest that an inhibitor of SCFbTrCP1 has been identified, which

prevents the polyubiquitination and degradation of IkBa (Nakajima

et al., 2008).

Researchers have also identified a small-molecule inhibitor of

Cdc4, the yeast ortholog of the mammalian Cullin RING E3 ligase

Fbw7 (F box and WD repeat domain-containing 7). A recent

X-ray crystal structure (Orlicky et al., 2010) revealed that the

inhibitor inserts between two of the b strands of the WD40

propeller domain of Cdc4, which are remote from the

substrate-binding site. Binding of the inhibitor induces a long-

range conformational change that distorts the substrate-binding

pocket and impedes recognition of the substrate. Thus, this

compound is one of the first allosteric inhibitors of an E3 ligase

to be identified and raises the possibility that other Cullin RING

E3 ligases with WD40 domains may possess analogous pockets

that could be targeted by inhibitors. A small-molecule inhibitor of

the SCFMet30 ligase was recently identified in a screen for small-

molecule enhancers of the drug rapamycin (Aghajan et al., 2010).

To our knowledge, none of these compounds has yet entered

clinical development, but they are proof-of-principle, demon-

strating that there is no particular fundamental barrier to identi-

fying inhibitors of the Cullin RING family of E3 ubiquitin ligases.

The Future of Ubiquitin Drug DiscoveryThere are striking parallels between the histories of protein phos-

phorylation and protein ubiquitination and their exploitation for

the development of drugs to treat diseases (Table 1). Both bio-

logical control mechanisms were identified many years ago,

but interest in targeting them for drug discovery only started to

take off in the 1990s. Indeed, the first compounds inhibiting

components of these systems entered clinical trials at around

the same time (Bortezomib—1997, Gleevec—1998), and these

drugs were among the fastest ever approved for clinical use

(Gleevec—2001, Bortezomib—2003). Both Gleevec and Borte-

zomib subsequently achieved ‘‘blockbuster’’ status with current

sales of about US$3 billion (Gleevec) and US$1.4 billion (Borte-

zomib) per annum.

However, that is where their similarities end. Since the devel-

opment of Gleevec, 15 other drugs targeting a specific protein

kinase have been approved for clinical use, but no other drug tar-

geting a particular component of the ubiquitin system has yet

been approved. In addition, kinase inhibitors currently under-

going clinical trials also outnumber the inhibitors of the ubiquitin

system by more than ten to one (Table 1).

Why has drug discovery in the ubiquitin system lagged so far

behind that of protein kinases, and what is needed to change

this state of affairs in the future? In retrospect, one factor driving

the kinase field forward at such a rapid pace is the ease with

which large and varied chemical libraries can be synthesized

and exploited to develop inhibitors of many protein kinases.

Further, receptor tyrosine kinases have extracellular domains

that can also be targeted with therapeutic antibodies. In

contrast, although E3 ubiquitin ligases outnumber protein

kinases, researchers still have not developed a general approach

for identifying inhibitors of many E3 ubiquitin ligases. This is

because, thus far, researchers have focused primarily on dis-

rupting the interaction between E3 ligases and their substrates,

which is specific to particular E3 ligase-substrate pairs. More-

over, finding compounds to disrupt the interface of two proteins

can be intrinsically more difficult to achieve than searching for

small molecules that block catalytic activity.

Surprisingly, little effort has been devoted to developing

compounds that disrupt the interactions between E2-conjugating

enzymes and E3 ligases. E2-E3 interactions are usually relatively

weak (Ye and Rape, 2009) and may therefore be relatively easy

to disrupt. Moreover, compounds that disturb the interaction

between anE2-conjugatingenzyme and anE3 ligase could, inprin-

ciple, exert their effects by binding to the E2, the E3, or the E2-E3

interface, creating the potential to identify three types of inhibitors

from a single screen. There are �40 E2-conjugating enzymes en-

coded by the human genome; therefore, on average, each E2

must interact productively with �15 E3 ligases. Compounds that

disrupt E2-E3 interactions by binding specifically to the E3 ligase

could be identified by counterscreening with another E3 ligase

that also forms a productive interaction with the same E2. Indeed,

focusing efforts on large families of E3 ligases, such as the Cullin

RING ligases, may lead to the development of chemical libraries

with the capability of disrupting many E2-E3 interactions.

By analogy with kinases, perhaps the key to developing inhib-

itors of specific E2-E3 interactions is to find compounds that

bind to small hydrophobic pockets on E3 ligases located

proximal to the E2-E3 interface itself or to identify allosteric inhib-

itors that disrupt the E2-E3 interaction by inducing long-range

conformational changes. The three-dimensional structure of an

E2-ubiquitin thiol ester-E3 ligase complex has yet to be reported,

but such a structure might be extremely helpful in understanding

how E2-E3 interactions could be disrupted. To crystallize such

a complex, it might be necessary to stabilize the E2-ubiquitin

thiol ester-E3 interactions by including a small molecule that

inactivates E3 ligase function without affecting its ability to

bind to the E2-conjugating enzyme.

Another area where more effort will probably be fruitful is the

production of chemical libraries that target the different families

of deubiquitinases. Although inhibitors of a few deubiquitinases

are under development, such as Usp2a, Usp7, Usp20, and

Uch-L3, other deubiquitinases are also potentially rewarding

drug targets but seem to have attracted little attention so far.

For example, Usp6 is an oncogene with transforming activity; re-

arrangements and fusions of this deubiquitinase are found in

a number of cancers (Oliveira et al., 2006). Moreover, the possi-

bility of developing drugs that increase the expression and/or

activity of deubiquitinases also should not be ignored. For

example, the deubiquitinase BAP1 interacts with BRCA1, an

E3 ligase frequently mutated in breast cancer. BAP1 enhances

BRCA1-mediated inhibition of breast cancer cell growth and

may be a tumor suppressor gene that functions in the BRCA1

growth control pathway (Jensen et al., 1998). Thus, drugs that

enhance the activity or expression of BAP1 could have thera-

peutic potential for treating cancer.

Experience with protein kinases has taught us that

compounds developed as inhibitors of one protein kinase

commonly turn out to inhibit other protein kinases even more


potently (Bain et al., 2007) and thus can become leads in

completely different drug discovery programs. Sorafenib (also

called Nexavar), an approved drug for the treatment of renal

cell carcinoma, was originally developed as an inhibitor of

a serine/threonine kinase Raf. However, now Sorafenib is

thought to exert its therapeutic benefit by inhibiting several tyro-

sine kinases, such as the PDGF receptor (Lierman et al., 2006).

Developing chemical libraries that target deubiquitinases is likely

to yield similar surprises and likely generate drug leads for

a number of these isopeptidases.

The success ofBortezomiband the advancement of the NAE-E1

inhibitor MLN4924 into clinical trials suggest that there is vast

potential to develop more drugs targeted to general components

of the ubiquitin system. Drugs that block the same target by

distinct mechanisms can have strikingly different efficacies

because their toxicities, half-lives in vivo, and pharmaco-dynamic

properties can vary substantially. Such targets might include other

E1-activating enzymes (e.g., the E1s for ubiquitination and SU-

MOylation) and other components of the proteasome. For

example, Bortezomib predominantly targets the chymotrypsin-

like activity of the proteasome, and drugs that inhibit the cas-

pase-like and trypsin-like activities of the proteasome may be

more potent inhibitors or have different effects than Bortezomib.

The 19S component of the proteasome is another underex-

plored target. The 19S possesses ATPase activity, a polyubiqui-

tin-binding site, and deubiquitinase activities, all of which could

be targeted for drug development. Another possible target is

p97/VCP, a protein that plays a key role in eliminating misfolded

proteins by the endoplasmic reticulum-associated degradation

pathway (ERAD). Indeed a small-molecule inhibitor of the

ATPase activity of p97/VCP has been discovered that blocks

proliferation of cancer cell lines (T.-F. Chou et al., 2008, FASEB

J., abstract). Novel proteasome inhibitors might also be useful in

transplantation as a therapy for antibody- and cell-mediated acute

rejection (Everly et al., 2008). For example, Bortezomib has shown

promise in reducing graft-versus-host disease and in reconstitut-

ing the immune system in some stem cell transplant patients.

Inflammatory and autoimmune disorders may be treated with

selective inhibitors to a distinct class of proteasome, called the

immunoproteasome. Expressed in monocytes and lympho-

cytes, the immunoproteasome regulates many facets of the

immune response, in part by shaping the antigenic repertoire

presented on class I major histocompatibility complexes. The

immunoproteasome contains orthologs of the proteolytic activi-

ties associated with the ‘‘constitutive’’ 26S proteasome,

including a component with chymotryptic-like activity, called

LMP7. Recently, researchers developed a relatively selective

inhibitor of LMP7, which prevents the production of interleukin-

2 and interferon-g by activated T cells and interleukin-23 by acti-

vated monocytes. Furthermore, this inhibitor showed promise in

treating arthritis in mouse models (Muchamuel et al., 2009).

Finally, it is also worth noting that Mycobacterium tuberculosis

is the only bacterial pathogen known to have a proteasome.

Recently, one compound, oxathiazol-2-one, was identified with

preferential inhibition of the bacterial proteasome over the

human proteasome (Lin et al., 2009). Indeed, a selective inhibitor

of this mycobacterial proteasome might be useful for treating

tuberculosis.

Predicting the future is notoriously difficult. However, given

the diverse approaches and avenues that remain unexplored

in developing drugs targeted at the ubiquitin system, the

authors of this article would be surprised if ubiquitin drug

discovery was not far more important in 10 years time than it

is today. Nevertheless, only time will tell if ubiquitin drug

discovery will eventually rival in its importance that of kinase

drug discovery.

ACKNOWLEDGMENTS

We are grateful to Ron Hay, Thimo Kurz, and the reviewers of this Perspective

for making suggestions that improved the article. P.C. is a Royal Society

Research Professor, and the work of his laboratory is supported by the UK

Medical Research Council, AstraZeneca, Boehringer Ingelheim, GlaxoSmithK-

line, Merck-Serono, and Pfizer.

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Leading Edge

Review

Pathogen-Mediated PosttranslationalModifications: A Re-emerging FieldDavid Ribet1,2,3 and Pascale Cossart1,2,3,*1Institut Pasteur, Unite des Interactions Bacteries-Cellules, Departement de Biologie Cellulaire et Infection, F-75015 Paris, France2INSERM, U604, F-75015 Paris, France3INRA, USC2020, F-75015 Paris, France


DOI 10.1016/j.cell.2010.11.019

Posttranslational modifications are increasingly recognized as key strategies used by bacterial andviral pathogens to modulate host factors critical for infection. A number of recent studies illustratehow pathogens use these posttranslational modifications to target central signaling pathways inthe host cell, such as the NF-kB andMAPkinase pathways, which are essential for pathogens’ repli-cation, propagation, and evasion from host immune responses. These discoveries open newavenues for investigating the fundamental mechanisms of pathogen infection and the developmentof new therapeutics.

Posttranslational modifications (PTMs) of proteins provide highly

versatile tools and tricks used by both prokaryotic and eukary-

otic cells to regulate the activity of key proteins. PTMs include

the addition of simple chemical groups, such as a phosphate,

acetyl, methyl, or hydroxyl groups; more complex groups,

such as AMP, ADP-ribose, sugars, or lipids; and small polypep-

tides, such as ubiquitin or ubiquitin-like proteins. They also

include modifications of specific amino acid side chains (e.g.,

deamidation of glutamine residues) and the cleavage of a peptide

bond (i.e., proteolysis).

PTMs represent efficient strategies to modify activities, half-

lives, or the intracellular localization of host proteins that are

critical for infection. The first report that a pathogen could

mediate a PTM occurred 40 years ago with the discovery that

diphtheria toxin, produced by Corynebacterium diphtheriae,

ADP-ribosylates and thus inhibits the host Elongation Factor-2

(EF-2) (Collier and Cole, 1969). This modification blocks transla-

tion in the intoxicated cells and thereby leads to cell death.

Since then, a considerable number of host PTMs mediated,

induced, or counteracted by different pathogen-encoded virulence

factors have been reported (for reviews, see Ribet and Cossart,

2010; Randow and Lehner, 2009). In this Review, we discuss new

discoveries in the modulation of PTMs by pathogens. In the first

part, we focus on ubiquitin and ubiquitin-like proteins, which have

emerged as central regulating modules targeted by both viral and

bacterial pathogens. We then discuss two recently identified

PTMs catalyzed by bacterial pathogens, AMPylation and eliminyla-

tion. In the third part, we describe how pathogens hijack certain

PTMs to preferentially target specific host pathways to promote

their replication,propagation, andescape from the immunesystem.

Ubiquitin and Ubiquitin-like Modifications Targetedby PathogensUbiquitination

Ubiquitination is the covalent attachment of ubiquitin, a small

polypeptide of 76 amino acids, to a target protein. Ubiquitin is

generally linked to the lysine residue of the target protein;

however, a cysteine, serine, threonine, or N-terminal amino

group of a protein can also be modified. This conjugation

requires the successive activities of an E1-activating enzyme,

an E2-conjugating enzyme, and then an E3 ligase. Ubiquitination

is a fundamental PTM involved in many different cellular func-

tions, including the trafficking of membrane proteins, endocy-

tosis, signal transduction, DNA repair, and transcription regula-

tion. Ubiquitin itself contains seven lysines, K6, K11, K27, K29,

K33, K48, and K63. Therefore, chains of ubiquitin can be formed

by attaching additional ubiquitin molecules to a lysine residue of

the previously attached ubiquitin.

K48-linked polyubiquitin chains play a fundamental role in

protein degradation by targeting proteins to the proteasome. In

contrast, K63-linked polyubiquitin chains are involved in nonpro-

teolytic processes, such as DNA repair and vesicular trafficking.

In addition to these ‘‘homotypic’’ K48- or K63-linked chains, in

which only one type of ubiquitin linkage is involved, mixed

K11/K63-linked chains have also recently been described

(Boname et al., 2010). The discovery of these ‘‘mixed’’ chains

highlights that ubiquitin chains are probably more diverse and

complex than appreciated until now.

Ubiquitination is reversible because eukaryotic cells encode

proteases that are specific for ubiquitin. These proteases, called

deubiquitinases (DUBs), remove ubiquitin from their targets or

cleave the bond between two linked ubiquitins.

Ubiquitination constitutes an attractive target for a wide range

of pathogens because it regulates many pathways in eukaryotic

cells. Indeed, viruses and pathogenic bacteria can modulate the

ubiquitination level of host proteins by inducing their monoubi-

quitination, their polyubiquitination with K48-linked chains

(which then triggers their degradation), their polyubiquitination

with other types of ubiquitin chains, or their deubiquitination (re-

viewed in Ribet and Cossart, 2010; Randow and Lehner, 2009).

Some pathogen-encoded effectors display E3 ubiquitin ligase

activities. An important fraction of these viral or bacterial E3


ligases shares structural homologies with eukaryotic E3 ligases,

which are classically divided into HECT and RING E3s depend-

ing on their structures and mechanistic properties (reviewed in

Kerscher et al., 2006). HECT E3 ligases transiently bind ubiquitin

before transferring it to the target protein. In contrast, RING E3

ligases do not link ubiquitin directly but rather facilitate ubiquiti-

nation by binding simultaneously to the charged E2 enzyme and

the protein target.

Recent studies have identified a new family of bacterial E3

ligases with a structural domain completely distinct from the

eukaryotic RING and HECT domains (Hicks and Galan, 2010).

Studies have also identified viral E3 ligases structurally distinct

from eukaryotic ones (Randow and Lehner, 2009). Whether

these new E3 ligases also exist in eukaryotes is still unknown.

Whereas pathogens may have acquired eukaryotic-like E3

ligases by horizontal transfer from diverse eukaryotic sources,

the noneukaryotic E3 ligases may represent novel structures

evolved by pathogens to mimic the function of these essential

enzymes of the host cell.

In addition to encoding their own E3 ligases, some pathogens

may encode adaptor proteins that bind host E3 enzymes and

redirect them to specific targets. For example, two decades

ago, a study found that this strategy is used by some human

papillomaviruses (HPVs), which are associated with the develop-

ment of uterine cervix cancer. The E6 oncoproteins of HPV sero-

type 16 and 18 recruit a host E3 ligase to induce the degradation

of the p53 tumor suppressor, thereby facilitating transformation

of the infected cells (Scheffner et al., 1990).

In addition to E3 ubiquitin ligases, pathogens also encode

DUB-like proteins. A few viral DUBs have been identified, but

their roles in vivo, as well as their host targets, are unknown.

In contrast, several DUB-like proteins have been characterized

in pathogenic bacteria. Salmonella enterica serovar Typhimu-

Figure 1. Posttranslational Modification of

Host Proteins during InfectionYersinia (blue) is an extracellular pathogen thatinjects effectors into the host cell’s cytoplasmusing a specialized type III secretion system(T3SS). Salmonella (red) triggers its own entryinto host cells and replicates in a remodeledvacuole. It also secretes T3SS-dependent effec-tors. After cell invasion, Listeria (green) escapesfrom vacuoles and resides free in the cytoplasm,where it replicates and starts moving using thehost cell’s actin. Interactions with host factorsare mediated by bacterial surface or secretedproteins. Effectors from all three of these bacteria(blue for Yersinia effectors, red for Salmonellaeffectors, and green for Listeria effectors) alterposttranslational modifications of host proteins(purple) to facilitate pathogens’ replication, propa-gation, and evasion from host immune responses .

rium (S. Typhimurium) is an invasive path-

ogen of the small intestine that, in mice,

causes a disease similar to human

typhoid fever. SseL, an effector secreted

by this bacterium, displays deubiquitinat-

ing activity in vitro. It suppresses ubiquiti-

nation and degradation of IkBa, a central

regulator of the NF-kB pathway (see below) (Figure 1) (Le Ne-

grate et al., 2008). Infection with a strain of S. Typhimurium lack-

ing sseL leads to the accumulation of ubiquitinated proteins at

the site of replicating intracellular bacteria (Rytkonen

et al., 2007). Strikingly, the decoration of intracytosolic bacteria

with polyubiquitinated proteins has recently been proposed as

a signal used by host cells to sense intracellular invaders

(Figure 1). This signal triggers cytosolic defense pathways,

such as autophagy, although the nature of ubiquitinated proteins

is unknown (Perrin et al., 2004; Thurston et al., 2009). Bacterial

DUBs may decrease this accumulation of polyubiquitinated

proteins and thus might represent a strategy developed by intra-

cellular bacteria to escape these specific host defense systems.

Interestingly, pathogen-encoded proteins can also be directly

ubiquitinated by the host cell machinery. A striking example in

which PTMs by the host cell strongly alter the behavior of bacte-

rial effectors is the Salmonella SopE and SptP proteins. These

two effectors contribute to the transient remodeling of the host

cell’s cytoskeleton during bacterial entry into the cell. SopE

acts as a GEF (guanine nucleotide exchange factor) and

activates host Rho-GTPases, resulting in actin cytoskeleton

rearrangement, membrane ruffling, and subsequent bacterial

uptake. In contrast, SptP acts as a GAP (GTPase-activating

protein) to deactivate Rho-GTPases and allow the recovery of

the actin cytoskeleton’s normal architecture a few hours after

infection. Although SopE and SptP are codelivered by Salmo-

nella, they exhibit different half-lives. SopE is rapidly polyubiqui-

tinated and degraded by the host proteasome, whereas SptP

exhibits much slower degradation kinetics (Kubori and Galan,

2003). Recent studies found that Salmonella also hijacks the

ubiquitination machinery to control one of its effectors, SopB,

which displays two different activities depending on whether

the protein is ubiquitinated or not (Patel et al., 2009; Knodler


et al., 2009). Thus, by actively co-opting the ubiquitination

machinery of the host cell, Salmonella regulates the half-lives

and activities of some of its key virulence factors.

SUMOylation

In addition to ubiquitin, other polypeptides can be covalently

linked to cellular proteins to modify their fate and functions.

These polypeptides, which belong to the ubiquitin-like protein

family, share high structural homology with ubiquitin, ranging

from �15% to 50% sequence similarity with it. SUMO (small

ubiquitin-like modifier) belongs to the ubiquitin-like protein family

and is ubiquitous in the eukaryotic kingdom. The human genome

encodes three functional SUMO isoforms that can be linked to

hundreds of different targets. Similar to the ubiquitin system,

the conjugation of SUMO onto the lysine of a target protein

requires an E1, an E2, and an E3 SUMO enzyme. In parallel,

deSUMOylases regulate the SUMOylation level of cellular

proteins by removing SUMO from its targets.

SUMOylation is a fundamental PTM involved in transcription

regulation, intracellular transport, stress responses, the mainte-

nance of genome integrity, and many other biological processes.

Although SUMOylation was first thought not to play a role in

protein degradation, recent findings show that SUMO can trigger

the recruitment of ubiquitin E3 ligases, such as RNF4 (RING

finger protein 4), leading to the ubiquitination and proteasomal

degradation of some SUMOylated proteins (Lallemand-Breiten-

bach et al., 2008; Tatham et al., 2008).

As with the ubiquitin system, several bacterial and viral factors

target or mimic components of the SUMOylation machinery,

thereby increasing or decreasing the SUMOylation level of host

proteins (reviewed in Boggio and Chiocca, 2006; Ribet and

Cossart, 2010). For example, KSHV (Kaposi’s sarcoma-associ-

ated herpes virus), a herpes virus responsible for Kaposi’s

sarcoma development, encodes an enzyme, K-bZip, which

displays E3 SUMO ligase activity. This protein directly partici-

pates in catalyzing SUMO conjugation to host targets, such as

p53 and Retinoblastoma (Rb) protein (Chang et al., 2010). These

modifications are proposed to play a role in modulating host

genes expression in the early stage of viral infection (Chang

et al., 2010).

VP35, a protein encoded by Ebola virus, does not display

E3-like activity, but it binds to the host E3 SUMO enzyme

PIAS1 (protein inhibitor of activated STAT 1) and increases the

SUMOylation level of IRF7 (interferon regulatory factor 7) (Chang

et al., 2009). This SUMOylation of IRF7 downregulates interferon

transcription and may contribute to the dampening of the anti-

viral response induced upon infection of Ebola virus (Chang

et al., 2009).

Gam1, a protein encoded by an avian adenovirus, has an

opposite effect on SUMOylation; it targets the host E1 SUMO

enzyme to proteasomal degradation, thereby inhibiting the

SUMOylation machinery and altering host transcription (Boggio

et al., 2004). Degradation of the SUMOylation machinery is

a strategy also used by Listeria monocytogenes, a food-borne

bacterial pathogen responsible for listeriosis. Indeed, infection

by L. monocytogenes leads to the degradation of Ubc9, the

human E2 SUMO enzyme (Ribet et al., 2010). Listeriolysin O is

a pore-forming toxin secreted by this bacterium, which plays

a fundamental role in bacterial virulence (Figure 1). Listeriolysin

O triggers the degradation of Ubc9, as well as the degradation

of some SUMOylated host proteins (Ribet et al., 2010). In

contrast to the ubiquitin system, which includes dozens of E2

enzymes in humans, the SUMO system has only one E2 enzyme.

Therefore, this degradation of Ubc9 leads to a blockade of the

SUMOylation machinery and to a global decrease in the level

of SUMO-conjugated host proteins in infected cells. Thus, by

decreasing SUMOylation in infected cells, Listeria may alter the

activities of host factors critical for infection (Ribet et al., 2010).

Pathogen-encoded deSUMOylasescan also cause a decrease

in the SUMOylation level of host proteins. Indeed, this is the case

for XopD, a protein injected by the plant pathogen Xanthomonas

campestris into the cytoplasm of plant cells. This protein is

a SUMO-specific protease, which induces deSUMOylation of

several host factors when it is expressed in plant cells (Hotson

et al., 2003). XopD is known to alter host transcription, to promote

pathogen multiplication, and to delay the onset of leaf chlorosis

and necrosis. However, the exact roles of deSUMOylation in

XopD’s effects are unknown (Kim et al., 2008).

In addition to the induction or inhibition of SUMOylation of host

proteins, viral proteins can be SUMOylated themselves.

However, the role that these modifications play in virulence is

unknown in most cases (Boggio and Chiocca, 2006). Surpris-

ingly, examples of bacterial factors directly SUMOylated by

host enzymes have not been identified. It is, however, likely

that future studies will unveil the existence of such modifications,

as well as their role in bacterial infection or in antibacterial

defenses.

Neddylation

Neddylation is another PTM that pathogens target during infec-

tion. Nedd8, which is a member of the ubiquitin-like protein

family, can be linked to cellular proteins in a fashion similar to

ubiquitin (reviewed in Rabut and Peter, 2008). The major class

of currently known Nedd8 substrates is Cullins. Cullins act as

scaffolding proteins in the assembly of multisubunit RING E3

ubiquitin enzymes, called Cullin RING ligases (CRLs). Neddyla-

tion of Cullins controls the activity of CRLs and thereby the ubiq-

uitination and degradation kinetics of CRLs substrates. As with

ubiquitin, Nedd8 can be deconjugated from its targets by dened-

dylases.

Bacterial and viral pathogens can interfere with the neddyla-

tion of host proteins. For example, the Epstein-Barr virus

encodes a protein BPLF1, which displays deneddylase activity

(Gastaldello et al., 2010). During infection, BPLF1 deneddylates

Cullins, thereby inhibiting the activity of CRLs and stabilizing

several CRL substrates. In particular, this leads to the deregula-

tion of the cell cycle and the establishment of an S-phase-like

cellular environment, which is required for efficient replication

of virus DNA (Gastaldello et al., 2010).

A recent study also reported that Cif (cycle-inhibiting factor),

a cyclomodulin translocated into cells by enteropathogenic

and enterohemorrhagic Escherichia coli, binds to Nedd8-conju-

gated CRLs of the host. This interaction inhibits the activity of the

CRLs, leading to a deregulation of the host cell cycle (Jubelin

et al., 2010). Proteins with in vitro deneddylase activity have

also been described in Chlamydia trachomatis, an obligate intra-

cellular bacterial pathogen. However, the role these deneddy-

lases play in infection remains unknown (Misaghi et al., 2006).


ISGylation

ISG15 (interferon stimulated gene 15) is an ubiquitin-like protein

with two ubiquitin domains. The expression of ISG15 is induced

in response to type I interferons (IFN), a family of cytokines

involved in the antiviral response. Consistent with this induction

in response to IFN, a growing number of studies are now high-

lighting the roles ISG15 plays in antiviral defense against several

types of viruses (reviewed in Skaug and Chen, 2010; Jeon et al.,

2010). Conjugation of ISG15 to target proteins requires the

activity of E1, E2, and E3 enzymes, which are also induced by

IFN. In contrast to the ubiquitin system, which includes hundreds

of E3 enzymes, one unique E3 ISG15 enzyme, namely HERC5,

modifies the vast majority of ISG15 substrates in human cells.

Like with other ubiquitin-like modifications, ISGylation is revers-

ible; specific proteases, called deISGylases, remove ISG15 from

its targets.

The antiviral activity of ISG15 can be due to either the

ISGylation of host proteins critical for infection or the direct

ISGylation of viral proteins (Skaug and Chen, 2010; Jeon et al.,

2010). This latter case has been described for the NS1 protein

of influenza A virus (NS1A), which is ISGylated during infection.

This modification of NS1A was linked to an impairment of influ-

enza replication, although the precise effect of the ISG15 addi-

tion on NS1A remains to be determined (Zhao et al., 2010;

Tang et al., 2010).

Interestingly, recent studies also proposed that the ISG15

conjugation system may modify broadly, and somehow nonspe-

cifically, newly synthesized proteins in a cotranslational manner

(Durfee et al., 2010). This implies that, in the context of an inter-

feron response, viral proteins, rather than cellular proteins, may

be the principal targets of ISGylation (Durfee et al., 2010).

Although only a small fraction of viral proteins might be

ISGylated, it was proposed that ISGylation of viruses’ structural

proteins, which precisely assemble into high-order structures,

might impair the production of infectious viral particles. Indeed,

this was demonstrated for the human papillomavirus HPV16.

ISGylation of a small proportion of its structural protein L1 was

sufficient to have a dominant-negative effect on virus infectivity

(Durfee et al., 2010). The authors postulated that the ISGylation

of host proteins could thus only be a side effect of the cell’s effort

to target viral proteins.

Consistent with the role of ISG15 in antiviral defense, several

viruses have evolved strategies to impair ISGylation (Skaug

and Chen, 2010; Jeon et al., 2010). In particular, studies

have identified several viral proteins that can either mimic

deISGylases or interfere with the ISGylation machinery of the

infected cell. Indeed, the papain-like protease of SARS corona-

virus and the ovarian tumor domain-containing proteases of

nairo- and arteriviruses all display ISG15-deconjugating activi-

ties (Lindner et al., 2005; Frias-Staheli et al., 2007). On the other

hand, NS1 protein of influenza B virus binds to ISG15 and inhibits

its conjugation to target proteins (Yuan and Krug, 2001). By

inhibiting ISG15 conjugation or increasing ISG15 deconjugation,

all these effector proteins were proposed to decrease the

potential antiviral effect of ISGylation.

The role of ISG15 in bacterial infections remains completely

unknown. According to the study by Durfee et al. (2010), the

participation of ISG15 in antibacterial defenses, if any, will prob-

ably rely on the ISGylation of cellular proteins rather than bacte-

rial proteins because the latter are not translated by the host cell

machinery. Nevertheless, investigating the role of ISG15 in infec-

tions by bacterial pathogens will undoubtedly provide exciting

insights into the field of host-pathogens interactions.

AMPylation and Eliminylation, New PTMs Mediatedby BacteriaAMPylation

AMPylation is the addition of an adenosine monophosphate

(AMP) group onto a threonine, tyrosine, or, possibly, serine

residue of a protein. The AMPylation of host proteins by bacterial

pathogens was recently detected in cells during an infection with

Vibrio parahaemolyticus, a human pathogen causing acute

gastroenteritis, and Histophilus somni, a pathogen responsible

for respiratory diseases and septicemia in cattle. Two virulence

factors produced by these extracellular bacteria, namely VopS

and IbpA, are able to reach the cytoplasm of host cells during

infection, where they use ATP to transfer an AMP moiety to

host Rho-GTPases (Figure 2) (Yarbrough et al., 2009; Worby

et al., 2009). This AMPylation alters the activity of Rho-GTPases,

which regulate the dynamics of the cell cytoskeleton.

The catalytic domain responsible for AMPylation was mapped

to the Fic domain (filamention induced by cAMP) of VopS and

IbpA. Fic domains are defined by a core sequence of nine amino

acids containing an invariant histidine residue that is essential for

the AMPylation (Yarbrough et al., 2009). Interestingly, proteins

containing Fic domains are found not only in prokaryotes but

also in eukaryotes, and the existence of eukaryotic proteins

able to catalyze AMPylation has been proposed (Worby et al.,

2009; Kinch et al., 2009). Thus, AMPylation might represent

a new and important posttranslational modification in eukaryotic

cells.

Legionella pneumophila is a human pathogen of the respira-

tory tract responsible for a severe form of pneumonia, called

Legionnaire’s disease. L. pneumophila encodes a factor, DrrA,

which AMPylates the host protein Rab1b, a small GTPase

involved in intracellular vesicular transport (Muller et al., 2010).

AMPylation of Rab1b leads to its constitutive activation, which

not only alters vesicular transport in infected cells but also

contributes to the formation of Legionella intracellular vacuoles

and aids bacterial replication.

Interestingly, the catalytic domain of DrrA is distinct from the

Fic domains observed in VopS and IbpA (Muller et al., 2010).

Thus, a wide diversity of both prokaryotic and eukaryotic

enzymes may catalyze AMPylation, a posttranslational modifica-

tion that might represent an unsuspected way of regulating

various signaling pathways in the cell.

Eliminylation

Phosphorylation was the first covalent protein modification

described. Since its discovery in the late 1950s, phosphorylation

has emerged as a common and fundamental PTM. Phosphory-

lation involves the reversible attachment of a phosphate group

to target proteins by forming a phosphoester bond. This addition

generally occurs on hydroxyl groups of serine, threonine, or tyro-

sine residues. Phosphorylation is reversible; phosphatases can

hydrolyze the phosphoester bond to release the phosphate

group and restore the amino acid in its unphosphorylated form.


Interestingly, a previously unknown enzymatic activity, called

phosphothreonine lyase, was recently identified in three different

bacterial factors (Li et al., 2007; Mazurkiewicz et al., 2008; Zhang

et al., 2007). These enzymes remove the phosphate group from

a threonine residue but, in contrast to classical phosphatases,

do not regenerate the hydroxyl group. Instead, this reaction,

nicknamed eliminylation, modifies threonine into dehydrobutyr-

ine, a residue that can no longer be phosphorylated (Brennan

and Barford, 2009).

The first factor identified with such activity is OspF, a protein

produced by Shigella flexneri, the causative agent of bacillary

dysentery in humans (Li et al., 2007). During infection, bacteria

directly secrete OspF into the host cell cytoplasm, where OspF

helps to dampen the host immune responses by irreversibly

dephosphorylating host MAP (mitogen-activated protein)

kinases (Figure 3) (Li et al., 2007; Arbibe et al., 2007). Phospho-

threonine lyases have been described only inS. flexneri,S.Typhi-

murium, and the plant pathogen Pseudomonas syringae, and

MAP kinases are the only known targets of this PTM. However,

we can expect that, as with AMPylation, some eukaryotic

enzymes may also display this activity and that eliminylation

might regulate numerous signaling pathways in eukaryotic cells.

Signaling Pathways Preferentially Targeted byPathogens by Alteration of Host PTMsSome pathogens produce several effectors that modulate the

activity of host cell proteins by stimulating or counteracting their

Figure 2. Pathogen-Mediated PTMs Target

the Cytoskeleton and ImmunoreceptorsBacteria effector proteins (green) control thedynamics of the host cell’s actin cytoskeleton byposttranslationally modifying Rho-GTPases (left).Viral effector proteins (blue) regulate posttransla-tional modification of immunoreceptors, such asthe major histocompatibility complex class I(MHC I) and the CD4 (cluster of differentiation 4)molecules (right), thereby decreasing their expres-sion at the cell surface and dampening immuneresponses.

PTMs. In this section, we will focus on

several key cellular pathways that are

preferentially targeted by pathogens

through these PTMs.

Regulation of the Cytoskeleton

Dynamics by PTMs

The niches occupied by pathogens within

their hosts are quite diverse. Whereas

some bacterial pathogens remain strictly

extracellular, other bacteria, as well as

viruses, invade host cells and replicate

therein. For viruses, entry into host cell

is strictly required for the synthesis of viral

proteins and the production of new infec-

tious viral particles. Bacteria take refuge

inside host cells to escape humoral

immune response and to replicate in

a well-protected environment. To enter

the cell and create such niches requires extensive remodeling

of the host cell cytoskeleton, a multiprotein assembly of struc-

tural and regulatory elements. Indeed, many pathogen-induced

PTMs target structural or regulatory components of the host

cell’s cytoskeleton.

Listeria monocytogenes is a bacterium that can induce its own

entry into a wide range of cells that are normally nonphagocytic.

This internalization requires interactions between surface

proteins of Listeria and host receptors. After successive PTMs,

these interactions trigger the recruitment of host factors and

the remodeling of host cell cytoskeleton required for internaliza-

tion of the bacteria (Figure 1). For example, the interaction

between the Listeria surface protein InlA and its cellular receptor

E-cadherin promotes Listeria’s invasion into epithelial cells of the

intestine. Activation of E-cadherin by InlA leads to phosphoryla-

tion and ubiquitination of E-cadherin by the Src kinase and

the Hakai E3 ligase, respectively. These PTMs trigger the recruit-

ment of the host’s clathrin-mediated endocytic machinery

followed by rearrangements of the actin cytoskeleton and inter-

nalization of the bacteria (Bonazzi et al., 2008).

In contrast, entry of Listeria into cells that do not express

E-cadherin is mediated by another surface protein, InlB, which

interacts with and activates Met, the hepatocyte growth factor

(HGF) receptor (Figure 1). Similar to HGF activation, Met activa-

tion by InlB induces its autophosphorylation and subsequent

monoubiquitination by the host E3 ligase Cbl. This leads to

the recruitment of the host’s clathrin-dependent endocytic


machinery, actin rearrangements, and ultimately, the internaliza-

tion of the bacteria (Veiga and Cossart, 2005; Veiga et al., 2007).

To avoid being killed, pathogens can also actively inhibit their

engulfment by professional phagocytes. The mechanisms

involved in this process may also require various pathogen effec-

tors to regulate the PTMs of host proteins (Figure 1). Pathogenic

Yersinia species are involved in human diseases, ranging from

enteric disorders to the plague. One virulence factor secreted

by Yersinia, YopH, displays potent phosphatase activity. It

decreases phosphorylation levels of host proteins involved in

focal adhesion complexes and impairs the cytoskeleton rear-

rangements required for bacterial uptake. Another factor of

Yersinia, YopT, is a protease that cleaves the membrane-

anchoring domain of host Rho-GTPases, leading to their irre-

versible detachment from the plasma membrane and their inac-

tivation (Figure 2 and Figure 1) (Shao et al., 2002). Thus, YopT

contributes to the inhibition of bacterial phagocytosis by pre-

venting rearrangements of the actin cytoskeleton.

Finally, some bacterial pathogens, such as Clostridium diffi-

cile, secrete several toxins that posttranslationally modify host

Rho-GTPases, leading to their constitutive activation, inactiva-

tion, or degradation (Figure 2). This alteration of Rho-GTPases

is widespread and allows bacteria to regulate the host cell’s

cytoskeleton in numerous ways, as well as gene transcription

Figure 3. Pathogen-Mediated PTMs Target

the MAP Kinase and NF-kB Signaling

PathwaysThe MAP kinase (left) and NF-kB (right) signalingcascades trigger immune responses in the hostcell during infections. Both bacterial (green) andviral (blue) effectors weaken these immuneresponses by inducing or counteracting post-translational modifications of key components inthese critical pathways.

and cytokine expression (reviewed in Ak-

tories and Barbieri, 2005).

Inhibition of the NF-kB Pathway

The NF-kB pathway is an example of

a pathway tightly regulated by ubiquitina-

tion (Figure 3). The NF-kB pathway plays

a central role in inflammation and in the

establishment of both innate and immune

responses. Specific signals, such as

cytokines or microbial signatures, acti-

vate this pathway by switching on the

IkB kinase (IKK) complex. This leads to

the phosphorylation of IkBa, an inhibitor

protein that sequesters transcription

factors of the NF-kB family in the cyto-

plasm. Phosphorylated IkBa is then

recognized by specific ubiquitin E3

ligases, polyubiquitinated with K48-

linked chains, and targeted to the protea-

some for degradation. Destroying IkBa

leads to the release of NF-kB transcrip-

tion factors, allowing them to translocate

into the nucleus and initiate transcriptionof various genes involved in host immune responses. Because

the NF-kB pathway plays a central role in immune responses,

there is a strong evolutionary pressure on pathogens to prevent

activation of this pathway during infection.

One possibility for dampening this pathway is to block the

ubiquitination of IkBa, thereby inhibiting its proteasomal degra-

dation and the translocation of NF-kB factors into the nucleus

(Figure 3). In numerous cases, factors achieve this goal by inter-

fering with the host ubiquitination machinery. For example,

S. flexneri secretes the effector OspG into the host cell’s

cytoplasm, where it binds to and inhibits UbcH5, a host E2

ubiquitin enzyme involved in IkBa ubiquitination (Kim et al.,

2005). The accessory protein Vpu (viral protein U) of HIV1 also

interferes with IkBa ubiquitination by inhibiting the E3 ubiquitin

ligase involved in IkBa’s modification (Bour et al., 2001). The

DUB-like SseL factor produced by S. Typhimurium inhibits

IkBa ubiquitination in response to the TNF-a cytokine, suggest-

ing that SseL acts directly by removing the K48-linked chains of

IkBa (Le Negrate et al., 2008).

Numerous factors also target the IKK complex directly

(Figure 3). For example, in addition to producing OspG, S. flex-

neri also secretes IpaH9.8, an effector with E3 ubiquitin ligase

activity. IpaH9.8 polyubiquitinates the NEMO/IKKg protein of

the IKK complex and targets it to the proteasome, thereby


impairing the phosphorylation and subsequent degradation of

IkBa (Rohde et al., 2007; Ashida et al., 2010). L. monocytogenes

intracellularly secretes InlC, which directly interacts with the

IKKa protein to block the phosphorylation of IkBa (Gouin et al.,

2010). Similarly, YopJ/P, an effector produced by pathogenic

Yersinia species, mediates the acetylation of the IKKa and

b proteins, which prevents their activation and subsequent

IkBa phosphorylation (Mittal et al., 2006).

Interestingly, commensal bacteria of the human intestine can

also act on the NF-kB pathway. Indeed, some bacterial fermen-

tation products, such as butyrate or other short-chained fatty

acids, can stimulate the local production of reactive oxygen

species in intestinal epithelial cells. This leads to the inactivation

of some redox-sensitive enzymes, such as E2 Nedd8 enzyme,

and therefore a decrease in the neddylation level of host

proteins. In this context, reduced neddylation levels, in particular

the decrease in Cullin-1 neddylation, have been associated with

a downregulation of the NF-kB pathway and hypothesized to

contribute to the inflammatory tolerance of the intestinal epithe-

lium toward commensal bacteria (Kumar et al., 2009).

Targeting of MAP Kinase Pathway

Similar to the NF-kB pathway, the MAP kinase pathway is

another central signaling cascade that is essential for the activa-

tion of host innate immune responses. Therefore, not surpris-

ingly, pathogens often target the MAP kinase pathway in order

to facilitate their infection (Figure 3). One effector protein

secreted intracellularly by Shigella is OspF, which possesses

phosphothreonine lyase activity. OspF irreversibly dephosphor-

ylates host MAP kinases and, therefore, was proposed to partic-

ipate in the dampening of host immune responses (Li et al., 2007;

Arbibe et al., 2007). Interestingly, other bacterial virulence

factors, such as SpvC from S. Typhimurium or HopAI1 from

the plant pathogen P. syringae, possess the same phospho-

threonine lyase activity as OspF and also target MAP kinases

of their hosts (Mazurkiewicz et al., 2008; Zhang et al., 2007). In

addition to these factors, the Yersinia YopJ/P effector can inac-

tivate host MAP kinases by catalyzing their acetylation (Mittal

et al., 2006; Mukherjee et al., 2006). Finally, the anthrax lethal

factor, a subunit of the Anthrax toxin encoded by Bacillus anthra-

cis, cleaves host MAP kinases, leading to their irreversible inac-

tivation (reviewed in Turk, 2007).

Regulation of Cellular Immunoreceptors

To avoid detection by the immune system, some pathogens

restrict the surface expression of fundamental molecules of the

immune system by subverting host ubiquitination (Figure 2).

For example, KSHV encodes two E3 ubiquitin ligases, K3 and

K5, which both target the host protein’s major histocompatibility

complex class I (MHC I). An essential player of the immune

response, MHC I alerts the immune system to intracellular path-

ogens by sampling the protein repertoire of host cells and then

presenting peptides to cytotoxic T lymphocytes. K3 rapidly

mediates the polyubiquitination of MHC I molecules at the

surface of the cell with K63-linked chains, leading to their endo-

cytosis and degradation. Interestingly, K5 also mediates polyu-

biquitination of MHC I but with mixed K63 and K11 chains,

instead of homotypic chains. Indeed, these mixed chains are

required for the internalization of MHC I by K5, thus highlighting,

for the first time, the putative importance of such mixed polyubi-

quitin chains in the control of immune responses (Boname

et al., 2010). Some herpesvirus E3 ubiquitin ligases downregu-

late MHC I molecules by triggering their degradation by the

ERAD (endoplasmic reticulum-associated protein degradation)

pathway (reviewed in Randow and Lehner, 2009). Some viral

proteins, such as HIV Vpu accessory protein, can act as adap-

tors of host E3 ubiquitin ligases to induce the proteasomal

degradation of other types of host immunoreceptors, such as

CD4 (cluster of differentiation 4) receptor on T cells (Schubert

et al., 1998). Finally, bacterial pathogens, such as Salmonella,

can decrease the expression of MHC class II molecules at the

cell surface by modulating their ubiquitination, which also leads

to the dampening of host immune responses (Lapaque et al.,

2009).

ConclusionResearchers have known for decades that pathogens interfere

with the host’s PTMs. However, the current ‘‘re-emergence’’ of

this field of research reflects the importance of controlling

PTMs during infection and the complexity of these processes

in host-pathogen interactions. In this Review, we focused on

how pathogens manipulate host PTMs and how they use these

PTMs to solve their own biological needs.

It should be stressed that pathogens may also actively co-opt

or be the passive targets of the host cell’s PTM machinery. As

mentioned above, pathogen-encoded proteins can indeed be

ubiquitinated, SUMOylated, or ISGylated, and like with host

proteins, PTMs of pathogen-encoded proteins regulate these

factors’ half-lives, activities, intracellular localization, or binding

to other host- or pathogen-encoded factors. Therefore, it is

tempting to speculate that the diversity of known PTMs affecting

pathogen-encoded proteins will greatly increase in the near

future.

As the number of studies reporting crosstalk between different

PTMs increases, an emerging idea is that PTMs are more

complex than originally anticipated. For example, in the NF-kB

signaling pathway alone, phosphorylation, SUMOylation, K63-

polyubiquitination, and K48-polyubiquitination act in synergy

to regulate the activation or the inhibition of transcriptional

responses. Targeting of these pathways by pathogens, there-

fore, often requires a tightly controlled orchestration of multiple

levels of PTMs.

Studies on pathogen interference with host protein PTMs has

provided numerous insights into cell biology over the years. In

particular, some pathogen effectors serve as invaluable tools

to study particular aspects of cell biology. For example, the

C3 exoenzyme from Clostridium ADP-ribosylates and inhibits

multiple Rho-GTPases. Therefore, the C3 protein has been used

successfully to highlight the specific role of the Rho-GTPase in

stress fiber formation and to study the regulation of the actin

cytoskeleton dynamics in eukaryotic cells (Ridley and Hall,

1992; Ridley et al., 1992).

Finally, the development of new technologies, such as

improvements in mass spectrometry (especially the SILAC

[stable isotope labeling of amino acids in cell culture] technique;

Mann, 2006), will undoubtedly increase the list of currently

known PTMs and facilitate the understanding of their roles in

host-pathogen interactions. Identifying pathogen-encoded


enzymes that catalyze specific PTMs critical for infection will

provide valuable new targets for drug development. Indeed,

the selective inhibition of these enzymes may constitute a prom-

ising strategy to counter these insidious invaders.

ACKNOWLEDGMENTS

We apologize to authors whose work could not be included because of space

constraints. Work in P.C.’s laboratory receives financial support from Institut

Pasteur, Inserm, INRA, ERC (advanced grant 233348), the Fondation le

Roch Les Mousquetaires, and the Fondation Louis-Jeantet. D.R. is supported

by a fellowship from the Association pour la Recherche sur le Cancer. P.C. is

an international research scholar of the Howard Hughes Medical Institute.

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Leading Edge

Review

Modifications of Small RNAsand Their Associated ProteinsYoung-Kook Kim,1 Inha Heo,1 and V. Narry Kim1,*1School of Biological Sciences and Center for National Creative Research, Seoul National University, Seoul 151-742, Korea


Small regulatory RNAs and their associated proteins are subject to diverse modifications that canimpinge on their abundance and function. Some of the modifications are under the influence ofcellular signaling, thus contributing to the dynamic regulation of RNA silencing.

IntroductionThe past decade has witnessed an explosion of research on

small regulatory RNAs that has yielded a basic understanding

of the many types of small RNAs in diverse eukaryotic species,

the protein factors involved, and the functions of key factors

along the RNA silencing pathways. Much more remains to be

learned, however, with recent studies unveiling interesting new

layers of regulation and complexity associated with small

RNAs. We now know that both small RNAs and their associated

protein factors can be modified at multiple steps in their biogen-

esis and effector pathways.

Insight into modifications of small RNAs came initially from

sequencing efforts, which made it clear that most microRNA

(miRNA) loci generate multiple isoforms (called isomiRs) apart

from the reference sequence (Morin et al., 2008). Alternative/

inaccurate processing partly explains the heterogeneity, but

a substantial portion of the variation is due to RNA modifications.

Small RNAs are modified either internally or externally by untem-

plated nucleotide addition, exonucleolytic trimming, 20-O-methyl

transfer, and RNA editing. Protein factors in RNA silencing path-

ways are also subject to various posttranslational modifications,

including phosphorylation, hydroxylation, ubiquitination, and

methylation. In this Review, we focus on the recent develop-

ments in the modifications of RNAs and proteins in RNA silencing

pathways.

Small RNA BiogenesisRNA silencing is a widespread mechanism of gene regulation in

eukaryotes. At the core of all RNA silencing pathways lie small

RNAs (20–30 nt in length) associated with the Argonaute family

proteins (Kim et al., 2009). Small RNAs provide the specificity

of regulation by base-pairing to the target nucleic acids while

the Argonaute proteins execute the silencing effects. The Argo-

naute (Ago) proteins are grouped into Ago and Piwi subfamilies,

and in animals, three types of small RNAs have been described:

microRNAs (miRNAs), small interfering RNAs (siRNAs), and Piwi-

interacting RNAs (piRNAs).

miRNAs (�22 nt) induce mRNA degradation and/or transla-

tional repression. Nucleotides 2–7, from the 50 end of the miRNA,

are referred to as the ‘‘seed’’ and are critical for hybridization to

the targets (Bartel, 2009). As a class, miRNAs are found in all

tissues, although each miRNA species displays a unique spatio-

temporal pattern of expression. An miRNA originates from a long

primary transcript (pri-miRNA) containing a local hairpin struc-

ture (Kim et al., 2009). In animals, the nuclear RNase III Drosha

liberates the hairpin-shaped precursor miRNA (pre-miRNA)

(Figure 1). The cytoplasmic RNase III Dicer removes the terminal

loop to produce a small RNA duplex, consisting of the functional

miRNA strand and the passenger (*) strand (miRNA/miRNA*).

The duplex then binds to the Argonaute loading complex

(comprised of Dicer, TRBP, and Ago), whose action leads to

the incorporation of the functional miRNA strand (mature miRNA)

into Ago. The plant miRNA system differs from its animal coun-

terparts in several aspects (Figure 2). The plant homolog of Dicer,

Dicer-like 1 (DCL1), cleaves both pri-miRNA and pre-miRNA in

the nucleus. Plant miRNAs generally show extensive comple-

mentary to their target mRNAs and induce endonucleolytic

cleavage of the targets.

Endogenous siRNAs (endo-siRNAs, �21 nt) are similar to

miRNAs in their binding to the Ago subfamily proteins, in their

dependence on Dicer for biogenesis, and in exerting their regu-

latory effects posttranscriptionally (Kim et al., 2009). But unlike

miRNAs, endo-siRNAs originate from long double-stranded

RNA precursors (dsRNAs), and their biogenesis does not require

processing by Drosha. Endo-siRNAs are abundant in lower

eukaryotes and in plants, whereas in mammals, they are found

in restricted tissues such as the ovary.

Piwi-interacting RNAs (piRNAs, 21–30 nt) associate with the

Piwi subfamily of Argonaute proteins. piRNAs mediate the

silencing of repetitive elements in gonads via transcriptional

and posttranscriptional silencing mechanisms. Production of

piRNAs is not dependent on RNase III nucleases, and the steps

and factors involved in their biogenesis remain largely unknown.

Modifications of Small RNAs30 End Modifications: Uridylation, Adenylation,

and 20-O-Methylation

The 30 ends of mature miRNAs are highly heterogeneous,

whereas the 50 ends are relatively invariable. The patterns

and sources of heterogeneity seem to vary depending on the

miRNA species and the cell types. The 30 end often contains

extra 1–3 nucleotides that do not match the genomic DNA

sequences. These untemplated nucleotides are added by


terminal nucleotidyl transferases that preferentially introduce

uridyl or adenyl residues to the 30 terminus of RNA.

The first indication of 30 end modification of small RNA came

from a hen1 mutant of Arabidopsis (Li et al., 2005). HEN1 is

a methyl transferase that adds a methyl group to the 20-OH at

the 30 end of RNA (Yu et al., 2005). In hen1 mutants, miRNAs

are reduced in abundance and become heterogeneous in size

due to uridylation at the 30 end. Because U tailing correlates

with the exonucleolytic degradation of mRNAs (Shen and

Goodman, 2004), it was postulated that uridylation induces

degradation of plant miRNAs and that the 20-O-methyl moiety

is required to protect small RNAs from uridylation and decay

(see below). Consistent with this notion, in green algae Chlamy-

domonas, a nucleotidyl transferase, MUT68, uridylates the 30

end of small RNA, and the RRP6 exosome subunit facilitates

small RNA decay in a manner dependent on MUT68 in vitro (Ibra-

him et al., 2010). Deletion of MUT68 results in elevated miRNA

and siRNA levels, indicating that MUT68 and RRP6 collaborate

in the turnover of mature small RNAs in plants.

Similar links between 20-O-methylation, uridylation, and decay

appear to exist in animals. A recent study on the zebrafish Hen1

homolog shows that piRNAs are uridylated and adenylated and

that piRNA levels are reduced in hen1 mutant germ cells (Kam-

minga et al., 2010). In flies and mice, piRNAs are methylated

by HEN1 orthologs, but the connection to stability control

remains unclear (Horwich et al., 2007; Kirino and Mourelatos,

2007; Ohara et al., 2007; Saito et al., 2007). In flies, dAgo2-bound

RNAs (mostly siRNAs) are protected by 20-O-methylation from

being uridylated/adenylated, which in turn induces 30

exonucleolytic trimming (Ameres et al., 2010). In nematode

worms, the role of 20-O-methylation has yet to be determined.

However, a subset of endo-siRNAs associated with an Ago

homolog CSR-1 is uridylated at the 30 end, and the uridyl trans-

ferase CDE-1 (also known as CID-1 or PUP-1) negatively regu-

lates these siRNAs, indicating that uridylation serves as a trigger

for decay (van Wolfswinkel et al., 2009).

Although mature miRNAs lack methylation in animals, uridyla-

tion plays a significant role in the control of miRNA biogenesis.

In mammalian embryonic stem cells, let-7 biogenesis is sup-

pressed by the Lin28 protein that binds to the terminal loop of

the let-7 precursors (Heo et al., 2008; Newman et al., 2008;

Rybak et al., 2008; Viswanathan et al., 2008). Of interest, Lin28

induces 30 uridylation of pre-let-7 by recruiting the terminal

nucleotidyl transferase TUT4 (also known as ZCCHC11) (Hagan

Figure 1. Modifications in the Animal

MicroRNA Pathway(Left) MicroRNAs (miRNAs) are subject to diversemodifications. Pri-miRNAs are edited by ADARs,which convert adenosine to inosine (I). RNA editinginhibits processing and/or alters target specificity.Pre-let-7 is regulated through uridylation. Lin28recognizes pre-let-7 and, in turn, recruits a nucleo-tidyl transferase TUT4 (mammal) or PUP-2(worms), which adds an oligo-uridine tail at the 30

end of RNA. The uridylated pre-miRNA is resistantto Dicer processing and subject to decay. TUT4also uridylates mature miRNA (miR-26), whichreduces miRNA activity. Another nucleotidyl trans-ferase GLD-2 adenylates mature miRNAs, whichreduces the activity of miRNA and/or increasesthe stability of specific miRNAs (such as miR-122).(Bottom) Mature miRNAs are degraded throughseveral mechanisms. In worms, a 50/30 exonu-clease XRN-2 degrades miRNAs that are releasedfrom Ago. In flies and humans, extensive pairingbetween miRNA/siRNA and target RNA triggerstailing as well as 30/50 trimming of miRNA/siRNA.(Right) Protein factors, which are involved in themiRNA pathway, are also subject to various post-translational modifications. Human Drosha isphosphorylated at two serine residues, S300/S302, by an unknown kinase. Phosphorylationlocalizes Drosha to the nucleus, where the pri-miRNA processing occurs. MAP kinases Erk1/2phosphorylate human TRBP at S142, S152,S283, and S286, which increases the proteinstability of TRBP and Dicer. Ago2 is regulated bymultiple modifications. A prolyl hydroxylaseC-P4H(I) hydroxylates P700 in human Ago2, whichenhances stability of Ago2 and increases P bodylocalization. Phosphorylation of human Ago2 atS387 by MAPKAPK2, which is induced by p38pathway, also promotes P body localization ofAgo2. However, the biological significance ofP body localization of Ago2 remains unclear. Inmice, a stem cell-specific E3 ligase, mLin41, ubiq-uitinates Ago2 and targets it for proteosome-dependent degradation.


et al., 2009; Heo et al., 2009). The oligo U-tail added by TUT4

blocks Dicer processing and facilitates the decay of pre-let-7.

The homologs of TUT4 may have related functions in other

organisms. In nematode worms, PUP-2 uridylates pre-let-7

in vitro and suppresses the let-7 function in vivo (Lehrbach

et al., 2009).

Let-7 is unlikely to be the only miRNA uridylated at the pre-

miRNA level. In support of this notion, untemplated 30 uridine is

frequently found in other mature miRNAs originating from the

30 arm of pre-miRNAs (but significantly less frequently in those

from the 50 arm) (Burroughs et al., 2010; Chiang et al., 2010).

Because untemplated uridylation is observed in cells lacking

Lin28, it will be interesting to determine which pre-miRNAs other

than pre-let-7 are controlled by uridylation and to identify addi-

tional factors required for pre-miRNA uridylation.

Although uridylation is generally thought to induce the decay

of small RNAs, adenylation may have the opposite conse-

quence. In cottonwood P. trichoacarpa, many miRNA families

are adenylated at their 30 ends, and adenylation prevents miRNA

degradation in in vitro decay assay (Lu et al., 2009). In the case of

mammalian miR-122, which is adenylated by cytoplasmic poly

(A) polymerase GLD-2 (or TUTase2), 30 end adenylation is also

implicated in its stabilization (Katoh et al., 2009). In the liver of

Gld-2 knockout mice, the steady-state level of mature miR-122

is reduced, and the abundance of target mRNAs of miR-122

increases.

However, a recent study indicates that GLD-2 adenylates

most miRNAs, and the adenylation may affect their activity rather

than stability (Burroughs et al., 2010). Deep sequencing of Ago-

associated small RNAs shows that adenylated miRNAs are

relatively depleted in the Ago2 and Ago3 complexes, suggesting

that adenylation may interfere with Ago loading. Similarly, it

has been reported that uridylation of mature miR-26 by TUT4

results in the reduction of miR-26’s activity without altering the

miRNA levels (Jones et al., 2009). Therefore, it remains an inter-

esting but yet unresolved issue whether or not uridylation/

adenylation affects the stability of miRNAs in animals. One may

speculate that 30 modified miRNAs enter the silencing complex

with altered frequencies, which in turn affects the small RNA’s

sensitivity to nucleases. Further examination is needed to iden-

tify the players involved in these processes, particularly the

nucleases that recognize a U/A tail, and to dissect their action

mechanisms.

miRNA Decay

Several nucleases degrade small RNAs (Figures 1 and 2). An

Arabidopsis enzyme SDN1 (small RNA degrading nuclease,

a 30-to-50 exonuclease) degrades single-stranded miRNAs

in vitro (Ramachandran and Chen, 2008). miRNAs accumulate

in a mutant lacking SDN1 and its related nucleases SDN2 and

SDN3, indicating that the SDN proteins may act redundantly to

degrade plant miRNAs. The 20-O-methyl group at the 30 end of

miRNAs, which is a general feature of plant miRNAs, has

a protective effect against SDN1 in in vitro assays. Of note, uridy-

lation causes a small but detectable protective effect in the same

in vitro assay, indicating that SDN1 is unlikely to be the nuclease

responsible for U-tail-promoted degradation. Given that RRP6

(a 30-to-50 exonuclease) facilitates decay of small RNAs in a

MUT68-dependent manner in Chlamydomonas extracts, multi-

ple enzymes may be involved in small RNA decay in plants,

playing partially overlapping but differential roles (Ibrahim et al.,

2010).

In C. elegans, XRN-2 (a 50-to-30 exonuclease) is involved in the

degradation of mature miRNAs (Chatterjee and Grosshans,

2009). Because miRNAs are tightly bound to and protected by

Ago, it is unclear how XRN-2 accesses the 50 end of an miRNA

for decay. Of interest, larval lysate promotes efficient release of

miRNA in vitro, implicating an as yet unknown factor that assists

the release of miRNA from the otherwise tightly associated Argo-

naute protein (Chatterjee and Grosshans, 2009). In Arabidopsis,

two XRN-2 homologs, XRN2 and XRN3, degrade the loop of

miRNA precursor following processing, but they do not affect

mature miRNA levels (Gy et al., 2007).

In mammals, a general nuclease for miRNAs has yet to be

identified. Knockdown of XRN-1 or an exosome subunit in

human cells results in only partial upregulation of miR-382, and

XRN-2 depletion does not have a significant effect (Bail et al.,

2010). Thus, it awaits further investigation whether or not there

is one major conserved pathway for miRNA decay in mammals.

There have been intriguing reports of regulated decay of

miRNAs. For instance, miR-29b is degraded in dividing cells

more rapidly than in mitotically arrested cells (Hwang et al.,

2007). In the central nervous system of Aplysia, the levels of

miR-124 and miR-184 decrease in 1 hr after treatment with

the neurotransmitter serotonin (Rajasethupathy et al., 2009).

Figure 2. RNA Modifications in the Plant miRNA PathwayIn plants, both pri-miRNA and pre-miRNA are cleaved by DCL1/HYL1complex. After cleavage, 30 ends of miRNA duplex are 20-O-methylated bya methyl transferase HEN1. The methylation protects miRNAs from uridylationand exonucleolytic degradation. In the green algae Chlamydomonas, the nu-cleotidyl transferase MUT68 attaches uridine residues at the 30 end of maturemiRNA lacking a methyl group. Then, the RRP6 exosome subunit, a 30-to-50

exonuclease, degrades the uridylated miRNAs. In Arabidopsis, a 30/50

exonuclease SDN1 is reported to degrade mature miRNAs.


Because U0126, an inhibitor of mitogen-activated protein kinase

(MAPK), blocks the reduction of miR-124, the decay process

may be dependent on the MAPK pathway. Of interest, a study

on mammalian neuronal cells shows that most miRNAs turn

over more rapidly in neurons than in other cell types (Krol et al.,

2010). Neuronal activation accelerates decay of the miRNAs,

whereas blocking neuronal activity stabilizes the miRNAs. It

will be exciting to discover the nuclease(s) and the upstream

signals for miRNA degradation in these systems.

Recently it has been shown that a polynucleotide phosphory-

lase (PNPase, a type I interferon-inducible 30-to-50 exonuclease)

binds specifically to several miRNAs (miR-221, miR-222, and

miR-106b) and induces rapid turnover in a human melanoma

cell line (Das et al., 2010). Because there is no apparent

commonality in terms of the sequences, it is unclear how

PNPase recognizes the miRNAs specifically.

As mentioned above, there is substantial evidence linking uri-

dylation/adenylation and exonucleolytic attack on small RNAs.

A recent study provides evidence that extensive complemen-

tarity between a small RNA and its target RNA triggers uridyl/

adenyl tailing as well as 30/50 trimming in flies and humans

(Figure 1) (Ameres et al., 2010). Animal small RNAs with high

complementarity to the targets, such as piRNAs and fly endo-

siRNAs, appear to be generally protected by 20-O-methylation

at the 30 end like plant small RNAs. It has been postulated that

animal miRNAs, which do not carry methylation, maintain only

partial complementarity with their targets so as to avoid tailing

and trimming of miRNAs. Of note, viruses seem to exploit a

related miRNA decay pathway to invade host cells more effec-

tively. Herpesvirus saimiri, a family of primate-infecting herpesvi-

ruses, expresses viral noncoding RNAs called HSURs (H. saimiri

U-rich RNAs). A recent report reveals that HSURs rapidly down-

regulate host miR-27 and that base-pairing between HSUR and

miR-27 is required for the degradation (Cazalla et al., 2010).

These discoveries imply an additional layer of stability control

of small RNAs, which is influenced by the interaction with the

target RNA.

miRNA Editing

Adenosine deaminases acting on RNAs (ADARs) convert

adenosine to inosine on the dsRNA region of small RNA precur-

sors (Figure 1 and Figure 3A). Because inosine (I) pairs with

cytosine instead of uridine, such edits could alter the structure

of small RNA precursor, thereby interfering with processing.

For instance, editing of pri-miR-142 by ADAR1 and ADAR2

suppresses Drosha processing (Yang et al., 2006), whereas

that of pre-miR-151 by ADAR1 interferes with Dicer processing

(Kawahara et al., 2007a). Because hyperedited dsRNAs can be

targeted by the nuclease Tudor-SN, RNA editing may also desta-

bilize small RNA precursors (Scadden, 2005). In rare cases, RNA

editing occurs in the seed sequence of miRNA, changing the

targeting specificity. In the brain, where ADAR is abundant,

miR-376 cluster miRNAs are frequently edited in the seed

region and are redirected to repress a different set of mRNAs

(Kawahara et al., 2007b). High-throughput sequencing of the

fly endo-siRNA pool also reveals evidence for RNA editing

(Kawamura et al., 2008). The precursors of endo-siRNAs (long

hairpins and sense-antisense pairs) may be targeted by ADARs,

although the functional significance of this siRNA modification is

unknown.

Posttranslational Protein ModificationsPhosphorylation of RNase III Enzymes

Human Dicer interacts with two related dsRNA-binding proteins,

TRBP and PACT. Although they do not influence Dicer process-

ing itself, TRBP and PACT stabilize Dicer and may also function

in RISC assembly (Chendrimada et al., 2005; Haase et al., 2005;

Lee et al., 2006). A recent study indicates that four serine

residues of human TRBP (S142, S152, S283, and S286) are

phosphorylated by the MAP kinase Erk, which controls cell

proliferation, survival, and differentiation (Figure 1) (Paroo et al.,

Figure 3. Modifications in the Endo-siRNA

and piRNA Pathways(A) Endogenous small interfering RNAs (endo-siRNAs) are processed from long dsRNAs ina Dicer-dependent manner and are loaded ontoAgo proteins. High-throughput sequencing datashow that the adenosine-to-inosine (I) editingoccurs in fly endo-siRNAs, likely by ADAR,although the role of RNA editing is unknown. Flyendo-siRNAs bound to dAgo2 are 20-O-methyl-ated by HEN1 homolog, which protects RNAsfrom uridyl/adenyl tailing and degradation. Inworms, a subset of endo-siRNAs, which are asso-ciated with an Ago homolog CSR-1, is uridylatedat the 30 end by the nucleotidyl transferase CDE-1.(B) piRNAs are generated from single-strandedRNA precursors that are processed by primaryprocessing and/or secondary processing (ping-pong amplification cycle). piRNAs are associatedwith Piwi subfamily proteins (PIWI). Animal piRNAsare 20-O-methylated by HEN1 orthologs. In zebra-fish, depletion of hen1 induces uridylation ofpiRNAs and facilitates decay, suggesting thatmethylation stabilizes piRNAs. However, the phys-iological significance of piRNA methylation in flies

and mammals remains unclear. PIWI proteins are methylated at arginine residues (sDMA, symmetrical dimethyl arginine) at their N termini by orthologs of themethyl transferase PRMT5. In flies and mice, TDRD proteins interact with PIWI proteins through sDMA and may play important roles in piRNA metabolism.


2009). Phosphorylation enhances protein stability of TRBP,

consequently elevating Dicer protein levels. Intriguingly, TRBP

phosphorylation preferentially increases growth-promoting

miRNAs such as miR-17, whereas tumor-suppressive let-7 is

reduced. The mechanism of selective downregulation of let-7

is unclear, but it may be an indirect effect. An interesting implica-

tion of these findings is that the MAPK/Erk pathway exerts its

effects, in part, by regulating miRNA biogenesis.

Drosha, a nuclear enzyme for pri-miRNA processing (Lee

et al., 2003), has recently been shown to be a direct target of

posttranslational modification (Tang et al., 2010). Mass spec-

trometry and mutagenesis studies reveal that human Drosha is

phosphorylated at serine 300 (S300) and serine 302 (S302)

(Figure 1). Phosphorylation of these residues is essential for

the nuclear localization of Drosha and is required for pri-miRNA

processing. Because both endogenous and overexpressed

Drosha localize to the nucleus constitutively, it is unclear whether

or not the phosphorylation at S300/S302 is a regulated process.

Understanding the physiological significance of this regulation

will require the identification of the kinase that phosphorylates

Drosha.

Argonaute2 Is a Target of Multiple Modifications

Ago2 is subject to multiple posttranslational modifications

(Figure 1). Human Ago2 binds to the type I collagen prolyl-4-

hydroxylase (C-P4H(I)) that hydroxylates Ago2 at proline 700

(Qi et al., 2008). Depletion of C-P4H(I) reduces the stability of

the Ago2 protein and, accordingly, downregulates siRNA-medi-

ated silencing. Furthermore, hydroxylation is required for Ago2

localization to the processing body (P body), a cytoplasmic

granule that is thought to be a site for RNA storage and degrada-

tion. P body localization of Ago2 is also enhanced by phosphor-

ylation at serine 387, which is mediated by the p38 MAPK

pathway (Zeng et al., 2008). However, given the controversy

over the direct role of P body in small RNA-mediated silencing,

the biological significance of P body localization of Ago2 remains

unclear.

Ubiquitination also plays a part in the control of Ago2. Mouse

Lin41 (mLin41 or Trim71), a stem cell-specific Trim-NHL protein,

inhibits the miRNA pathway (Rybak et al., 2009). As an E3 ubiq-

uitin ligase, mLin41 ubiquitinates Ago2 and targets it for protea-

some-dependent degradation. Of interest, mLin41 is a target of

let-7 miRNA, suggesting that mLin41 and let-7 may be engaged

in a reciprocal negative feedback loop. Recently, other Trim-NHL

proteins have been reported to associate with the Argonaute

proteins and affect miRNA pathway. Mei-P26 (fly) inhibits miRNA

biogenesis, whereas TRIM32 (mouse) and NHL-2 (worm) acti-

vate the miRNA pathway (Hammell et al., 2009; Neumuller

et al., 2008; Schwamborn et al., 2009). Their mechanism of

action appears to be different than that of mLin41 because the

E3 ligase activity of Mei-P26 and TRIM32 is dispensable for their

effects and because NHL-2 enhances miRNA activity without

a change in miRNA levels.

Tudor Regulates PIWI Proteins

The PIWI (P element-induced wimpy testis) clade proteins bind

to Piwi-interacting RNAs (piRNAs) and silence transposable

elements in gonads. Mouse has three PIWI homologs (MILI,

MIWI, and MIWI2), and there are three PIWI proteins in flies

(Aubergine [Aub], AGO3, and Piwi) (Kim et al., 2009). Recent

studies have revealed that PIWI proteins carry symmetrical

dimethyl arginine (sDMA) at their N termini. Arginine methylation

of PIWI is mediated by a methyl transferase PRMT5 (dPRMT5/

capsuleen [csul]/dart5 in Drosophila) (Figure 3B) (Heo and Kim,

2009; Siomi et al., 2010). sDMA is recognized by Tudor

domain-containing proteins (TDRDs), which are critical for germ-

line development. In both flies and mice, deletion of TDRDs alters

piRNA abundance and/or composition, indicating that TDRDs

play important roles in the piRNA metabolism through specific

binding to the sDMAs of PIWI proteins. How TDRDs act in the

piRNA pathway at a molecular level awaits further investigation.

PerspectivesAs we delve deeper and wider into the small RNA world, the

emerging landscape becomes ever more complex on both the

RNA and protein sides. High-throughput analyses have uncov-

ered a considerable heterogeneity in small RNA populations.

Some isomiRs are expressed differentially in certain tissues,

suggesting that these variations may be associated with specific

regulatory functions (Chiang et al., 2010). Biochemical and

genetic studies also provide substantial evidence for the regula-

tory roles of the modifications discussed in this Review. Thus, it

is likely that at least some of the observed heterogeneity reflects

multiple layers of regulation. We should be cautious, however, in

extrapolating the current evidence because it is unclear how

much fraction of the small RNA and protein modifications trans-

late into functional consequences and whether certain modifica-

tions simply reflect the noise of RNA metabolism.

In addition to the functionality issue, a number of key questions

remain to be answered. Are there conserved pathways and

enzymes for RNA and protein modifications? If so, what are

the similarities and differences? 20-O-methylation is applied to

many small RNA pathways, but the details differ significantly in

different systems. For instance, plant HEN1 acts on dsRNA

duplexes, whereas animal HEN1 homologs methylate ssRNA

loaded on Argonaute proteins. Uridylation/adenylation is carried

out by a family of ribonucleotidyl transferases. How each

member selectively recognizes its substrates is largely unknown.

RNA stability is likely to play important roles in RNA silencing

pathways. Decay pathways of small RNA are beginning to be

unraveled, but there is no consensus between different species

as yet. One possibility is that multiple enzymes act in parallel as

in the mRNA decay pathway, which involves several 30 exonucle-

ases, 50 exonucleases, and endonucleases. Some of the decay

enzymes may function redundantly, and it remains one of the

major challenges in the field to identify them. Protein modifica-

tion is also emerging as one of the key regulatory layers.

Outstanding questions include which enzymes are involved,

what the in vivo significance of such modifications is, and

whether the protein modifications are developmentally regu-

lated. Future studies will reveal new types of modifications, addi-

tional regulatory factors, and their biological relevance.

The RNA silencing machinery should respond accurately to

developmental and environmental cues. Most signaling path-

ways are thought to be connected to RNA silencing, but we

are just beginning to understand the molecular links between

RNA silencing and cell signaling. What the upstream signals

are, how certain RNAs and proteins get specifically recognized,


and what the downstream effects of the modifications are await

elucidation. We also need to understand the interplay between

different modifications. There appears to be a crosstalk between

certain modifications of RNA (such as methylation, uridylation,

and decay), which may influence their fate and function. It is likely

that there is a crosstalk between the different posttranslational

modifications in the proteins involved in the biogenesis and

effector functions of small RNA silencing pathways. Under-

standing these networks will undoubtedly provide ample oppor-

tunities to manipulate RNA silencing and will reveal new lessons

about gene regulation.

ACKNOWLEDGMENTS

We thank members of V.N.K.’s laboratory for helpful discussions and

comments. This work was supported by the Creative Research Initiatives

Program (20100000021) and the National Honor Scientist Program

(20100020415) through the National Research Foundation of Korea (NRF)

and the BK21 Research Fellowships (I.H.) from the Ministry of Education,

Science and Technology of Korea. We apologize to authors whose work has

not been covered because of space limitations.

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The ER UDPase ENTPD5 Promotes ProteinN-Glycosylation, the Warburg Effect,and Proliferation in the PTEN PathwayMin Fang,1 Zhirong Shen,1 Song Huang,1 Liping Zhao,1 She Chen,2 Tak W. Mak,3 and Xiaodong Wang1,2,*1Howard Hughes Medical Institute and Department of Biochemistry, University of Texas Southwestern Medical Center at Dallas,

5323 Harry Hines Boulevard, Dallas, TX 75390, USA2National Institute of Biological Sciences, Zhongguancun Life Science Park, Beijing 102206, China3The Campbell Family Institute for Breast Cancer Research, Princess Margaret Hospital, Toronto, ON M5G 2M9, Canada*Correspondence: [email protected]

DOI 10.1016/j.cell.2010.10.010

SUMMARY

PI3K and PTEN lipid phosphatase control the level ofcellular phosphatidylinositol (3,4,5)-trisphosphate,an activator of AKT kinases that promotes cellgrowth and survival. Mutations activating AKT arecommonly observed in human cancers. We reporthere that ENTPD5, an endoplasmic reticulum (ER)enzyme, is upregulated in cell lines and primaryhuman tumor samples with active AKT. ENTPD5hydrolyzes UDP to UMP to promote protein N-glyco-sylation and folding in ER. Knockdown of ENTPD5 inPTEN null cells causes ER stress and loss of growthfactor receptors. ENTPD5, together with cytidinemonophosphate kinase-1 and adenylate kinase-1,constitute an ATP hydrolysis cycle that convertsATP to AMP, resulting in a compensatory increasein aerobic glycolysis known as the Warburg effect.The growth of PTEN null cells is inhibited bothin vitro and in mouse xenograft tumor models.ENTPD5 is therefore an integral part of the PI3K/PTEN regulatory loop and a potential target for anti-cancer therapy.

INTRODUCTION

Class I phosphatidylinositol 3-kinases (PI3Ks) and lipid phospha-

tase PTEN balance cellular response to growth and survival

signals (reviewed by Engelman et al., 2006). In response to acti-

vation of receptor tyrosine kinases, PI3K phosphorylates phos-

phatidylinositol 4,5-bisphosphate (PIP2) at the 3-OH position of

the inositol ring to generate phosphatidylinositol 3,4,5-trisphos-

phate (PIP3) that recruits and activates serine/threonine kinase

AKT (Whitman et al., 1988; Franke et al., 1997; Stephens et al.,

1998). AKT subsequently activates many downstream targets

for cell growth and survival, including the rapamycin-sensitive

mTOR complex 1 (mTORC1), which then phosphorylates

p70S6K and translation initiation factor 4E-BP1 to accelerate

the translational rate, thus accommodating rapid growth (Fingar

et al., 2002). PTEN, by dephosphorylating PIP3 back to PIP2,

antagonizes the signal generated by PI3K (Maehama and Dixon,

1998). The importance of the PI3K/PTEN pathway has been

manifested by frequent PI3K gain of function, or PTEN loss of

function, in a variety of human cancers (reviewed by Yuan and

Cantley, 2008; Keniry and Parsons, 2008).

AKT activation also contributes to the elevation of aerobic

glycolysis seen in tumor cells, known as the Warburg effect

(Elstrom et al., 2004; Warburg, 1925). AKT promotes cell-surface

expression of glucose transporters while sustaining activation of

hexokinase and phosphofructose kinase-1 (PFK1), thus acceler-

ating influx and capture of glucose for glycolysis (reviewed by

Vander Heiden et al., 2009). Of interest, in cancer cells, there is

invariant expression of the embryonic M2 splice version of pyru-

vate kinase, an enzyme working in the last step of glycolysis,

instead of a more active M1 splicing isoform expressed in

most of the adult tissues (Christofk et al., 2008). The combined

effects of more glucose entering into the glycolysis pathway

and slowing down pyruvate kinase activity build up intermediate

metabolites for synthesis of growth-enabling macromolecules.

One noticeable example is the entry of glucose-6-phosphate

to the pentose shunt pathway to generate ribose for nucleotide

synthesis (reviewed by Vander Heiden et al., 2009).

Another outlet of glucose-6-phosphate is to form UDP-

glucose, a substrate for protein glycosylation. In mammalian

cells, most secreted proteins and membrane proteins are glyco-

sylated at the asparagine (Asn) sites, i.e., N-glycosylated. Of

interest, receptor tyrosine kinases that promote cell growth

and proliferation, such as the epidermal growth factor receptor,

EGFR, are much more highly N-glycosylated than receptors

whose functions do not (Lau et al., 2007). Most of the glycosyla-

tion reactions happen in the Golgi apparatus, with two known

exceptions. One is the dolichol-linked 14 sugar core glycan

(Glc3Man9GlcNAc2) that is synthesized in cytoplasm and ER

membrane before being flipped into the lumen of ER, where it

is transferred to the Asn of the nascent polypeptide chain (re-

viewed by Helenius and Aebi, 2004). Another is reglucosylation

in ER after the third and second glucose (Glc) on the core glycan

are trimmed by glycosidase I and II. Trimming and reglucosyla-

tion by UDP-glucose:glycoprotein glucosyltransferase (UGGT)

generate monoglucosylated structures that are recognized by

Cell 143, 711–724, November 24, 2010 ª2010 Elsevier Inc. 711

calnexin/calreticulin, an ER molecular chaperone system for

N-glycosylated proteins (reviewed by Ellgaard et al., 1999).

The removal and addition of glucose allow the binding and

release of calnexin/calreticulin to and from nascent polypeptide

chains until the proteins are correctly folded and transferred to

Golgi for further glycosylation. If proteins are misfolded beyond

repair, they are subjected to degradation by the ER-associated

protein degradation system (ERAD) (reviewed by Fewell et al.,

2001).

During a study using the embryonic fibroblasts (MEFs) from

the PTEN null mice and PTEN heterozygous littermates (Stam-

bolic et al., 1998), we made a surprising finding that an ER

UDP hydrolysis enzyme is upregulated by AKT activation. This

enzyme, ENTPD5, seems to mediate many of the observed

cancer-related phenotypes associated with AKT activation.

RESULTS

PTEN Knockout MEFs Have an Elevated Activity thatHydrolyzes ATP to AMPAs reported previously, the PTEN null MEFs showed elevated

levels of phosphorylated AKT and p70S6 kinase, whereas the

total protein level of these two kinases remained the same as

in PTEN heterozygous MEFs (Stambolic et al., 1998) (Figure 1A).

We noticed that S-100 cell extracts (prepared after collecting the

supernatants of 100,000 3 g spin of broken cells) from PTEN null

MEFs had a lower ATP level compared to that from the heterozy-

gous MEFs (Figure 1B, columns 7 and 8). Given that cellular ATP

levels are relatively stable, we reasoned that the difference in

their ATP contents occurred during S-100 preparation, which

took about 1 hr. Indeed, as shown in Figure 1B, the ATP levels

in PTEN null MEFs were only slightly lower than those in the

heterozygous MEFs if the measurement was carried out immedi-

ately after cells were harvested (Figure 1B, columns 1 and 2).

When the broken cell suspension, or supernatants after

10,000 3 g spin (S-10), or S-100 were incubated on ice for 1 hr

before the ATP levels were measured, ATP concentrations in

the extracts from PTEN knockout MEFs were much lower than

those from heterozygous MEFs (Figure 1B, columns 3–8). Such

an observation indicated that there was higher ATP hydrolysis

activity in the PTEN knockout cell extracts. To measure this

activity directly, we incubated a-P32-ATP with the S-100 extracts

and analyzed the radioactivity using thin layer chromatography.

As shown in Figure 1C, more radiolabeled ATP was hydrolyzed

when incubated with the S-100 from PTEN null MEFs, and the

nucleotide was hydrolyzed all the way to AMP.

To sort out whether the accelerated ATP hydrolysis was due to

a specific activity or a combination of nonspecific ATPases, we

fractioned the same amount of S-100 extracts from PTEN null

and PTEN heterozygous MEFs side by side on a Q Sepharose

ion-exchange column. The fractions from each column run

were dialyzed, and ATPase activity was measured by adding

each column fraction to the S-100 from PTEN heterozygous

MEF, which served as the baseline activity. A single peak of

elevated ATP-to-AMP activity centered at fractions 11–13 was

observed in fractionated S-100 from PTEN null MEFs, whereas

much less activity was seen in the corresponding fractions

from PTEN heterozygous MEFs (Figure 1D).

ENTPD5 Is Responsible for the Elevated ATPase Activityin PTEN Knockout CellsWe decided to purify the ATPase from large-scale cultured PTEN

knockout MEFs. We took 800 mg of S-100 from PTEN null MEFs

and put it through five chromatographic steps (Figure 2A). The

ATP hydrolysis activity was measured as in Figure 1D, and the

active fractions from each column step were pooled, dialyzed,

and loaded onto the next column. Finally, after a Superdex 200

gel-filtration column, the active fractions were loaded onto

a 100 ml Mini Q column and the bound protein was eluted with

a linear salt gradient. Fractions of 100 ml were collected and as-

sayed. Shown in Figure 2B, a single ATP hydrolysis peak

centered at fraction 6 was observed. When these fractions

were analyzed by SDS-PAGE followed by silver staining,

a protein band just below the 50 kDa molecular weight marker

correlated perfectly with the activity.

This protein was excised from the gel and subjected to mass

spectrometry analysis. The enzyme was identified as ectonu-

cleoside triphosphate diphosphohydrolase 5, ENTPD5, a mem-

ber of the ENTPD enzyme family known to hydrolyze tri- and/or

diphospshonucleotide to monophosphonucleotide (reviewed

by Robson et al., 2006).

To verify that ENTPD5 is indeed the enzyme that caused the

higher rate of ATP-to-AMP conversion in PTEN null MEFs, we

first did a western blotting analysis of ENTPD5 in these MEFs.

As shown in Figure 2C (bottom), ENTPD5 was only prominently

detected in PTEN null extracts, but not in PTEN heterozygous

extracts (Figure 2C, lanes 1 and 2). When mouse ENTPD5 was

exogenously expressed in the PTEN heterozygous MEFs, the

extracts from these cells showed the ability to hydrolyze ATP

to AMP just like that from PTEN null cells (Figure 2C, lanes

3–5). Moreover, when ENTPD5 was knocked down in PTEN

null MEFs with two different siRNA oligos, the ATP-to-AMP

conversion was diminished in each case, and a control siRNA

oligo had no effect (Figure 2C, lanes 6–10).

To confirmthat the elevated level ofENTPD5 isdue todeletion of

PTEN, we transfected a wild-type PTEN cDNA, or the phospha-

tase active site mutant (PTEN C124S), into PTEN null MEFs.

Indeed, restoring PTEN expression in these cells lowered phos-

phoAKT and diminishedENTPD5expression, whereas the catalyt-

ically dead mutant PTEN had no effect (Figure 2D, lanes 2 and 3).

Consistently, treatment of PTEN null MEFs with a PI3 kinase inhib-

itor also lowered the level of ENTPD5 (Figure 2E, lanes 2 and 3).

The upregulation of ENTPD5 in PTEN null cells is at transcrip-

tional level. Its mRNA is 6-fold higher in PTEN null MEFs

compared to that in PTEN heterozygous MEFs (Figure S1 avail-

able online). The promoter region of ENTPD5 is negatively regu-

lated by the FoxO family of transcription factors (Figure S2),

which upon phosphorylation by AKT, are displaced from the

nucleus into the cytoplasm (Brunet et al., 1999).

To directly demonstrate the nucleotide hydrolysis activity of

ENTPD5, we generated recombinant human ENTPD5 protein

in insect cells using a baculovirus vector and purified the enzyme

to homogeneity (Figure 3A, right). The purified enzyme was then

incubated with ATP, ADP, CTP, CDP, GTP, GDP, UTP, and UDP,

and the released phosphate was measured. Unexpectedly, the

purified recombinant ENTPD5 could only hydrolyze UDP and

GDP (Figure 3A, left).

712 Cell 143, 711–724, November 24, 2010 ª2010 Elsevier Inc.

UMP or GMP Is a Required Cofactorfor the ATP Hydrolysis ActivityDuring our ENTPD5 purification efforts, we noticed that a small

molecule cofactor was required for the observed ATP-to-AMP

hydrolysis activity. S-100 extracts from PTEN null MEFs lost

ATP-to-AMP converting activity after dialysis (Figure 3B,

lane 4), and the activity was restored with addition of a small

molecule fraction prepared by a 10 kDa cutoff filter (Figure 3B,

lane 6). There was no difference in such a small molecule in

PTEN heterozygous and PTEN null MEFs (Figure 3B, lanes 6

Lu

min

es

ce

nc

e I

nte

ns

ity

0

100000

200000

300000

PT

EN

+/-

PT

EN

-/-

PT

EN

+/-

PT

EN

-/-

PT

EN

+/-

PT

EN

-/-

PT

EN

+/-

PT

EN

-/-

Sta

rt

Bro

ke

n c

ells

S-1

0

S-1

00

B

PT

EN

+/-

PT

EN

-/-

AMP

ADP

ATP

C

800800

10001000

12001200

14001400

16001600

0

250250

500500

750750

10001000

Na

Cl

(mM

)

800800

10001000

12001200

14001400

16001600

0

250250

500500

750750

10001000

UV

Ab

s. (m

AU

)U

V A

bs. (m

AU

)

Na

Cl

(mM

)

+/-

Sta

rt

-/-

Start

+/-

Flo

w

-/-

Flo

w

3 4 5 6 7 8 9 1011121314 1516

17+

18

Fraction №

S-100 (+/-)

+/-

-/-

D

AMP

ADP

ATP

AMP

ADP

ATP

UV

Ab

s. (m

AU

)U

V A

bs. (m

AU

)

PTEN

pAKT

AKT

pP70S6K

PTEN+/-

PTEN-/-

P70S6K

Actin

A

*

Figure 1. Identification of ATP Hydrolysis Activity in PTEN Knockout MEF Cells

(A) Total cell extracts from PTEN+/� and PTEN�/� cells were prepared as described in Experimental Procedures. Aliquots of 10 mg protein were subjected to 10%

SDS-PAGE followed by western analysis of PTEN (asterisk denotes a cross-reactive band), phosphorylated AKT (pAKT), AKT, phosphorylated P70S6 kinase

(pP70S6K), P70S6 kinase, and b-actin.

(B) Cell extracts were prepared from PTEN+/� and PTEN�/� cells, and at indicated steps of preparation, aliquots of 20 ml samples were incubated on ice for 1 hr

followed by immediate measurement of ATP using a Cell Titer-Glo kit. Error bar represents standard deviation of two independent experiments.

(C) Aliquots of 30 mg of S-100 fractions from PTEN+/� or PTEN�/� cells were incubated with a-P32-labeled ATP and analyzed by TLC as described in the

Experimental Procedures. Positions for ATP, ADP, or AMP were indicated.

(D) 6 ml each of S-100 from PTEN+/� or PTEN�/� cells (3.5 mg/ml) was separated by a 1 ml Q Sepharose HP column with a salt gradient elution as indicated.

Fractions of 1 ml were collected and dialyzed overnight at 4�C. 10.5 ml of each fraction was mixed with another 10.5 ml of undialyzed S-100 from PTEN+/� cells

and assayed for ATP hydrolysis activity as in (C). The positions of ATP, ADP, and AMP were indicated. FPLC histograms were presented in top panels.


and 8), and the molecule was also present in S-100 from HeLa

cells (Figure 3B, lane 10), which have a wild-type PTEN.

Based on its biochemical properties, we deduced that the

cofactor is a nucleotide. Testing a variety of nucleotides revealed

that uracil and guanine, either in tri-, di-, or monophosphate

form, substituted the small molecule fraction from cells

(Figure 3C, lanes 1–15). In contrast, thymidine nucleotides

have no activity, whereas CMP only showed a slight activity.

To see whether the conversion of UTP/UDP to UMP is neces-

sary for the observed activity, we tested various forms of

nonhydrolyzable uracil, including UTPgS, UTPaS, and UMP-

PNP (Figure 3D). All of these nucleotides worked except UTPaS,

Start PTEN-/- S-100

SP HP

Q HP

Phenyl HP

Super-dex 200

Mini Q

A

Ve

c

PT

EN

PT

EN

-C

D

PTEN-/- MEF

Lane 1 2 3

Tranfection

ENTPD5

pAKT

AKT

ß-Actin

PTEN

D

Lane 1 2 3

Cell Line +/- -/-

LYDMSOTreatment

ENTPD5

pAKT

AKT

ß-Actin

E

3 4 5 6 7 8 9 1110

250KD

150KD

100KD

75KD

50KD

37KD

25KD

20KD

15KD

10KD

AMP

ADP

ATP

120mM127.5mM NaCl

B

Fraction №

ENTPD5

+/-

-/-

AMP

ADP

ATP

ENTPD5

+/- V

ecto

r

+/- E

NT

PD

5 1#

+/- E

NT

PD

5 2#

+/-

-/-

-/- s

iR

NA

G

FP

-/- s

iR

NA

E

NT

PD

5 1

#

-/- s

iR

NA

E

NT

PD

5 2

#

C

*

1 2 3 4 5 6 7 8 9 10Lane

Figure 2. Purification and Characterization of ENTPD5

(A) Diagram of the purification scheme for ATP hydrolysis activity from S-100 of PTEN�/� MEF cells.

(B) Final step of purification. (Top) Aliquots of 30 ml of indicated fractions from the Mini Q column were subjected to 4%–10% gradient SDS-PAGE gels followed by

staining using a silver staining kit from Invitrogen. (Bottom) Aliquots of 3 ml of indicated fractions were incubated with 10.5 ml of undialyzed S-100 from PTEN+/�

MEF cells and assayed for ATP hydrolysis activity.

(C) (Lanes 1–5) ATP hydrolysis activity in S-100 from PTEN+/� MEF cells expressing exogenous ENTPD5. PTEN+/� vector or PTEN+/� ENTPD5 1# and 2# (two

individual clones with different expression levels of ENTPD5) were established as described in the Experimental Procedures. Cell lysates (S-100) from indicated

cell lines were prepared, and aliquots of 30 mg were used for ATP hydrolysis assay. (Lanes 6–10) ENTPD5 expression in PTEN�/� MEFs was knocked down as

described in the Experimental Procedures. The cells were harvested, and S-100 were prepared and normalized for ATP hydrolysis assay. Positions of ATP, ADP,

and AMP are indicated. (Bottom) Aliquots of 10 mg protein of indicated samples were subjected to 10% SDS-PAGE followed by western analysis of ENTPD5.

Asterisk denotes cross-reactive proteins.

(D) PTEN�/� MEF cells were transfected with 4 mg plasmid DNA containing vector control or cDNA encoding PTEN or PTENcs as indicated. At 24 hr after trans-

fection, cells were harvested and total cell lysates were prepared. Aliquots of 10 mg of protein were loaded onto 10% SDS-PAGE followed by western analysis of

levels of PTEN, AKT, phosphorylated AKT(pAKT), ENTPD5, and b-actin as indicated.

(E) PTEN+/� and PTEN�/� MEF cells were treated with DMSO or LY294002 (50 mM) for 24 hr. Aliquots of 20 mg total cell extracts were subjected to 10% SDS-

PAGE followed by western analysis using indicated antibodies.

See also Figure S1 and Figure S2.


which could not be hydrolyzed to UMP, indicating that the

conversion to UMP is critical for this cofactor to function. The

same holds true for guanine nucleotides (Figure S3).

UMP, ENTPD5, UMP/CMP Kinase-1, and AdenylateKinase-1 Constitute an ATP-to-AMP Hydrolysis CycleBased on the facts that purified ENTPD5 is unable to hydrolyze

ATP directly and the assay also contained S-100 from PTEN

heterozygous MEFs, we realized that there must be more factors

in the S-100, which are also required to hydrolyze ATP to AMP.

These factors presented in cells regardless of their PTEN status.

For example, when we added purified, recombinant ENTPD5

and UMP to the dialyzed S-100 from large-scale cultured HeLa

cells, the ATP-to-AMP hydrolysis was reconstituted (Figure 4A,

lanes 1–6). This observation made purification of these factors

easier because HeLa cells can be grown in large quantity in

Lane

+/- +/-

1 2 3 4 5 6 7 8 9 10

+/- +/- +/--/- -/- -/- -/- -/-

Dialysis

S-100

Small Molecule from +/- +/--/- -/- Hela

AMP

ADP

ATP

B

1000

250

62.5

15.625

4 1000

250

62.5

15.625

4 1000

250

62.5

15.625

4

Tri-phosphate Di-phosphate Mono-phosphate

Conc.(μM)

Uracil

Guanine

Cytosine

Thymidine

AMP

AMP

AMP

AMP

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15Lane

C

Lane 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16

Small Molecule - UMP UTPαSUTPγS PNP-UMP

10

00

25

0

62

.5

16

4 10

00

25

0

62

.5

16

4 10

00

25

0

62

.5

16

4 10

00

25

0

62

.5

16

4Conc.(μM)

AMP

ADP

ATP

D

17 18 19 20 21

Substrates

16

12

8

4

0No

rm

ali

ze

d [

Pi]

Fo

ld I

nc

re

as

e

Reaction 0hr

Reaction 2hr

ATP ADP CTP CDP GTP GDP UTP UDP

A

Figure 3. Small Molecule Requirement for ENTPD5-Mediated ATP Hydrolysis

(A) (Right) the recombinant human ENTPD5 was generated and purified as described in the Experimental Procedures. An aliquot of 120 ng recombinant ENTPD5

was subjected to SDS-PAGE followed by Coomassie brilliant blue staining. Arrows indicate recombinant ENTPD5. (Left) The nucleotide hydrolysis reactions were

carried out in triplicate by mixing 0.1 mg/ml ENTPD5 with 50 mM indicated nucleotides. After 2 hr incubation at 30�C, released free phosphate was measured by

malachite green assay as described in the Experimental Procedures. Data shown are representative of three independent experiments. Error bars indicate SEM.

(B) Small molecule (<10 kDa) was extracted from either PTEN+/� or PTEN�/� MEF cells or HeLa S3 cell lysates (S-100 fractions) as described in the Experimental

Procedures. Aliquots of 10.5 ml undialyzed cell lysates (lane 1 and 2) or dialyzed cell lysates (lane 3–10) fromPTEN+/� (lanes 1, 3, 5, 7, and 9) orPTEN�/� (lanes 2, 4,

6, 8, 10) MEFs (3.5 mg/ml) were mixed with another 10.5 ml buffer A (lane 1 to 4) or small molecule recovered from PTEN+/� (lanes 5 and 8), PTEN�/� cells (lanes 6

and 7), or from HeLa S3 cells (lanes 9 and 10) and were assayed for ATP hydrolysis activity. The positions of ATP, ADP, and AMP were indicated.

(C) Aliquots of 10.5 ml dialyzed S-100 from PTEN�/� MEF cells (3.5 mg/ml) were incubated in the presence of indicated final concentration of UTP, UDP, and UMP;

or GTP, GDP, and GMP; or CTP, CDP, and CMP; or TTP, TDP, and TMP as indicated at 30�C with a-P32-labeled ATP in a total volume of 30 ml at 30�C for 1 hr

followed by TLC to resolve radioactive adenosine nucleotides. Position of AMP on TLC plate is indicated.

(D) Aliquots of dialyzed S-100 prepared from PTEN�/� MEF cells were mixed with buffer A (lane 1) or indicated final concentration of indicated nucleotides and

assayed for ATP hydrolysis activity.

See also Figure S3.


suspension. To identify these factors, we fractionated HeLa cell

S-100, using a Q Sepharose column, and collected both the

flowthrough (Q-FL) and column-bound fractions eluted with

300 mM NaCl (Q-30). Neither fraction alone was able to hydro-

lyze ATP to AMP, although the Q-30 fraction, when ENTPD5

and UMP were present, hydrolyzed ATP to ADP (Figure 4A, lanes

13 and 14). When both the Q-FL and Q-30 fractions were

included, the ATP-to-AMP activity was fully reconstituted (Fig-

ure 4A, lane 18).

We purified the activity present in the Q-30 fraction. The

activity present in the Q-30 fraction was purified by subjecting

HeLa S-100 onto four sequential column chromatographic steps

and finally onto a Mini Q column (Figure 4B, left). The activity

was eluted from this column with a linear salt gradient from 40

to 120 mM NaCl, and fractions eluted from the column were as-

sayed in the presence of recombinant ENTPD5, UMP, and the

Q-FL fraction (Figure 4B, right-bottom). A peak of activity was

observed at fractions 8–10. The same fractions were subjected

to SDS-PAGE followed by silver staining, and two protein bands

close to 37 and 20 kDa markers correlated perfectly with activity

(Figure 4B, right-top). Both bands were identified by mass

spectrometry as human UMP/CMP kinase-1 (CMPK1).

Fraction № 4 5 6 7 8 9 10 11 12

40 mM

120 mM NaCl

AMP

ADP

ATP

20KD

25KD

37KD

50KD

75KD

Start Hela S-100

Q HP

SP HP

Heparin HP

Super-dex 200

Mini Q

B

Fraction № ST 13 14 15 16 17 18 19 2220 21 23 24 25

AMP

ADP

ATP

AK1

C

rENTPD5 + -+ + - + - + - + - + - + - + - +

UMP - + - - + + - - + + - - + + - - + +

Hela S-100 Q-FL Q-30 Q-FL + Q-30

Lane 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17

A

AMP

ADP

ATP

18

AMP

ADP

ATP

Lane 1 2 3 4 5 6 7 8

PN

Gase F

AK

1

CM

PK

1

ENTPD5

9 10 11 12

Glycosylated

ENTPD5

UnGlycosylated

ENTPD5

PNGaseF

AK1

CMPK1

ENTPD5 +- - ++ - + +

AK1 + - - + + - + +

CMPK1 +- - + +- + +

UMP + + + + + -+ +

D

N/T

Figure 4. Reconstitution of ENTPD5-Mediated ATP Hydrolysis

(A) HeLa S3 cell S-100 was fractionated by Q Sepharose column to two fractions (Q-FL [flowthrough] and Q-30). Q-30 represents fraction eluted with 300 mM

NaCl. Aliquots of 15 ml buffer A (lane 1 and 2), or dialyzed HeLa cell S-100 (lanes 3–6), or Q-FL (lanes 7–10), or Q-30 (lanes 11–14), or Q-FL combined with Q-30

(7.5 ml each) (lanes 15–18) were mixed with (lanes 1, 2, 4, 6, 8, 10, 12, 14, 16, and 18) or without (lanes 3, 5, 7, 9, 11, 13, 15, and 17) ENTPD5 in the presence (lanes 2,

5, 6, 9, 10, 13, 14, 17, and 18) or absence (lanes 1, 3, 4, 7, 8, 11, 12, 15, and 16) of 100 mM UMP and assayed for ATP hydrolysis activity.

(B) (Left) Diagram of the purification scheme for the required factor in Q-30. (Right) Final step of purification of CMPK1. (Top) Aliquots of 60 ml indicated Mini Q

fractions that were subjected to 4%–10% gradient SDS-PAGE followed by silver staining. Arrow indicates the protein band correlated with ATP hydrolysis

activity. (Bottom) Aliquots of 5 ml indicated fractions that were mixed with 15 ml of dialyzed Q-FL fraction in the presence of 100 mM UMP and 18 ng recombinant

ENTPD5 and were assayed for ATP hydrolysis activity.

(C) 3 ml of the Q-FL fraction was concentrated to 600 ml with a spin column and analyzed on a Supdex-200 column (10/30). Fractions of 1 ml were collected, and

aliquots of 7.5 ml of indicated fractions were combined with 7.5 ml dialyzed Q-30 fraction, 100 mM UMP, and 18 ng recombinant ENTPD5 and were assayed for ATP

hydrolysis activity. Positions of radioactive ATP, ADP, and AMP are indicated. (Bottom) Aliquots of 10 ml of indicated fractions were subjected to 10% SDS-PAGE

followed by western blotting analysis using an antibody against human adenylate kinase 1 (AK1).

(D) (Left) Aliquots of recombinant AK1 (lane 1), ENTPD5 (lane 2), and CMPK1 (lane 3) (final concentration, 1 mg/ml) were incubated alone or were sequentially

combined as indicated (lane 4 to 8) in the presence (lane 1 to 7) or absence (lane 8) of UMP (100 mM) for ATP hydrolysis activity. Position of ATP, ADP, or

AMP was indicated. (Right) Aliquots of 10 mg recombinant AK1 (lane 9), ENTPD5 (lane 10) or ENTPD5 pretreat with PNGase F (NEB) (50 units/mg ENTPD5)

(lane 11), and CMPK1 (lane 12) were subjected to 10% SDS-PAGE followed by Coomassie brilliant blue staining.


The identification of UMP/CMP kinase in the Q-30 fraction

shed light on why UMP is a cofactor for the ATPase activity

and how ENTPD5 plus this enzyme generates ADP from ATP.

In this reaction, UMP is phosphorylated into UDP by CMPK1

and ATP, generating ADP. UDP is subsequently hydrolyzed by

ENTPD5 to UMP, completing the cycle with net conversion of

ATP to ADP.

With this knowledge, we then made an educated guess that the

third protein factor present in the Q flowthrough fraction should

be an adenylate kinase, which converts two ADP into one ATP

and AMP, causing the ATP-to-AMP conversion seen in PTEN

null cell extracts. To confirm this, we took the Q flowthrough frac-

tion and subjected it to a gel-filtration column and collected the

fractions eluted from the column to assay for ATP-to-AMP hydro-

lysis in the presence of UMP, purified recombinant ENTPD5, and

the Q-30 fraction that contains CMPK1. An ATP-to-AMP activity

peak centered at fractions 17 and 18 was observed (Figure 4C,

top). When these factions were subjected to western blotting

analysis using an antibody against adenylate kinase-1 (AK1),

the detected western blotting band correlated perfectly with the

activity peak (Figure 4C, bottom). The correlation was maintained

with additional chromatographic steps (data not shown).

We subsequently generated recombinant CMPK1 and AK1 in

bacteria and purified them to homogeneity (Figure 4D, lanes 9

and 12). Purified recombinant ENTPD5 expressed in insect cells

runs as a triplet on an SDS-PAGE gel that could be shifted down

to a doublet after treatment by PNGase F, indicating that

ENTPD-5 is glycosylated (Figure 4D, lanes 10 and 11).

These purified recombinant proteins allowed us to reconsti-

tute the ATP-to-AMP hydrolysis cycle. Only when all three

enzymes and UMP were present, efficient ATP-to-AMP conver-

sion was observed (Figure 4D, lanes 1–8).

ENTPD5 Is an ER EnzymeAfter purification and identification of ENTPD5 from PTEN null

cells, we realized that ENTPD5 is identical to a previously purified

ER UDPase (Trombetta and Helenius, 1999). Although we iden-

tified and purified ENTPD5 from the S-100, the enzyme most

likely fractionated there as a result of broken ER from physical

shearing during the cell-breaking process. When we expressed

an ENTPD5-GFP fusion protein in cells, the GFP signal was co-

localized with the coexpressed ER-DsRed marker (Figure 5A).

The ER location of ENTPD5 and its preferred specificity for

UDP suggested that ENTPD5 functions in the process of reglu-

cosylation catalyzed by UGGT for calnexin/calreticulin-mediated

protein folding (Trombetta and Parodi, 2003). In the process,

UDP is generated after the conjugated glucose gets transferred

to the glycosidase I/II trimmed core glycan on N-glycosylated

proteins. UDP-glucose is made in cytosol and transported into

ER through the UDP-sugar transporter, which is an antiporter

that must exchange out one molecule of UMP for each UDP

sugar conjugate imported into the ER (Hirschberg et al., 1998).

UDP therefore needs to be hydrolyzed to UMP to prevent end

product feedback inhibition of UGGT, as well as to serve as

a substrate for the antiporter (Trombetta and Helenius, 1999).

UMP is phosphorylated back to UDP by CMPK1 in the cytosol,

and the generated ADP is converted to ATP and AMP by AK1

(diagramed in Figure 5B).

Knockdown of ENTPD5 Causes ER Stress and GrowthInhibitionBecause cells with an activated PI3K/AKT pathway increase

their cellular protein translation level, cells need to evolve a corre-

sponding system in ER to accommodate the high demand for

protein folding process. It is possible that cells may do so by up-

regulating ENTPD5 to increase the conversion of UDP to UMP in

ER, thereby promoting N-glycosylation and folding. Thus,

reducing the level of ENTPD5 in cells with active AKT should

induce ER stress. In addition, because many growth-promoting

cell membrane receptors are highly N-glycosylated, loss of

function of ENTPD5 could affect their folding process, resulting

in their reduction and, subsequently, cell growth arrest. To test

this hypothesis, we engineered several cell lines based on

the PTEN null MEFs in which the expression of ENTPD5 could

be knocked down with the addition of doxcycline (Dox),

which turned on a Tet-suppressor-controlled shRNA-targeting

ENTPD5. The results from a representative cell line were shown

in Figure 5C. Comparing to PTEN null MEFs expressing GFP

shRNA, addition of Dox to the culture media resulted in success-

ful knockdown of ENTPD5 expression in these cells. As a result,

an ER stress marker, GRP78/BiP, was induced, and cellular

N-glycosylation level, as measured by PHA blotting, was down

(Figure 5C, lanes 5–8). Of interest, the levels of receptor tyrosine

kinases, including EGFR, Her-2/Erb-2, and type I insulin-like

growth factor receptor (IGF-IR) b, were significantly decreased

after ENTPD5 knockdown.

To confirm that the above-mentioned cellular effects after

ENTPD5-targeting shRNA expression were specific, we intro-

duced into these cells a cDNA encoding ENTPD5 with silent

mutations in the shRNA target sequence. In these cells, although

the endogenous ENTPD5 was still knocked down after addition

of Dox (Figure 5D, lanes 2, 4, and 6), the expression of an

shRNA-resistant wild-type transgene (three flag tags were fused

to ENTPD5 coding sequence so it migrated higher) led to

complete reversal of BiP induction, lowered glycosylation, and

downregulation of these growth factor receptors (Figure 5D,

lane 4). In contrast, introducing an E171A mutant that abolishes

UDP hydrolysis activity of ENTPD5 was not able to rescue these

phenotypes (Figure 5D, lane 6). In addition to BiP, another ER

stress marker, CHOP, was also induced when ENTPD5 was

knocked down (Figure 5D).

Consistent with the loss of growth factor receptors after

ENTPD5 knockdown, cell growth was also dramatically attenu-

ated. As shown in Figure 5E, when ENTPD5 in PTEN null MEFs

was knocked down after addition of Dox, very few colonies

grew on the culture dish after 10 days, although the same

number of cells was plated initially, and they were cultured under

the same condition (Figure 5E, left row). The growth inhibition

was rescued when the shRNA-resistant ENTPD5 cDNA was ex-

pressed (Figure 5E, middle row), whereas the inhibition was

exacerbated if an enzymatic dead mutant of ENTPD5 was ex-

pressed instead (Figure 5E, right row).

ENTPD5 Promotes Aerobic GlycolysisOne implication of elevated ENTPD5 expression is that a

significant percentage of cellular ATP is consumed through

the ENTPD5/CMPK1/AK1 enzyme cascade. To maintain the


ENTPD5-GFP

ENTPD5-GFP

DsRed

ER-DsRed Merge

Mer ge

A

N-acetylglucosamine

manose UUU uridine

glucose PPP phosphate

2xATP 2xADP AMP

2xUMP 2xUDP

ENTPD5

CMPK1

AK11/2 1/2

2xPi

CytosolER Lumen

ATP 2xAMP

+PiInput Output

B

UP UUP

UGGT FoldingGlycoprotein

Cytosol

ER Lumen

UDP-GlucoseUMP

Phosphate

PhosphateTransporter

Antiporter

ENTPD5

UP UUP

UP UUPi

PTEN-/- MEF shRNA ENTPD5Rescue Vec sr ENTPD5 sr ENTPD5 E171A

Cell line

- Dox+ Dox

E

Lane 1 2 43 5 6 7 8Dox +- + + +- - -

Time 2d 4d 4d2d

Cell LineGFP ENTPD5PTEN-/- sh RNA

ENTPD5

GRP78/BiP

EGFR

ß-Actin

*

PH

A B

lot

Her-2/ErbB-2

IGFRß

C

PTEN-/- MEFsh RNA ENTPD5

Vec

sr E

NT

PD

5

sr E

NT

PD

5 E

171A

Lane 1 2 3 54 6Dox - - -+ + +

Cell Line

ENTPD5sr ENTPD5-3Flag

GRP78/BiP

*

EGFR

ß-Actin

PH

A B

lot

Rescue

Her-2/ErbB-2

IGFRß

D

CHOP

Figure 5. Biological Function of ENTPD5 in PTEN�/� MEF Cells

(A) PTEN�/� MEF cells were cotransfected with mouse ENTPD5-GFP and free DsRed or with ENTPD5-GFP and ER-localized DsRed (ER-DsRed). ENTPD5-GFP

colocalized with ER-DsRed (bottom row), but no obvious codistribution with free DsRed was observed (top row). Scale bars, 10 mm.

(B) Working model for ENTPD5. See the text for details.

(C) PTEN�/� MEF cells with doxycycline (Dox)-inducible expression of shRNA-targeting ENTPD5 was generated as described in the Experimental Procedures.

After 2 or 4 days induction with Dox (0.125 mg/ml), cells were harvested and total cell lysates were prepared as described in the Extended Experimental Proce-

dures. Aliquots of 10 mg protein were subjected to SDS-PAGE followed by western blotting analysis using the indicated antibodies. Glycosylation was visualized

by PHA blot as indicated. Asterisk denotes decreased glycosylated proteins.


intracellular ATP level, the extra ATP consumed should come

from either increased oxidative phosphorylation or glycolysis.

We therefore tested both by measuring oxygen consumption

and lactate production, respectively. Consistent with previous

reports, there was not much difference in the respiration rate of

PTEN null and PTEN heterozygous MEFs (Figure S4), but

PTEN null cells showed �40% higher lactate production in their

cultured medium (Figure 6A, columns 4 and 5). When ENTPD5

was ectopically expressed in two PTEN heterozygous MEF lines,

lactate production was increased, and the level of increase

correlated with that of ENTPD5 expression (Figure 6A, columns

2 and 3). Consistently, when ENTPD5 was knocked down with

addition of Dox as in Figure 5D, lactate production was signifi-

cantly decreased (Figure 6B, columns 1 and 2). In PTEN null

MEFs harboring Dox-inducible ENTPD5-targeting shRNA that

also expressed shRNA-resistant ENTPD5 transgene, addition

of Dox did not result in a decrease in lactate production, and

the basal lactate production also became higher, correlated

with the higher than endogenous expression of the transgene

(Figure 6B, columns 3 and 4, and Figure 5D). In contrast, when

the catalytic site mutant ENTPD5 transgene was expressed to

the similar level, lactate production still reduced with the addition

of Dox (Figure 6B, columns 5 and 6).

If the ATP hydrolysis cycle initiated by ENTPD5 discovered

in vitro is operational in cells, and the extra ATP consumed by

a higher level of ENTPD5 is compensated by increased glycol-

ysis, glucose starvation of these cells should result in much

faster decrease of intracellular ATP level compared to cells

with lower ENTPD5 expression. Indeed, when intracellular ATP

concentrations in PTEN heterozygous and PTEN null MEFs

were measured after glucose withdrawal from the culture media,

that in PTEN null MEFs decreased to about half of the original

level within the first hour, while there was little change of ATP

in PTEN-heterozygous MEF within 2 hr (Figure 6C).

To confirm that the faster ATP level dropping in PTEN null cells

was due to higher expression of ENTPD5, the same set of MEFs

as in Figure 6B was subjected to glucose starvation after

ENTPD5 was knocked down with the addition of Dox. Knock-

down of ENTPD5 for 2 days in PTEN null MEFS caused the total

cellular ATP level to decrease (Figure 6D, columns 3 and 4).

However, the ATP level did not decrease further after glucose

starvation for 1 hr, whereas cells without Dox treatment

consumed 50% of original ATP during this period (Figure 6D,

columns 1 and 2). The MEFs expressing the shRNA-resistant

ENTPD5 did not lower their ATP level with Dox treatment, but

their ATP levels were even more drastically lowered after glucose

starvation, possibly due to ENTPD5 overexpression (Figure 6D,

columns 5–8). In contrast, cells expressing a similar level of the

catalytic dead mutant ENTPD5 behaved the same as cells

without transgene expression (Figure 6D, columns 9–12).

The observed decrease of glycolysis after ENTPD5 knock-

down could be due to lowered tyrosine kinases receptors and

AKT activity (Figure 5C and Figure S5), which stimulates the

glucose transporter activity on cell surface (Kohn et al., 1996;

Plas et al., 2001). In addition, knockdown of ENTPD5 may reduce

the production of ADP/AMP, which allosterically activate glycol-

ysis enzymes such as phosphofructose kinase (PFK) (Gevers

and Krebs, 1966). To distinguish these possibilities, cellular

fructose-6-phosphate and fructose-1,6-bisphosphate were

measured using LC-Mass. As shown in Figure 6E, the former

was lowered by �20% after ENTPD5 knockdown (Figure 6E,

left), whereas the latter dropped by �60% (Figure 6E, right).

These results suggested that ENTPD5 indeed affects glucose

influx to cells, but its major impact on glycolysis is to directly

activate glycolysis enzymes such as PFK by hydrolyzing ATP.

ENTPD5 Expression Correlates with AKT Activation inHuman Cancer Cell Lines and Primary Tumor SamplesPTEN mutation and AKT activation are common features for

human cancer. To check whether what was observed in PTEN

null MEFs is also true for human cancer cells, we screened

a panel of human cancer cell lines for the expression of PTEN,

activated AKT, and ENTPD5. As shown in Figure 7A, AKT activa-

tion was seen in human prostate cancer lines C42 and LNCaP

cells, and in these two cell lines, elevated ENTPD5 expression

was also observed.

We also examined ENTPD5 expression and AKT activation in

primary human tumor samples by staining two adjacent sections

from a formalin-fixed, paraffin-embedded human primary pros-

tate cancer sample with rabbit monoclonal antibodies against

human ENTPD5 and phosphoAKT, respectively. The specificity

of this anti-ENTPD5 antibody was verified by western blotting

analysis using LNCaP cell lines with or without their ENTPD5

knocked down (Figure S6A). The staining intensity for ENTPD5

in tumor was significantly greater compared with adjacent normal

tissue and was correlated with pAKT staining (Figure 7B). Out of

10 samples from patients between age 57 and 76, only one tumor

sample from a 57-year-old patient and another sample collected

from a patient who had just gone through therapy did not show

strong ENTPD5 staining, and the same tumors were also negative

for pAKT (Figure S6B2 and Figure S6B10). The remaining eight

samples all showed greater tumor staining of pAKT and ENTPD5

(Figure 7B and Figures S6B3–S6B9).

Because microarray data of many tumors are publicly avail-

able, we also analyzed a group of recently publicized microarray

data from human prostate cancer samples (Bermudo et al.,

2008). We found that ENTPD5 is highly expressed in all 20 tumor

samples compared to normal prostate epithelium cells (Fig-

ure S7). In addition, after clustering all gene expression profiles

from prostate tumor microarray data using SOM (self-organiza-

tion method), we identified dozens of genes associated with

AKT activation, including Her-3, PI3KCB, Ras, S6 kinase,

CD36, IL8, EGF, osteropontin, and FoxO1, which are signifi-

cantly coregulated with ENTPD5 (Figure S7).

(D) Rescue cell lines with expression of shRNA-resistant wild-type or catalytic dead mutant (E171A) ENTPD-5 were established as described in the Extended

Experiments Procedures. Same as in (C), after 2 days culture, cells were harvested, and total cell lysates (10 mg/lane) were subjected to SDS-PAGE followed

by western analysis as indicated. Glycosylation was visualized by PHA blot analysis.

(E) Rescue cell line was plated at density of 5 3 104/100 mm dish and treated with Dox as in (C). Cell medium was changed each 3 days. After 10 days culture, the

plates were stained by methylene blue.


ENTPD5 Is Important for Cancer Cell GrowthTo verify the functional significance of ENTPD5 expression in

human cancer cells, we generated a cell line from the original

LNCaP cells in which an shRNA against human ENTPD5 could

be induced by Dox. In these cells, knockdown of ENTPD5 by

adding Dox to the culture media also lowered N-glycosylation

(Figure 7C, comparing lanes 7 and 8, 9 and 10, and 11 and 12)

and induced BiP expression (Figure 7C, lanes 8, 10, and 12).

B

lact

ate

(nm

ol/m

g pr

otei

n)

400

800

1200

0

Dox - + - -+ +

Cell Line PTEN-/- MEF shRNA ENTPD5

Rescue Vec sr ENTPD5 sr ENTPD5E171A

PTEN+/- PTEN-/-Cell Line

- Glucose 0 1 h 2 h 0 1 h 2 h

ATP

(nm

ol/m

g pr

otei

n)

C

0

2

4

6

8

10

0

0.2

0.4

0.6

0.8

1.0

1.2

Rel

ativ

e Fr

ucto

se-6

-P C

onte

nt

PTEN-/- MEFshRNA ENTPD5

- +Dox

0

0.2

0.4

0.6

0.8

1.0

1.2

Rel

ativ

e Fr

ucto

se-1

, 6-2

P C

onte

nt

PTEN-/- MEFshRNA ENTPD5

- +Dox

EF-6-P F-1,6-2P

Clone № 1# 2#

Stable transfected with Vec ENTPD5-Flag

PTEN+/- -/-Cell Line

lact

ate

(nm

ol/m

g pr

otei

n)400

200

600

800

0

ENTPD5ENTPD5 Flag

ß-Actin

A

- Glucose

Rescue

Dox

Cell Line

D

0

2

4

6

8

10

12AT

P (n

mol

/mg

prot

ein)

ENTPD5sr ENTPD5 3Flag

ß-Actin

- - - - - -+ + + + + +

sr ENTPD5 sr ENTPD5 E171A

- - -+ + +

Vec

PTEN-/- MEF shRNA ENTPD5

Figure 6. ENTPD5 Promotes ATP Hydrolysis and Glycolysis In Vivo

(A) Lactate in the culture media of PTEN+/� and PTEN�/� MEF cells (columns 4 and 5) as well as PTEN+/� MEF clones stably transfected with vector control

(column 1) or Flag-tagged mouse ENTPD5 (column 2, clone 1 and column 3, clone 2, as used in Figure 2C) was measured as described in the Extended

Experimental Procedures, and the value was normalized to total protein amount.

(B) Lactate in the culture media of ENTPD5 knockdown and rescue cell lines was measured as in (A) except pretreatment with or without Dox for 2 days.

(C)PTEN+/� andPTEN�/� MEF were deprived of glucose for indicated time periods, and intracellular ATP was determined as described in Extended Experimental

procedures.

(D) The intracellular ATP was measured 1 hr after glucose starvation on ENTPD5 knockdown and rescue cell lines as in (B).

(E) ENTPD5 was knocked down as in (B), and the intracellular fructose-6-phosphate (left) and fructose-1,6-bisphosphate (right) were separated and quantified by

HPLC mass spectrometry (ABI 3200 Q TRAP LC/MS/MS Systems). The relative amount of metabolite is normalized to total ion count (TIC). All experiments were

repeated at least two times, and the error bar represents standard deviation.



These phenotypes were rescued by the expression of a wild-

type shRNA-resistant ENTPD5 transgene, but not by the active

site mutant ENTPD5 (Figure 7C, lanes 13–24). Several growth

factor receptors were also checked in these cell lines after Dox

treatment. As shown in Figure 7D, EGFR and Her2/ErbB-2

were significantly down, and IGFRbwas slightly down (Figure 7D,

lanes 1–4). They were restored to the normal level by the expres-

sion of shRNA-resistant ENTPD5 transgene (Figure 7D, lanes 5

and 6), but not the active site mutant (Figure 7D, lanes 7 and

8). Consistently, when the cell number was measured after

4 day knockdown of ENTPD5, only about half of LNCaP cells

were there, compared to a control knockdown cell line, and

the defect was rescued by expression of wild-type ENTPD5

transgene, but not active site mutant (Figure 7E).

To test whether knocking down ENTPD5 in LNCaP cells also

has an effect on their growth in vivo, we implanted the LNCaP

cells bearing a Dox-inducible shRNA targeting ENTPD5 in matri-

gel in nude mice. As a control, LNCaP cells with a Dox-inducible

shRNA targeting GFP were also implanted. After the xenograft

tumors reached the size of 500 mm3, a cohort of seven mice

were fed with Dox-containing water. The level of ENTPD5 in

these tumors was measured after 6 weeks. Compared with

mice fed with normal water, the ENTPD5 levels in ENTPD5-tar-

geting shRNA containing tumors from mice fed with Dox-con-

taining water were significantly lower except in one mouse

(Figure 7F). Whereas ENTPD5-targeting shRNA containing

tumors in mice fed with normal water continued to grow, the

tumors in mice fed with Dox-containing water shrank (Figure 7G).

Amazingly, when these tumor samples were analyzed under

a microscope after fixing and staining with hematoxylin and

eosin, there were very few tumor cells left in the matrigel in

tumors grown in Dox-fed mice, whereas in mice fed with normal

water, the matrigel was filled with tumor cells (Figure 7H). The

GFP shRNA-containing tumors did not respond to Dox treatment

and continued to grow during the period of experiment.

DISCUSSION

ENTPD5 Is an Important Link in the PI3K/PTENSignaling LoopThe experimental data reported here identify ENTPD5, an ER

UDPase, as an important link in the PI3K/PTEN/AKT signaling

loop. We reason that ENTPD5 upregulation is important for

AKT-activated cells to cope with elevated translational activity

that generates more nascent polypeptide chains destined for

the ER.

ENTPD5 is a member of the ectonucleoside triphosphate

diphosphohydrolase family, which consists of seven other

members (reviewed by Robson et al., 2006). ENTPD 1–3

(CD39, CD39L1, and CD39L3) are typical ectoenzymes, whereas

the other five members have a predominant intracellular localiza-

tion including ER, Golgi apparatus, and lysosomal/auto-

phagic vacuoles. The functions of these organelle-associated

ENTPDases are still largely unexplored, but judging by their loca-

tion and substrate preference, it would not be surprising if they all

turn out to regulate protein glycosylation.

Among members of this group of enzymes, however, ENTPD5

is the only intracellular ENTPDase that is transcriptionally upre-

gulated in PTEN null cells (Figure S1). The mRNA of an extracel-

luar ENTPDase, ENTPD2, although expressed at a much lower

level than ENTPD5, was also elevated in PTEN null cells (Fig-

ure S1). The significance of such is unknown.

ENTPD5 Contributes to Warburg EffectOne of the surprising findings reported here is how quickly ATP

can be consumed as a result of ENTPD5 upregulation. One

naturally raised question is where the extra consumed ATP

comes from. After measuring both oxygen consumption and

lactate generation, we found that the lactate production was

elevated in these PTEN null cells, whereas oxygen consumption

did not change (Figure 6A and Figure S4). When ENTPD5 was

knocked down, higher lactate production returned to normal

(Figure 6B). Moreover, simply ectopically expressing ENTPD5

in PTEN heterozygous MEF elevated their lactate production

(Figure 6A). These results indicate that ENTPD5 is a critical player

in causing the Warburg effect, i.e., elevated lactate production

under aerobic conditions, in these PTEN null cells.

In addition to being part of the activation loop for AKT that

promotes glucose uptake into cells (Kohn et al., 1996; Plas

et al., 2001), a major effect of ENTPD5 on glycolysis might be

its ability to generate ADP/AMP through the aid of CMPK1 and

AK1. Elevated AMP levels (and to a lesser extent, ADP) activate

phosphofructokinase and inhibit fructose diphosphatase to drive

glycolysis and prevent gluconeogenesis, resulting in higher

lactate production (Gevers and Krebs, 1966). Consistently, the

fructose-1,6-bisphosphate level dropped to a much lower level

than that of fructose-6-phosphate when the ENTPD5 in PTEN

null cells was knocked down (Figure 6E).

In addition to UDP, ENTPD5 also use GDP as a substrate and

hydrolyze it to GMP (Figure 3A and Figure S3). It is interesting to

note that GDP-conjugated sugars are another group of major

substrates for glycosylation. The significance of hydrolyzing

GDP by ENTPD5 is not clear because it is believed that GDP

sugars are transferred to proteins in the Golgi.

ENTPD5 Is Potentially an Anticancer TargetThe current study highlighted ENTPD5 as a critical link in the

PI3K/PTEN pathway that promotes cell growth and survival,

a pathway that is often activated in cancer cells. We saw good

correlation between ENTPD5 expression and AKT activation in

both cultured prostate cancer cell lines and primary human pros-

tate carcinoma samples (Figures 7A and 7B and Figures S6 and

S7). Therefore, inhibition of this enzyme, similar to knockdown,

can potentially generate benefits for anticancer activity. It should

induce more severe ER stress in cancer cells with active AKT due

to higher protein traffic through the secretory pathway. It may

cause synthetic lethality in these cells, which otherwise maintain

survival advantage and resistance to common anticancer drugs.

It will also lower many growth factor receptors on the cell surface

due to their high N-glycosylation nature, a phenomenon that may

reflect the evolutionary connection between fast growth and

nutrient availability in mammalian cells (Lau et al., 2007). Among

such receptors, EGFR, Her2/ErB2, and IGFR levels were down

after ENTPD5 knockdown (Figure 5 and Figure 7), whereas

a nongrowth-promoting TGFb receptor did not change (data

not shown).


G

LNCaPshRNA GFP -DoxLNCaPshRNA GFP +DoxLNCaPshRNA ENTPD5 -DoxLNCaPshRNA ENTPD5 +Dox

1 2 3 4 5 6Weeks after Dox

Tuno

r si

ze c

hang

e af

ter

Dox

(%)

50%

70%

90%

110%

130%

150%

170%

190%

*

B

a b

c d

ENTPD5 pAKT

LNCaP shRNA GFP ENTPD5

Lane 1 2 3 4 5 6 7 8Dox - + - + - + - +

Rescue

No

ne

sr

EN

TP

D5

sr

EN

TP

D5

E1

72

A

0

20

40

60

80

100

120

E

Cel

l gro

wth

inhi

bitio

n (%

)

- DOX - DOX

+ DOX + DOX

H

500um 100um

LNC

aP s

h R

NA

EN

TPD

5 tu

mor

s

DU

145

LAPC

4

PC3

LNC

aPC

42

MC

F-7

MB

A23

1SK

BR

-3

T47-

D

Prostate Breast

PTEN

pAKT

ENTPD5

AKT

ß-Actin

A

GRP78/BiP

Lane 1 2 3 54 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24Dox - - -+ + + - - -+ + + - - -+ + + - - -+ + +

Time 2d 4d 6d 2d 4d 6d 2d 4d 6d 2d 4d 6dLNCaP shRNA GFP LNCaP shRNA ENTPD5

Rescue None sr ENTPD5 sr ENTPD5 E172A


PH

A B

lot

*

Cell Line

C

ß-Actin

D

Dox - - -+ + +

sr E

NTP

D5

sr E

NTP

D5

E17

2ANon

e

- +Lane 1 2 3 4 5 6 7 8

Cell Line (LNCaP shRNA) GFP ENTPD5


EGFR

ß-Actin

Rescue

Her-2/ErbB-2

IGFRß

ENTPD5

Ponceau S

-Dox +Dox

LNCaP shRNA ENTPD5 tumorsF

Figure 7. Knockdown of ENTPD5 in LNCaP Cells Decreases Glycosylation, Expression of Cell Surface Receptors, and Tumor Progression

(A) Aliquots of 20 mg of total cell extracts from indicated cell lines were subjected to 10% SDS-PAGE followed by western blotting analysis using antibodies as

indicated.

(B) Immunohistochemical staining for ENTPD5 (a and c) and pAKT (b and d) in adjacent sections of a human prostatic carcinoma sample (a/b 103 and c/d 203

lenses). Scale bar, 200 mM (a and b); 100 mM (c and d). Arrows indicate tumor.

(C) Inducible ENTPD5 knockdown and rescue stable cell lines were treated with or without Dox (0.0625 mg/ml) for indicated time periods. Aliquots of 10 mg of total

cell extracts were subjected to 10% SDS-PAGE for western blotting analysis of ENTPD5, BiP, glycosylated proteins (PHA blot), and b-actin. Asterisk indicates

decreased glycosylated proteins.

(D) Indicated cell lines were plated and treated with or without Dox for 6 days, total cell lysates were prepared, and aliquots of 10 mg protein were subjected to

SDS-PAGE followed by western blotting analysis using indicated antibodies.

(E) Indicated cells (1 3 104) were seeded in 96-well plates and then treated with and without Dox (1 mg/ml) for 4 days. Cell contents were measured.

(F–H) 2 3 106 LNCaP cells with Dox-inducible shRNA target GFP or ENTPD5 were injected subcutaneously into the flank of nude mice as described in the

Extended Experimental Procedures. When the tumors reached a volume of �500 mm3, mice were fed with normal or Dox-containing water.


Chronic inhibition of ENTPD5 may cause liver and male fertility

defects because mice with ENTPD5 deficiency show hepatop-

athy and aspermia (Read et al., 2009). These defects in mice,

however, only become obvious after 1 year of age. Given the

poor prognosis of PI3K/PTEN mutations in human cancers and

potential synthetic lethal effect of AKT activation and ENTPD5

inhibition, developing ENTPD5 inhibitors for cancer therapy

may be a worthwhile pursuit.

EXPERIMENTAL PROCEDURES

General Reagents and Methods

General chemicals are from Sigma unless otherwise described. We obtained

a-P32-labeled ATP from GE Healthcare. All other nucleotides are from Sigma.

Nonhydrolyzable uracil and guanine nucleotide analogs are from Gena Biosci-

ence (Germany). HRP-conjugated E-type PHA is from USBioLogical

(Ca#P3371-25). Puromycin, blasticidin, and hygromycin, which are used for

establishment and maintenance of stable cell lines, are purchased from Inviv-

ogen (Ca#ant-pr-1, ant-bl-1, and ant-hg-1, respectively). G418 is from Calbio-

chem (Ca#345810). The sources of antibodies used are listed in the Extended

Experimental Procedures.

Cell Culture

PTEN+/� and PTEN�/� MEF cells are established previously (Stambolic et al.,

1998). The sources of all other cell lines used and their culture conditions are

described in the Extended Experimental Procedures.

In Vitro ATP Hydrolysis Assay

The ATP hydrolysis assays were carried out by incubation-indicated cell

extracts or purified enzymes with a-P32-ATP and were analyzed by thin layer

chromatography (TLC). The detailed method was described in the Extended


Purification of ENTPD5 and CMPK

All purification steps were carried out at 4�C. All chromatography steps were

carried out using an automatic fast protein liquid chromatography (FPLC)

station (Pharmacia). The details of purification methods were described in

the Extended Experimental Procedures.

ENTPD5 Expression and ENTPD5 shRNA Constructs

All ENTPD5 expression and shRNA constructs were made as described in the

Extended Experimental Procedures.

Preparation of Recombinant ENTPD5, CMPK1, and Adenylate

Kinase

Human ENTPD5 recombinant protein was generated using Bac-to-Bac Bacu-

lovirus Expression Systems (Invitrogen Cat# 10359-016). Human CMPK1 and

AK1 were generated by bacterial expression. The details of methods were

described in the Extended Experimental Procedures.

Measurement of Lactate Production in Cell Culture Medium

We purchased Lactate Assay kit from Biovision (Cat#K627-100). Measure-

ment of Lactate concentration in cell culture medium was performed accord-

ing the manufacturer’s instruction and described in detail in the Extended


Measurement of Intracellular Fructose-6-P and Fructose-1,6-2P

The preparation and measurement of these two phosphosugars by LC-Mass

were described in the Extended Experimental Procedures.

Cell Survival Assay

Cell survival analysis was performed using the Cell Titer-Glo Luminescent Cell

Viability Assay kit (Promega) following manufacturer’s instruction with minor

modification. In brief, 25 ml of Cell Titer-Glo reagent was added to the cell

culture medium. Cells were placed on a shaker for 10 min and were then

incubated at room temperature for an additional 10 min. Luminescent reading

was carried on a Tecan SPECTRAFluor Plus reader (Tecan).

SUPPLEMENTAL INFORMATION

Supplemental Information includes Extended Experimental Procedures, seven

figures, and one table and can be found with this article online at doi:10.1016/j.

cell.2010.10.010.

ACKNOWLEDGMENTS

We would like to express our gratitude to Drs. Fenghe Du and Liping Liu for

excellent technical assistance. We are grateful for Dr. Aijun Liu from the 301

Hospital in Beijing for providing the human prostate tumor samples and

Dr. Benjamin Tu from University of Texas Southwestern for help with the phos-

phofructose sugar measurement. We thank Mr. Gregory Kunkel and Dr. Lai

Wang for critical reading of the manuscript. This work is also supported by

a grant from the National Cancer Institute (NCI) (PO1 CA 95471) and the

National High Technology Projects 863 from Chinese Ministry of Science

and Technology.

Received: May 14, 2010

Revised: September 10, 2010

Accepted: October 7, 2010

Published online: November 11, 2010

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(F) Tumors lysates were extracted after 6 weeks, and aliquots of 20 mg of protein were subjected to 10% SDS-PAGE and transfer to nitrocellulose filter. The filter

was stained with Ponceau S staining (bottom) followed by western blotting analysis using an antibody against ENTPD5 (top).

(G) Time course of tumor shrinkage caused by ENTPD5 knocking down. Tumor size was measured, and statistic analysis was performed as described in the

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Stepwise Histone Replacement by SWR1Requires Dual Activation with HistoneH2A.Z and Canonical NucleosomeEd Luk,1,* Anand Ranjan,1 Peter C. FitzGerald,2 Gaku Mizuguchi,1 Yingzi Huang,1 Debbie Wei,1 and Carl Wu1,*1Laboratory of Biochemistry and Molecular Biology2Genome Analysis Unit

National Cancer Institute, National Institutes of Health, Bethesda, MD 20892, USA

*Correspondence: [email protected] (E.L.), [email protected] (C.W.)DOI 10.1016/j.cell.2010.10.019

SUMMARY

Histone variant H2A.Z-containing nucleosomes areincorporated at most eukaryotic promoters. Thisincorporation is mediated by the conserved SWR1complex, which replaces histone H2A in canonicalnucleosomes with H2A.Z in an ATP-dependentmanner. Here, we show that promoter-proximalnucleosomes are highly heterogeneous for H2A.Z inSaccharomyces cerevisiae, with substantial repre-sentation of nucleosomes containing one, two, orzero H2A.Z molecules. SWR1-catalyzed H2A.Zreplacement in vitro occurs in a stepwise and unidi-rectional fashion, one H2A.Z-H2B dimer at a time,producing heterotypic nucleosomes as intermedi-ates and homotypic H2A.Z nucleosomes as endproducts. The ATPase activity of SWR1 is specificallystimulated by H2A-containing nucleosomes withoutensuing histone H2A eviction. Remarkably, furtheraddition of free H2A.Z-H2B dimer leads to hyperstim-ulation of ATPase activity, eviction of nucleosomalH2A-H2B, and deposition of H2A.Z-H2B. Theseresults suggest that the combination of H2A-contain-ing nucleosome and free H2A.Z-H2B dimer acting asboth effector and substrate for SWR1 governs thespecificity and outcome of the replacement reaction.

INTRODUCTION

The eukaryotic genome is packaged into chromatin within the

cell nucleus. The fundamental packaging unit of chromatin is

the nucleosome, which consists of an octameric histone core

around which 147 base pairs (bp) of DNA are wrapped in �1.7

left-handed superhelical turns, plus linker DNA of variable length

between adjacent nucleosome core particles (Kornberg and

Lorch, 1999). The canonical nucleosome, containing two each

of the four main histones, H2A, H2B, H3, and H4, is representa-

tive of the bulk of chromatin in the cell nucleus. However, a minor

fraction of the nucleosome population is assembled from nonal-

lelic histone variants, which have an important role in major chro-

mosome activities of the cell, including transcription, DNA repli-

cation, and repair (Ausio, 2006).

The widely conserved histone H2A.Z variant shares 60%

sequence identity with the canonical H2A histone and plays

essential, nonredundant roles in higher eukaryotes (Guillemette

and Gaudreau, 2006). In yeasts, H2A.Z is not essential, but cells

exhibit slow growth (Carr et al., 1994; Santisteban et al., 2000),

chromosome instability (Carr et al., 1994; Krogan et al., 2004),

gene silencing defects (Meneghini et al., 2003), and sensitivity

to genotoxic and environmental stress (Jackson and Gorovsky,

2000; Kobor et al., 2004; Mizuguchi et al., 2004). Crystallo-

graphic studies have shown that H2A.Z-containing nucleosomes

are in general structurally similar to canonical nucleosomes but

possess distinct internal and surface features (Suto et al.,

2000). Biophysical studies also reported differences in nucleo-

some stability, positioning, and higher-order interactions

(Zlatanova and Thakar, 2008). Of interest, it has been recently

demonstrated that purified nucleosomes containing both

histone H2A.Z and the histone H3.3 variant are the least stable

among native nucleosomes to salt-induced dissociation

(Jin and Felsenfeld, 2007; Zhang et al., 2005).

Genome-wide mapping of nucleosome distribution indicates

that the vast majority of budding yeast promoters have a stereo-

typical chromatin architecture, characterized by two well-posi-

tioned nucleosomes (+1 and �1) flanking an 80–230 base pair

region that is relatively depleted for histones and is commonly

referred to as the ‘‘nucleosome-free region’’ (NFR) (Cairns,

2009; Jiang and Pugh, 2009b; Weiner et al., 2010). With its

NFR-proximal edge covering the transcription start site (TSS),

the +1 nucleosome acts as a barrier that occludes the TSS and

helps position downstream nucleosomes in the coding region

(Jiang and Pugh, 2009b). Formaldehyde crosslinking and chro-

matin immunoprecipitation (ChIP) experiments conducted on

the budding yeast Saccharomyces cerevisiae first demonstrated

that histone H2A.Z (called Htz1) is enriched at intergenic regions

upstream of PHO5 and GAL1 even under repressed conditions

(Santisteban et al., 2000). It was subsequently shown in

genome-wide studies that H2A.Z is dramatically enriched at

the promoter-proximal +1 and �1 nucleosomes (Albert et al.,

2007; Raisner et al., 2005), with enrichment diminishing progres-

sively away from the promoter (Albert et al., 2007). The presence


of H2A.Z nucleosomes surrounding most yeast promoters in the

absence of transcription has led to the proposal that H2A.Z-con-

taining nucleosomes help poise genes for transcription (Jin and

Felsenfeld, 2007; Li et al., 2005; Santisteban et al., 2000; Zhang

et al., 2005). In metazoans, H2A.Z is localized to nucleosomes

proximal to promoters of active genes (Rando and Chang,

2009). More recently, H2A.Z has also been implicated in DNA

repair (Morrison and Shen, 2009) and in suppression of spurious

noncoding transcription (Zofall et al., 2009). The molecular func-

tion of H2A.Z in transcription and DNA repair remains obscure.

Previous studies have shown that the 14 subunit S. cerevisiae

SWI/SNF-related SWR1 complex (SWR1) is required for the

incorporation of H2A.Z (Kobor et al., 2004; Krogan et al., 2003;

Mizuguchi et al., 2004). Human counterparts of SWR1, named

SRCAP and p400, have also been identified (Gevry et al., 2007;

Ruhl et al., 2006). SWR1 is itself enriched at promoters, coinci-

dent with the maxima of H2A.Z distribution (Venters and Pugh,

2009). The recruitment of SWR1 to promoters is attributed in

part to the bromodomain-containing Bdf1 subunit of SWR1

and its interaction with acetylated histone H3 and H4 tails (Altaf

et al., 2010; Koerber et al., 2009; Raisner et al., 2005).

How SWR1 carries out the ATP-dependent replacement of

nucleosomal H2A with H2A.Z is not well understood. Studies

from our laboratory have shown that the histone replacement

reaction can be sufficiently reconstituted in vitro with purified

components (Luk et al., 2007; Mizuguchi et al., 2004). This basic

reaction has also been demonstrated with purified components

from mammalian and insect cells and can be enhanced by

acetylation of the nucleosomal substrate (Altaf et al., 2010;

Kusch et al., 2004; Ruhl et al., 2006). In the replacement reaction,

the H2A.Z-H2B dimer is delivered as a unit to SWR1 (Mizuguchi

et al., 2004; Ruhl et al., 2006) specifically to its Swc2 subunit.

Delivery is assisted by an H2A.Z-specific chaperone Chz1,

which is displaced upon H2A.Z-H2B binding (Luk et al., 2007).

A second binding site for H2A.Z-H2B was recently localized to

the N-terminal domain of the Swr1 ATPase subunit (Wu et al.,

2009). The binding of H2A.Z-H2B to SWR1 is independent of

ATP (Wu et al., 2005).

Other important steps of the histone replacement reaction

involve the ATP-dependent eviction of nucleosomal H2A-H2B

and insertion of H2A.Z-H2B. However, the mechanisms by which

these steps are carried out are obscure. It is also unclear whether

SWR1 replaces one or both histone H2A-H2B dimers in a canon-

ical nucleosome with H2A.Z-H2B, producing heterotypic (AZ) or

homotypic (ZZ) H2A.Z-containing nucleosomes. In vitro reconsti-

tution by salt dialysis shows that the two species can be reconsti-

tuted from purified histones and DNA (Chakravarthy et al., 2004;

Suto et al., 2000). Therefore, it is of particular interest to determine

whether promoter-proximal H2A.Z nucleosomes are organized in

the AZ or ZZ state because they are indistinguishable by standard

ChIP procedures (Albert et al., 2007; Raisner et al., 2005). Muta-

tion of the ATP-binding pocket of the Swr1 ATPase subunit and

studies with nonhydrolyzable ATP analogs documented that

ATP hydrolysis is an absolute requirement for the histone

replacement reaction (Mizuguchi et al., 2004). How the ATPase

activity of the SWR1 complex is transduced to the eviction of

H2A-H2B and insertion of H2A.Z-H2B and whether the ATPase

activity is regulated in the course of the reaction are unknown.

In this study, we investigated whether the homotypic and

heterotypic states of H2A.Z-containing nucleosomes are

present at the promoters of budding yeast in vivo. We found

that promoter-proximal nucleosomes are highly heterogeneous

in histone variant composition, with substantial representation

of nucleosomes containing one, two, or zero H2A.Z molecules.

To further understand this phenomenon, we developed an

in vitro assay to distinguish between compositional states and

found that the histone replacement reaction is stepwise and

unidirectional, i.e., progressing from AA (canonical) to AZ to ZZ

nucleosomes. Further investigation of the underlying mechanism

showed that ATP hydrolysis by the SWR1 complex is specifically

activated by H2A-containing nucleosome and additionally by

H2A.Z-H2B dimer, leading to histone replacement. These results

lead to a model in which specific activation of SWR1 by the two

in vivo histone substrates drives the stepwise, unidirectional

pathway of histone H2A.Z replacement.

RESULTS

Both AZ and ZZ Nucleosomes Are Presentin Saccharomyces cerevisiae

To investigate whether budding yeast nucleosomes contain one

or two copies of the H2A.Z variant, we performed coimmunopre-

cipitation (co-IP) analysis with the use of a diploid strain in which

one allele of HTZ1, the gene encoding S. cerevisiae H2A.Z, is left

untagged and the second HTZ1 allele bears a Flag epitope tag

(HTZ1FLAG) to facilitate purification. Cells in asynchronous

culture were fixed with formaldehyde to preserve nucleosome

integrity, and mononucleosomes were generated by MNase

digestion (Figure S1A available online). We immunoprecipitated

Htz1Flag-containing nucleosomes with anti-Flag antibodies and

analyzed their composition by reversal of crosslinking and

western blotting. Probing with anti-Htz1 antibodies showed

that untagged Htz1 copurifies with Htz1Flag, indicating the pres-

ence of homotypic H2A.Z (ZZ) nucleosomes in yeast cells

(Figure 1A). Moreover, reprobing the same blot with anti-H2A

antibodies shows copurification of H2A with Htz1Flag, demon-

strating the existence of heterotypic H2A.Z (AZ) nucleosomes

as well (Figure 1B). Based on the experimentally determined

ratios (Z:ZF = 0.29, Figure 1A; A:ZF = 0.49, Figures 1C and 1D),

we calculated the relative distribution of ZZ and AZ nucleosomes

to be �35% and �65%, respectively (Figure S1B).

In principle, AZ nucleosomes could be generated from AA

nucleosomes by stepwise replacement with H2A.Z-H2B dimers.

This replacement could occur in a replication-independent

manner in all phases of the cell cycle, including the G1 phase

(S. Sen and C.W., unpublished data). In addition, AZ nucleo-

somes could arise as a consequence of disruption of ZZ nucle-

osomes and reassembly with a mixed histone dimer pool during

DNA replication in S phase. The latter contribution can be mini-

mized in our analysis by the use of yeast cells arrested in G1

phase by a-mating factor (Figure 1F). Under these conditions,

a haploid yeast strain carrying Htz1Flag as the sole copy still

exhibits substantial copurification of H2A with Htz1Flag (�75%

compared to asynchronous cells) (Figure 1E).

We measured the relative proportion of H2A.Z to H2A bound to

chromatin in G1-arrested cells by quantitative western blotting


using purified bacterially expressed Htz1 and H2A as protein

standards (Figures 1G and 1H). Htz1 constitutes �9% of total

H2A-like histones in chromatin, comparable to previous results

obtained for mammalian cells (4%) (West and Bonner, 1980).

ZZ and AZ Nucleosomes Are Enriched at PromotersThe presence of homotypic and heterotypic H2A.Z nucleosomes

in G1-arrested cells prompted us to map their genomic

locations. Both AZ and ZZ nucleosomes could be enriched at

promoters genome-wide, or they could be differentially distrib-

uted among distinct sets of genes. To distinguish between these

possibilities, we used sequential IP to fractionate the heteroge-

neous nucleosome population into three subpopulations

representing ZZ, AZ, and AA nucleosomes. We first immunopuri-

fied Htz1Flag-containing nucleosomes from haploid yeast cells

(expressing Htz1Flag as sole source) with the use of anti-Flag

antibodies to isolate ZZ and AZ nucleosomes in the bound

fraction, followed by secondary IP of the eluate with anti-H2A

antibodies to separate ZZ from AZ nucleosomes (Figure S2A).

Western blotting of bound and flowthrough fractions confirmed

that IP was highly efficient (Figure S2B). In addition, the

flowthrough fraction from the first anti-Flag IP, which is quantita-

Figure 1. Isolation of Homotypic ZZ and

Heterotypic AZ Nucleosomes

(A and B) Histone co-IP analysis of mononucleo-

somes prepared from fixed diploid HTZ1FLAG/

HTZ1 cells (yEL021). (A) The SDS-PAGE and

anti-Htz1 (a-Htz1) western analyses of MNase-

treated nuclear extract (Input), flowthrough of

anti-Flag IP (FT), and anti-Flag immunoprecipi-

tates eluted with Flag peptides (Flag eluate). 20,

10, 5, and 2 ml of the Flag eluate was loaded in

lanes 3, 4, 5, and 6, respectively. Lane 3 was

imaged from a separate western blot. The ratio

of untagged Htz1-to-Htz1Flag for the Flag eluate

is 0.29 ± 0.08 (average and range of two western

analyses). The membrane was stripped and

reprobed with anti-H2A (a-H2A) antibodies in (B).

(C and D) The Flag eluate of (A) was quantified by

a-Htz1 and a-H2A western analyses using

recombinant Htz1 and H2A standards. The esti-

mated molar ratio of H2A to Htz1Flag in the Flag

eluate is 0.49.

(E) Co-IP and western analyses of the Flag eluate

from G1-arrested, asynchronous haploid cells

(yJL036). Numbers indicate quantification of the

Htz1Flag western signal relative to H2A.

(F) FACS analysis ofasynchronous andG1-arrested

cells.

(G and H) Quantification of total H2A and Htz1 poly-

peptides in the nuclear extract (Input) of G1-ar-

rested cells. Asterisk (*) indicates a cross-reactive

band. The two panels in (H) are imaged from the

same western blot.

See also Figure S1.

tively depleted for AZ and ZZ nucleo-

somes, was subjected to additional IP

with anti-H2A to give a bound fraction

highly enriched for AA nucleosomes.We mapped the locations of each distinct nucleosomal pop-

ulation by hybridization of amplified, fluorescently labeled DNA

to oligonucleotide tiling microarrays covering two yeast chro-

mosomes (chromosomes 3 and 6 plus other selected regions),

at 10 bp resolution, for both DNA strands. The results are pre-

sented as normalized ratios of nucleosomal to genomic DNA

fluorescence (Figure 2A and Figure S3). Consistent with

previous studies (Albert et al., 2007; Raisner et al., 2005), we

confirmed that Htz1-containing nucleosomes (Z total) map

predominantly to the promoter-proximal +1 and �1 nucleo-

somes, with enrichment tapering off away from the promoter

(Figure 2). Of interest, we found that the subpopulations of

AZ and ZZ nucleosomes are similarly enriched at most

promoters (Figure 2A). This is especially evident in the normal-

ized average profiles for 466 nucleosomes in the +1 position

(Figure 2B). The relative abundances of AZ and ZZ nucleo-

somes at the +1 location are highly correlated (R = 0.89),

arguing against differential enrichment of AZ or ZZ nucleo-

somes for a specific subset of promoters (Figure S2C). The

average AZ and ZZ nucleosome profiles surrounding the

promoter region also show differences. ZZ nucleosome enrich-

ment is more restricted to the �1 and +1 positions, whereas AZ


enrichment is comparatively lower and declines more gradually

away from the promoter (Figure 2B).

Substantial Presence of AA Nucleosomes at PromotersPrevious studies of H2A.Z enrichment at promoters genome-

wide did not include canonical (AA) nucleosomes, which are

commonly assumed to be depleted at promoters. To test this

assumption, we determined the genomic distribution of the puri-

fied AA nucleosome subpopulation on tiling microarrays

(Figure S2A). As anticipated, the normalized AA nucleosome

distribution is similar to that observed for the total nucleosome

pool (total) (Figures 2A and 2B and Figure S3). However, the abun-

dance of AA nucleosomes at promoters is surprisingly substan-

tial, despite enrichment of the ZZ and AZ variants. This is espe-

cially evident at the �1 and +1 nucleosome positions, where

H2A.Z is thought to be predominant but in fact exhibits a similar

abundance to canonical H2A (Figure 2B). We conclude that

steady-state histone variation at promoter-proximal nucleo-

somes is quite heterogeneous in a population of budding yeast,

showing significant levels of both variant and canonical nucleo-

somes. Clustering analysis of H2A.Z nucleosome distributions

for the TATA-containing and TATA-less promoters shows that

histone heterogeneity appears to be a common feature of most

yeast promoters, irrespective of gene category (Figure S2D).

SWR1 Generates Nucleosomal AZ Intermediate and ZZEnd Product In VitroThe steady-state level of H2A.Z at promoter-proximal nucleo-

somes is a product of opposing H2A.Z assembly and disas-

sembly pathways in vivo. Incorporation of H2A.Z in nucleosomes

Figure 2. Genomic Distribution of the AA, AZ, and ZZ Nucleosomes

(A) Tiling microarray data of a representative region in chromosome 3 showing the genomic distribution of the Z total (orange), ZZ (green), AZ (purple), AA (red),

and total (blue) nucleosomes. The data are presented as the normalized ratio of nucleosomal and genomic DNA signals. Gray bars indicate coding regions.

(B) Normalized average nucleosome distribution in and around the +1 nucleosome center of 466 genes (Jiang and Pugh, 2009a). Circles illustrate the estimated

positions of the �1, +1, +2, +3, and +4 nucleosomes.

See also Figure S2 and Table S2.


is catalyzed by the SWR1 chromatin remodeling complex, which

could convert AA nucleosomes to the ZZ state by replacing both

nucleosomal H2A-H2B dimers with Htz1-H2B in a concerted

reaction. Alternatively, SWR1 could replace the H2A-H2B dimers

in a stepwise manner involving AZ nucleosomes as a reaction

intermediate. To distinguish these models, we developed

a new histone replacement assay.

In this assay, immobilized arrays of canonical nucleosomes

are incubated with SWR1 purified from an htz1D strain,

Htz1Flag-H2B dimers, and ATP as previously described (Fig-

ure 3A). The chromatin product is then subjected to MNase

digestion to liberate mononucleosomes. Because Htz1 bears

a 33Flag epitope tag, replacement of one nucleosomal H2A-

H2B dimer with Htz1Flag-H2B retards the native electrophoretic

mobility of the nucleosome, and replacement of two dimers

retards the mobility further. Thus, in an ATP-dependent, limited

replacement reaction, three nucleosomal species with discrete

mobility can be resolved by nondenaturing PAGE (Figures 3A

and 3B). We examined the identities of each nucleosome

species by western blotting and confirmed that the top, middle,

and bottom gel bands correspond to ZZ nucleosomes with two

Flags (ZFZF), AZ nucleosomes with one Flag (AZF), and AA nucle-

osomes, respectively (Figure S4A).

The detection of AZ nucleosomes in a partial replacement

reaction suggests that the heterotypic H2A.Z nucleosome may

be a reaction intermediate. To investigate this possibility, we

monitored the progression of the SWR1-catalyzed replacement

reaction in vitro. Consistent with the hypothesis, we found that

the AZ species briefly accumulates upon the addition of ATP,

reaching a maximum at 30 min, followed by a gradual decrease

over time (Figure 3C and Figure S4B). By contrast, the ZZ

Figure 3. In Vitro Assay Showing the Step-

wise Assembly of AZ and ZZ Nucleosomes

(A and B) Overview and experiment for the in vitro

histone replacement assay. Bead-bound canon-

ical nucleosome arrays (depicted with three nucle-

osomes for simplicity) were incubated with

Htz1Flag-H2B dimer (chaperoned by Chz1, not de-

picted), SWR1, and ATP for 1 hr (step 1). After

washing, the chromatin was digested with MNase

to liberate mononucleosomes (step 2), which were

subsequently analyzed by nondenaturing PAGE

(step 3). AA (bottom), AZF (middle), and ZFZF

(top) nucleosomes were detected by SYBR green

staining.

(C) In vitro histone replacement time course.

SWR1-mediated histone replacement reactions

were stopped at various times by bead pull-

down and washing. Nucleosomal products were

analyzed as described in (A). (Middle) Densito-

metric measurement of the indicated gel region.

(Right) Peak height versus time.

species continues to accumulate past

30 min, reaching a plateau where ZZ

nucleosomes represent the bulk of the

nucleosome population, and AA nucleo-

somes are correspondingly diminished

to a minor fraction (Figure 3C and Fig-

ure S4B). Thus, reaction kinetics suggests that SWR1 converts

AA nucleosomes to the AZ and ZZ species in a stepwise manner.

Data of the above experiment do not inform whether a fully

replaced ZZ nucleosome is the reaction end product or

a substrate for additional rounds of H2A.Z replacement (i.e.,

H2A.Z replacing H2A.Z). We addressed this question by first

generating a mixed population of immobilized AA, AZ, and ZZ

nucleosomes by a partial replacement reaction in which Htz1-

H2B dimers provided to SWR1 bear a fluorescent Alexa633 label

on Htz1 and a Flag tag on H2B (Htz1Alexa-H2BFlag dimers) (Fig-

ure 4A). (Analysis of an aliquot by MNase digestion confirms

that mononucleosome products exhibit retarded electrophoretic

mobility and Alexa633 fluorescence depending on the extent of

replacement—the bottom band corresponding to unreplaced

nucleosomes, and the middle and top bands to nucleosomes

containing one and two Htz1Alexa-H2BFlag dimers, respectively

[Figures 4A and 4B, lane I].) A second round of SWR1-mediated

histone replacementusing untagged, unlabeled Htz1-H2B dimers

enabled us to evaluate whether the two Htz1Alexa-H2BFlag dimers

in the ZZ nucleosome were replaceable, as shown by a loss of

Alexa633 fluorescence, SYBR green staining, and Htz1 content

in the top nucleosome band (Figure 4A). However, all three indica-

tors remained essentially unchanged after the second SWR1

reaction, indicating that SWR1 does not catalyze replacement

of ZZ nucleosomes with new H2A.Z-H2B dimers (Figure 4B).

This experiment also permitted us to confirm directly that the

heterotypic AZ nucleosome (middle band) is a substrate for

SWR1-catalyzed histone replacement by virtue of a potential

increase in Htz1 content without a change in electrophoretic

mobility (Figures 4A and 4B, lane II). We found that the middle

band indeed shows a major increase in the Htz1 western blotting


signal, demonstrating that the AZ nucleosome, like the AA nucle-

osome (bottom band) is a substrate for SWR1 activity (Figure 4B,

bottom, lane II). Taken together, these results provide compel-

ling evidence that the AZ and ZZ nucleosomes are a bona fide

intermediate and end product, respectively, of the SWR1-medi-

ated histone replacement reaction.

No Reverse Replacement of ZZ Nucleosomes with H2A-H2B DimersWe confirmed that SWR1 does not replace ZZ nucleosomes with

H2A.Z-H2B dimers using immobilized ZZ nucleosome arrays

reconstituted from bacterially expressed histones. Incubation

of these arrays with Flag-tagged histone dimers, SWR1, and

ATP showed that SWR1 failed to replace ZZ nucleosomes with

Htz1Flag-H2B dimers even when dimers were in excess relative

to nucleosomes (Figure 4C).

Next, we examined whether AZ and AA nucleosomes could be

produced from the ZZ species though a reverse reaction by incu-

bation of immobilized ZZ nucleosome arrays with excess

H2AFlag-H2B dimers, SWR1, and ATP. This reaction also failed

to produce detectable incorporation of H2AFlag above back-

ground in the bead-bound chromatin fraction (Figure 4D). We

also tested whether the related INO80 remodeling complex

could mediate a reverse replacement reaction and found no

detectable ATP-driven exchange of H2AFlag into ZZ nucleosomal

arrays under reaction conditions (Figure 4D). Thus, other mech-

anisms may be responsible for the displacement of H2A.Z and

reassembly of the canonical nucleosome. By contrast, incuba-

tion of AA nucleosome arrays with saturating H2A-H2B dimers

(60 nM) gave a small but reproducible level of ATP-dependent

incorporation of H2AFlag into chromatin (Figure 4E), consistent

with previous findings (Mizuguchi et al., 2004). We conclude

that the histone replacement pathway mediated by SWR1 is

unidirectional, with strong substrate specificity for H2A-contain-

ing nucleosomes and the Htz1-H2B dimer.

Canonical Nucleosomes Specifically Stimulate SWR1ATPaseHistone variant replacement by the multicomponent SWR1

complex involves interaction with at least three essential

substrates: ATP, the H2A-containing nucleosome, and the

Htz1-H2B dimer. The differential utilization of H2A-containing

nucleosomes suggests that SWR1 recognizes H2A- over

H2A.Z-containing nucleosomes. Specific recognition could be

Figure 4. AA or AZ Nucleosomes Together with Htz1-H2B Dimer Are the Specific Substrates for SWR1

(A and B) Overview and experiment for the in vitro histone replacement assay. Nucleosomal arrays bearing a mixed population of AA, AZ, and ZZ nucleosomes

were marked with Htz1Alexa-H2BFlag dimers. After incubating with unlabeled, untagged Htz1-H2B, SWR1, and ATP, two potential scenarios depicted in II and III

could occur. (B) is the experiment. (Red) Htz1Alexa. (Flag) H2BFLAG. (Scan) Densitometric analysis of the a-Htz1 western blot.

(C–E) Standard histone replacement assay (Mizuguchi et al., 2004). Immobilized AA or ZZ nucleosomal arrays were incubated with SWR1 (or INO80), native Flag-

epitope-tagged histone dimers, and ATP where indicated. 60 nM of dimers and �15 nM nucleosome equivalents were used. The arrays were washed with 0.4 M

KCl before SDS elution and western analysis. (Top) SDS-eluted fraction of the chromatin-bound histones. (Bottom) Free histones in the supernatant fraction. AA

and ZZ ovals indicate the type of nucleosomal arrays used. ZF/B, Htz1Flag-H2B dimer; AF/B, H2AFlag-H2B dimer.

See also Figure S3.


a consequence of differential nucleosome binding and/or activa-

tion of the ATPase activity of SWR1. We examined whether AA

and ZZ nucleosomes differentially stimulate the ATPase activity

of SWR1 with the use of a real-time fluorescence assay that

monitors production of inorganic phosphate from ATP hydrolysis

(Brune et al., 1994).

Previously, we reported that the SWR1 complex exhibits

nucleosome-stimulated ATPase activity as shown by hydrolysis

of P32-ATP (Mizuguchi et al., 2004). This was confirmed by the

fluorescence assay, which shows strong stimulation of ATP

hydrolysis by conventional nucleosomes, and not by naked

DNA (Figure 5A and Figure S5A). Analysis of initial rates indicates

an �2.5-fold increase of ATP hydrolysis at saturating nucleo-

some and ATP levels. Strikingly, similar concentrations of ZZ

nucleosomes failed to stimulate the ATPase activity of the

SWR1 complex (Figure 5A and Figure S5A). This demonstrates

that SWR1 can functionally discriminate between conventional

and variant nucleosomes. By contrast, INO80 and SWI/SNF

exhibit no differential stimulation of ATPase activity by saturating

levels of AA and ZZ nucleosomes (Figures 5B and 5C and Fig-

ure S5). Of interest, both free H2A-H2B and Htz1-H2B dimers

failed to stimulate the ATPase activity of SWR1 in the absence

Figure 5. AA, but Not ZZ, Nucleosomes Stimulate SWR1 ATPase

(A–C) ATPase assay for chromatin remodelers. Inorganic phosphate (Pi) produced during ATP hydrolysis was monitored in real time by MDCC-PBP, which

increases in fluorescence upon phosphate binding (Brune et al., 1994). Reactions were performed at 23�C in the absence (orange) or presence of 15 nM AA nucle-

osomes (red), ZZ nucleosomes (green), or free DNA (blue). ATP was added �20 s before the first measurement (zero time) to final concentrations of 62.5 mM for

SWR1 and 500 mM for INO80 and SWI/SNF. Relative fluorescence was set as zero at zero time for all reactions.

(D) ATPase assay for SWR1 in the absence (orange) or presence of 15 nM recombinant Htz1-H2B dimers (Z/B, black), H2A-H2B dimers (A/B, gray), or AA nucle-

osomes (red).

(E) ATPase assay for SWR1 in the presence of AA nucleosomes and various ATP concentrations. Phosphate concentrations (calculated based on a linear phos-

phate standard curve) were plotted against time. Initial rate (vo) was determined by the slope of the linear part of each curve (0–300 s).

(F and G) Plots of initial rate versus substrate (ATP) concentrations for 1 nM SWR1 and 0.1 nM INO80 in the presence or absence of 15 nM AA nucleosomes, ZZ

nucleosomes, or DNA. The kinetic parameters Vmax and KM were determined by nonlinear fitting of the Michaelis-Menten curve over plotted values.

(H) Turnover number kcat (obtained from dividing Vmax by total enzyme concentration) and KM for the ATPase of SWR1, INO80, and SWI/SNF in the presence or

absence of 15 nM AA nucleosomes, ZZ nucleosomes, or DNA. Error bars represent the range of two measurements.

See also Figure S5.


of nucleosomes, suggesting that H2A-specific recognition must

be in the context of nucleosome architecture (Figure 5D).

We determined kinetic parameters for ATP hydrolysis by the

SWR1 complex (Figures 5E, 5F, and 5H). SWR1 has an enzyme

turnover rate (kcat) of 0.1 s�1 in the presence or absence of DNA.

The kcat remains essentially the same when SWR1 is incubated

with ZZ nucleosomes but increases to 0.25 s�1 (2.5-fold) with

saturating AA nucleosomes (Figures 5F and 5H). Hence, binding

of H2A-containing nucleosomes to SWR1 stimulates ATPase

activity by increasing the catalytic efficiency of the enzyme.

Figure 6. Further Binding of H2A.Z-H2B Dimer Hy-

peractivates SWR1 ATPase and Evicts Nucleo-

somal H2A-H2B

(A) Standard histone replacement assay (Mizuguchi et al.,

2004). Immobilized AA nucleosomal arrays (reconstituted

with H2AHA histone) were incubated with SWR1,

Htz1Flag-H2B (Z/B), histone chaperones, and ATP where

indicated. (Top) Western analyses of chromatin-bound

histones eluted by SDS. (Bottom) Western analyses of

unincorporated histones. 22 nM of Chz1 or FACT was

added to facilitate possible histone eviction.

(B) ATPase assay for SWR1 in the presence of 15 nM AA

nucleosomes and 15 nM Htz1-H2B dimers (purple). Red

is the AA only control.

(C) Kinetic parameters of SWR1 ATPase in the presence of

15 nM AA nucleosomes and 15 nM Htz1-H2B dimers

(purple). For comparison, the curve and parameters for

AA alone (red) are reproduced from Figures 5H and 5F,

respectively. Errors represent the range of two measure-

ments.

(D) Specificity of SWR1 ATPase hyperstimulation. ATPase

assay was performed as described in Figure 5A except

with different combinations of nucleosomes (Nuc) and

histone dimers. Z/B, Htz1-H2B dimers; A/B, H2A-H2B

dimers; AA, AA nucleosomes; ZZ, ZZ nucleosomes; (–)

control, no dimer or nucleosome.

See also Figure S6.

The Michaelis constant (KM), which represents

the ATP concentration at half maximal velocity

(1/2 Vmax), shows little change for canonical

and variant nucleosomes (5 mM and 7 mM,

respectively). In comparison, the stimulated

SWI/SNF has a kcat of 5.5 s�1 and KM of 80.5

mM, values consistent with previous determina-

tions (Smith and Peterson, 2005) (Figure 5H).

Nucleosome Stimulation of SWR1ATPase Is Not Sufficient for H2A-H2BEvictionThe stimulation of SWR1 ATPase by incubation

with conventional nucleosomes raised the

question of whether such ATP hydrolysis would

be sufficient for eviction of the nucleosomal

H2A-H2B dimer to facilitate Htz1-H2B deposi-

tion. To test this hypothesis, we incubated

SWR1 with immobilized arrays of conventional

nucleosomes carrying epitope-tagged histone

H2AHA and monitored H2AHA-H2B eviction in the supernatant

fraction by western blotting. Whereas the SWR1-catalyzed

Htz1 replacement reaction in the presence of Htz1-H2B dimer

(and histone chaperone) occurs robustly with quantitative evic-

tion of H2AHA, we did not detect any eviction of histone H2AHA

in the absence of Htz1Flag-H2B dimer (Figure 6A). Thus, the stim-

ulation of ATP hydrolysis provided solely by canonical nucleo-

some effector is inadequate for eviction of nucleosomal

H2A-H2B. Moreover, eviction of H2A-H2B and insertion of

Htz1-H2B appear to be coupled processes.


Htz1-H2BDimer andCanonical NucleosomeSpecificallyActivate SWR1The requirement for free Htz1-H2B dimers for SWR1-mediated

H2A.Z replacement prompted us to investigate whether addition

of Htz1-H2B to SWR1 further increases ATP hydrolysis. Indeed,

we observed a clear hyperstimulation of the ATPase activity of

SWR1 when saturating Htz1-H2B dimers (15 nM), and canonical

nucleosomes are both provided to SWR1 in the reaction (Figures

6B and 6D). The hyperstimulated ATPase activity exhibits a kcat

of 0.45 s�1, which represents an additional 1.8-fold increase in

the kcat relative to the stimulation by nucleosomes only (a 4.1-

fold increase in total), with little change of KM (Figure 6C). We

observed less hyperstimulation when H2A-H2B dimers were

substituted for Htz1-H2B at the same molar concentration

(Figure 6D, left). Importantly, a 4-fold increase of H2A-H2B

dimers (�60 nM) hyperstimulated ATPase activity to nearly

maximal level (Figure 6D, right), whereas hyperstimulation of

the ATPase activity upon addition of Htz1-H2B or H2A-H2B

dimers (at either concentration) to ZZ nucleosomes was much

lower (Figure 6D). Given that incorporation of new H2A in canon-

ical nucleosomal arrays is low (Figure 4E) under conditions

wherein ATPase activity is high, these findings indicate that

high ATPase activity per se is not sufficient for histone replace-

ment. It is the presence of the correct in vivo substrates that

ensures efficient coupling of the high ATPase activity to histone

replacement.

DISCUSSION

The steady-state level of H2A.Z at promoter-proximal nucleo-

somes is a consequence of the opposing pathways of H2A.Z

incorporation and H2A.Z eviction. Our observation of three

distinct variant states of promoter nucleosomes in a cell popu-

lation is complementary to previous mapping studies of H2A.Z

in budding yeast (Albert et al., 2007; Raisner et al., 2005; San-

tisteban et al., 2000). The comparable representation of AA

and ZZ states suggests that the AA-to-ZZ and ZZ-to-AA path-

ways are balanced for many genes, without one pathway domi-

nating. However, this balance can be shifted, for example, at

highly transcribed promoters (top 10% RNA Pol II occupancy)

in which the ZZ and AZ states are underrepresented relative

to the AA state for the +1 nucleosome position (Figure S2E),

suggesting that H2A.Z eviction is occurring at a faster rate

than incorporation. The greater restriction of the ZZ than AZ

state to +1 and �1 nucleosome positions is interesting and

may be a consequence of the stepwise nature of the histone

replacement reaction and the local concentration of SWR1

recruited to gene promoters (Venters and Pugh, 2009; Yoshida

et al., 2010).

Our in vitro studies show that SWR1 is capable of stepwise

deposition of H2A.Z-H2B into canonical nucleosomes, coupled

with H2A-H2B eviction, to give a fully replaced variant nucleo-

some. However, once incorporated, H2A.Z cannot be evicted

by SWR1, even in excess of either H2A.Z-H2B or H2A-H2B

dimers under otherwise identical reaction conditions. Therefore,

the SWR1-mediated pathway of H2A.Z replacement is unidirec-

tional, terminating with ZZ nucleosomes. It is possible that

a reverse reaction from the ZZ to AA nucleosome state requires

different conditions, cofactors, or modifications of the SWR1

enzyme or histone substrates. Alternatively, a return to the AA

state may occur through separate pathways. For example, other

SWI/SNF family members might possess the capability for

specific replacement of nucleosomal H2A.Z-H2B with H2A-

H2B, but we have not observed such activity for INO80,

a chromatin remodeling complex paralogous to SWR1, under

conditions in which INO80 displays robust nucleosome- or

DNA-stimulated ATPase and histone octamer sliding activities

(Figure 5, Figure S5, and unpublished data).

More likely, the well-documented high histone H3 turnover

rate at promoters implies promoter-specific nucleosome

disassembly, i.e., eviction of H2A.Z-H2B and H3-H4, and subse-

quent nucleosome reassembly with new histones (Dion et al.,

2007; Rufiange et al., 2007), thereby completing the dynamic

cycling of H2A and H2A.Z at gene promoters (Figure 7A).

These processes are likely to be mediated by a combination of

SWI/SNF family enzymes (Barbaric et al., 2007; Gutierrez

Figure 7. A Model for SWR1-Mediated

Histone Replacement

(A) Promoter nucleosome cycle. An AA nucleo-

some at the +1 promoter-proximal position is con-

verted to the AZ and ZZ states by SWR1 via a step-

wise, unidirectional pathway. The ZZ nucleosome

is subsequently converted back to the AA state

through pathways most likely involving nucleo-

some eviction and reassembly with an AA nucleo-

some (dotted gray arrows).

(B) SWR1 catalytic cycle. SWR1 stochastically

binds to one face of an AA nucleosome and the

H2A.Z-H2B dimer, leading to hyperstimulation of

ATPase activity (deep red) and a conformational

change in SWR1 (shown) required for histone

replacement. The newly incorporated Z face of

the AZ nucleosome deactivates the ATPase and

stops further histone replacement activity. The

AZ nucleosome dissociates and rebinds stochas-

tically on the A face for a second replacement

reaction.


et al., 2007; Lorch et al., 2006), RNA polymerase (Weiner et al.,

2010), and core histone chaperones (Corpet and Almouzni,

2009; Das et al., 2010). In addition, the in vivo lability of H2A.Z-

containing nucleosomes as reflected in salt sensitivity should

also contribute (Henikoff et al., 2009; Jin and Felsenfeld, 2007;

Zhang et al., 2005). Indeed, histone modifications at promoters

correlate with the signatures of newly deposited histones, such

as H3K56Ac and H4K16 deAc (Rando and Chang, 2009).

The directional nature of the H2A.Z replacement pathway

implies that SWR1 must functionally differentiate between ZZ

and AA (or AZ) nucleosomes. We have traced this differentiation,

at least in part, to a specific, 2.5-fold increase of the ATPase

activity (kcat) of SWR1 induced by AA, but not ZZ, nucleosomes.

However, this level of stimulation is insufficient for the eviction of

H2A-H2B from nucleosomes. Only after further addition of free

H2A.Z-H2B dimers is the ATPase activity of SWR1 hyperstimu-

lated (4-fold increase of kcat), concurrent with H2A-H2B eviction

and H2A.Z-H2B deposition. However, a hyperstimulated SWR1

ATPase is only necessary, but not sufficient, to mediate robust

histone replacement, as saturating free H2A-H2B dimers can

hyperstimulate SWR1 ATPase to nearly maximal level but with

substantially reduced histone replacement (Figure 4E and Fig-

ure 6D). This finding implies that unique features of H2A.Z-H2B

dimer, in addition to stimulating ATP hydrolysis, enhance histone

replacement by allosterically coupling the ATPase motor to

histone transactions. This additional molecular specificity seems

biologically necessary, given that H2A-H2B dimers should be in

excess over H2A.Z-H2B dimers in vivo.

Overall, our data suggest a model in which SWR1 binding to

and recognition of its two in vivo histone substrates (one face

of the AA nucleosome and the H2A.Z-H2B dimer) lead to hyper-

stimulation of ATPase activity as well as a conformational

change in SWR1 required for displacement of H2A-H2B and

insertion of H2A.Z-H2B (Figure 7B). The order of SWR1 binding

to nucleosomes and H2A.Z-H2B dimers should be stochastic.

The newly incorporated Z face of the AZ nucleosome deactivates

the ATPase and stops further histone replacement. The AZ

nucleosome subsequently dissociates from and reassociates

with SWR1 in a stochastic fashion (Figures 7B and 7C). In the

second round, recognition by SWR1 of the A face of the AZ

nucleosome and new H2A.Z-H2B dimer binding restimulates

SWR1 activity to catalyze replacement of the remaining nucleo-

somal H2A-H2B with H2A.Z-H2B. Functional recognition of the A

face of an AA or AZ nucleosome and the requirement for free

H2A.Z-H2B dimer ensures that only these effectors, which are

also substrates for SWR1, are productively utilized. This

provides a way of controlling the specificity and outcome of

the replacement reaction, which terminates with the ZZ

nucleosome.

The SWR1 complex contains multiple ATP-binding subunits,

including Swr1, actin, actin-related proteins Arp4 and Arp6,

and the Rvb1-Rvb2 dodecamer, members of the AAA+ family

of ATPases (Jha and Dutta, 2009; Mizuguchi et al., 2004). We

have previously found that a mutation (K727G substitution) in

the ATP-binding motif of the Swr1 subunit is sufficient to

abrogate Htz1 replacement in vivo and in vitro without affecting

assembly of the SWR1 complex (Mizuguchi et al., 2004). The

ATPase activity of the purified mutant enzyme is neither stimu-

lated by AA nucleosomes nor hyperstimulated by further addition

of Htz1-H2B dimer (Figure S6). These findings indicate that the

Swr1 ATPase is the key subunit whose activity is governed,

directly and/or indirectly, by the histone effectors.

It will be interesting to define the molecular determinants

within the canonical nucleosome and the H2A.Z-H2B dimer

that are specifically recognized by the SWR1 complex, to identify

the SWR1 components interacting with the nucleosome, and to

follow the fate of the evicted H2A-H2B dimer. Other questions

are the importance of the two Htz1-binding modules in SWR1

(Swc2 and the N terminus of the Swr1 subunit itself); the relation-

ship between ATPase activity, DNA translocase activity, and un-

wrapping of nucleosomal DNA; the timing and coupling of H2A

eviction and H2A.Z insertion; and the structural transformations

of SWR1 that accompany these events. Our present findings and

the biochemical assays that we have developed should facilitate

future investigations on the mechanism of histone H2A.Z

replacement.


Immunopurification of AA, AZ, and ZZ Nucleosomes

Crude chromatin was isolated from formaldehyde-fixed yeast cells as

described in Liang and Stillman (1997) and digested with MNase to mononu-

cleosomal level. Sequential IP was performed with the use of anti-Flag M2

agarose (Sigma) and anti-H2A antibodies (Active Motif) bound to nProtein A

Sepharose (GE Healthcare).

Amplification and Labeling of Nucleosomal DNA for Microarray

Analysis

Nucleosomal DNA and MNase-treated genomic DNA control were treated with

alkaline phosphatase (CIP, NEB) and end-repair enzyme mix (End-It kit, Epi-

centre) before being amplified by ligation-mediated PCR (Johnson et al.,

2008). Labeling was performed using the BioPrime Plus labeling kit (Invitrogen)

according to the manufacturer’s protocol.

Microarrays

Custom tiling microarrays were designed based on the Agilent 4 3 180K plat-

form. Each microarray contained �150,000 biological probes spanning

selected genomic regions. The tiling probes were spaced, on average, 10 bp

apart and covered both the sense and antisense DNA strands.

Normalization ofMicroarray Data for Different Nucleosomal Species

Given that Htz1 is the only H2A variant in budding yeast, normalization of mi-

croarray data was performed based on the assumptions that, to a first approx-

imation, the sum of Z total and AA nucleosomes is equal to the total nucleo-

some signal and that the sum of AZ and ZZ nucleosomes is equal to the Z

total nucleosome signal. Details are provided in Figure S3 and legend.

In Vitro Histone Replacement Assay

The SWR1 histone replacement assay was performed according to Mizuguchi

et al. (2004) except the immobilized nucleosomal arrays (80 ng DNA equiva-

lents) were digested with 0.16 U/ml MNase (+ 2 mM CaCl2) to liberate the nucle-

osomes. The reactions were stopped with 10 mM EDTA before analysis by

nondenaturing PAGE.

In Vitro Histone Replacement Assay Using Fluorescently Labeled

Htz1-H2B Substrate

To generate the mixed AA, AZ, and ZZ nucleosomal substrate for the experi-

ment in Figure 4B, AA nucleosomal arrays were incubated with SWR1 pre-

charged with Htz1Alexa-H2BFlag and with Htz1-H2BFlag. The resulting chromatin

had comparable levels of AA, AZ, and ZZ nucleosomes, which also exhibited

comparable Alexa633 fluorescence for the AZ and ZZ nucleosomal species. In

the chase step, the labeled nucleosomes were incubated with SWR1


precharged with the unlabeled, untagged Htz1-H2B. After washing, the nucle-

osomal products were released by MNase digestion and analyzed by nonde-

naturing PAGE as described above.

ATPase Assay

ATPase assay was performed based on the procedure described in Brune

et al. (1994). In this assay, inorganic phosphate (Pi) produced during ATP

hydrolysis is monitored by the fluorophore-modified phosphate-binding

protein MDCC-PBP (Phosphate Sensor, Invitrogen), which increases in fluo-

rescence upon Pi binding. Measurements were performed at 23�C on a Wallac

Victor plate reader using a 405 nm excitation, 460 nm emission filter set.

ACCESSION NUMBERS

The GEO accession number for the microarray data sets is GSE24618.


Supplemental Information includes Extended Experimental Procedures, six

figures, and three tables and can be found with this article online at doi:10.

1016/j.cell.2010.10.019.

ACKNOWLEDGMENTS

We thank W.H. Wu for the yeast strain SWR1-FL htz1D and J. Landry for the

yeast strain yJL036. We also thank H. Cam and C. Rubin for advice on micro-

array techniques, F. Pugh and members of the Wu lab for critical reading of the

manuscript, and anonymous reviewers for helpful suggestions. This work was

supported by the intramural research program of the National Cancer Institute

(C.W.) and by the Leukemia and Lymphoma Society (E.L. and A.R.).


Revised: August 25, 2010


Published: November 24, 2010

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Sororin Mediates Sister ChromatidCohesion by Antagonizing WaplTomoko Nishiyama,1 Rene Ladurner,1 Julia Schmitz,1,4 Emanuel Kreidl,1 Alexander Schleiffer,1 Venugopal Bhaskara,1

Masashige Bando,2 Katsuhiko Shirahige,2 Anthony A. Hyman,3 Karl Mechtler,1 and Jan-Michael Peters1,*1Research Institute of Molecular Pathology, Dr. Bohr-Gasse 7, A-1030 Vienna, Austria2Institute of Molecular and Cellular Biosciences, University of Tokyo, Yayoi, Tokyo 113-0032, Japan3Max Planck Institute of Molecular Cell Biology and Genetics, Pfotenhauerstrasse 108, D-01307 Dresden, Germany4Present address: World Health Organization, Avenue Appia 20, CH-1211 Geneva 27, Switzerland


DOI 10.1016/j.cell.2010.10.031

SUMMARY

Sister chromatid cohesion is essential for chromo-some segregation and is mediated by cohesin boundto DNA. Cohesin-DNA interactions can be reversedby the cohesion-associated protein Wapl, whereasa stably DNA-bound form of cohesin is thought tomediate cohesion. In vertebrates, Sororin is essentialfor cohesion and stable cohesin-DNA interactions,but how Sororin performs these functions isunknown.We show that DNA replication and cohesinacetylation promote binding of Sororin to cohesin,and that Sororin displaces Wapl from its bindingpartner Pds5. In the absence of Wapl, Sororinbecomes dispensable for cohesion. We proposethat Sororin maintains cohesion by inhibiting Wapl’sability to dissociate cohesin from DNA. Sororin hasonly been identified in vertebrates, but we showthat many invertebrate species contain Sororin-related proteins, and that one of these, Dalmatian,is essential for cohesion in Drosophila. The mecha-nism we describe here may therefore be widelyconserved among different species.

INTRODUCTION

In eukaryotic cells, sister chromatids remain physically con-

nected from the time of their synthesis during DNA replication

until their separation during mitosis or meiosis. This sister chro-

matid cohesion is essential for biorientation of chromosomes on

the spindle and for DNA-damage repair (reviewed in Nasmyth

and Haering, 2009; Onn et al., 2008; Peters et al., 2008). Cohe-

sion is mediated by cohesin complexes. Three cohesin subunits,

the ATPases Smc1 and Smc3 and the kleisin Scc1/Rad21/

Mcd1, form triangular structures that have been proposed to

mediate cohesion by embracing sister chromatids (Gruber

et al., 2003; for an illustration of this ‘‘ring model,’’ see Figure 6C

below). Scc1 binds to a fourth core subunit, called Scc3 in yeast

and stromal antigen (SA) in vertebrates, where somatic cells

contain two SA paralogs (SA1 and SA2). Scc1 and SA proteins

are further associated with a heterodimer of two proteins, called

Wapl and Pds5, the latter of which also exists in two isoforms in

vertebrates (Pds5A and Pds5B; Gandhi et al., 2006; Kueng et al.,

2006).

Cohesin complexes are loaded onto DNA before replication (in

animal cells already in telophase) and establish cohesion during

replication. In the subsequent mitosis, cohesion is dissolved by

removal of cohesin from chromosomes. In vertebrate cells, this

process occurs in two steps (Waizenegger et al., 2000): the

bulk of cohesin is removed from chromosomes in prophase by

a mechanism that depends on Polo-like kinase 1 (Plk1/Plx1)

and Wapl (Gandhi et al., 2006; Kueng et al., 2006). At centro-

meres, small amounts of cohesin are protected from the

prophase pathway by Shugoshin, and these complexes can

only be removed from chromosomes by the protease separase

(reviewed in Sakuno and Watanabe, 2009). This process occurs

only in metaphase because a surveillance mechanism called the

spindle assembly checkpoint (SAC) prevents separase activa-

tion until all chromosomes have been bioriented. The SAC

inhibits APC/CCdc20 (anaphase-promoting complex/cyclosome

associated with Cdc20), a complex whose ubiquitin ligase

activity is required for separase activation (reviewed in Peters,

2006).

How cohesion is established and maintained is poorly under-

stood. Fluorescence recovery after photobleaching (FRAP)

experiments in mammalian cells revealed that cohesin binds to

DNA much more stably after than before DNA replication, sug-

gesting that cohesion depends on an unidentified event during

DNA replication that stabilizes cohesin on DNA (Gerlich et al.,

2006). The dynamic mode of cohesin binding to DNA might

depend on Wapl because depletion of this protein from mamma-

lian cells does not only interfere with the prophase pathway but

also increases the residence time of cohesin on chromatin during

interphase (Kueng et al., 2006).

The only molecular event during DNA replication that is known

to be essential for cohesion establishment is acetylation of cohe-

sin (Ben-Shahar et al., 2008; Unal et al., 2008; Zhang et al., 2008).

This modification occurs on two lysine residues in the ATPase

domain of Smc3 (K112/113 in budding yeast) and is catalyzed

by the acetyltransferase Eco1. The lethality of yeast that is

caused by deletion of the ECO1 gene can be suppressed by

changing K112/113 to residues that might functionally mimic


Figure 1. Sororin Is Required for Cohesion in S Phase

(A) FISH of Sororin-depleted S phase cells. HeLa cells were synchronized in S phase by double thymidine arrest and transfected with control or Sororin siRNA.

Four hours after release from the second thymidine arrest, cells were labeled with BrdU for 15 min, pre-extracted, and subjected to FISH with a probe specific for


acetylated lysine but also by deletion of the WPL1/RAD61 gene,

which encodes a Wapl ortholog, and by mutations in Pds5 (Ben-

Shahar et al., 2008; Rowland et al., 2009; Sutani et al., 2009; Unal

et al., 2008). Cohesin is also acetylated in mammalian cells on

Smc3 residues K105/106 (Zhang et al., 2008), where two Eco1

orthologs exist, called Esco1 and Esco2 (Hou and Zou, 2005).

In vertebrate cells, cohesin-DNA interactions are also regu-

lated by Sororin. This protein was identified as a substrate of

APC/CCdh1, a form of the APC/C that is active during mitotic

exit and G1 phase, and Soronin was found to be essential for

cohesion in mammalian cells (Rankin et al., 2005). Interestingly,

Sororin depletion also reduces the number of cohesin com-

plexes that bind stably to DNA during G2 phase, indicating that

Sororin is required for the formation of stable cohesin-DNA inter-

actions (Schmitz et al., 2007). However, it is unknown how

Sororin performs this function, and whether the role of Sororin

is related to the function of cohesin acetylation. Furthermore, it

is unknown how widespread the role of Sororin is because

Sororin has only been identified in vertebrates.

Here we provide evidence that Sororin is recruited to chro-

matin-bound cohesin complexes in a manner that depends on

DNA replication and Smc3 acetylation, that Sororin causes

a conformational rearrangement within cohesin by displacing

Wapl from Pds5, and that these molecular events stabilize cohe-

sin on DNA by antagonizing Wapl’s ability to dissociate cohesin

from DNA. Furthermore, we show that distant orthologs of So-

rorin exist in many metazoan species, and that one of these

proteins, Dalmatian, is required for cohesion in Drosophila. We

therefore propose that sister chromatid cohesion depends on

stabilization of cohesin on DNA by Sororin-related proteins.

RESULTS

Sororin Is Required for Cohesion during S PhaseWe had previously shown that Sororin is required for cohesion in

G2 phase (Schmitz et al., 2007). To address whether Sororin’s

function is already needed during S phase, we used RNA inter-

ference (RNAi) to deplete Sororin from HeLa cells that had

been synchronized in the cell cycle and pulse-labeled these cells

with bromodeoxyuridine (BrdU). Cells in S phase were identified

by immunofluorescence microscopy (IFM) using BrdU anti-

bodies, and the distance between sister chromatids was

measured by DNA fluorescence in situ hybridization (FISH) using

a probe for an arm region on chromosome 21. On average, FISH

signals were twice as far separated in BrdU-positive, Sororin-

depleted cells than in control cells (Figures 1A and 1B), indicating

that Sororin is already required for cohesion during S phase. At

variance with these results, it has been reported that Sororin-

depleted cells only lose cohesion during metaphase and that So-

rorin is therefore not required for cohesion in early mitosis (Diaz-

Martinez et al., 2007). However, in time-lapse microscopy exper-

iments we observed that most Sororin-depleted cells failed to

congress chromosomes, consistent with the existence of cohe-

sion defects before metaphase (Figures S1A–S1D available on-

line). The function of Sororin is therefore not restricted to mitosis

and is instead already needed during or shortly after DNA

replication.

Sororin Associates with Chromatin during the Periodof the Cell Cycle Where Cohesion ExistsWe next analyzed the intracellular distribution of Sororin.

Previous IFM and fractionation experiments had shown that So-

rorin associates with chromatin in interphase, but Sororin could

not be detected on mitotic chromosomes (Rankin et al., 2005).

Because our antibodies could not detect Sororin in IFM experi-

ments, we tagged Sororin at its carboxy-terminus with a localiza-

tion-affinity purification (LAP) tag that contains green fluorescent

protein (GFP; Figure S1E). We modified the Sororin gene on

a bacterial artificial chromosome (BAC), enabling gene expres-

sion from the endogenous promoter (Poser et al., 2008). We

used a mouse BAC for these experiments to enable RNAi

‘‘rescue’’ experiments and generated clonal HeLa cell lines

that had stably integrated this BAC. The LAP-tagged version of

mouse Sororin could substitute for the cohesion function of

endogenous human Sororin when this was depleted by RNAi

(Figures S1F and S1G), and in tandem affinity purification exper-

iments mouse Sororin-LAP was found associated with human

cohesin (Figures S1H and S1I), indicating that this tagged

version of Sororin behaves similarly to endogenous Sororin.

We therefore analyzed by IFM the intracellular distribution of So-

rorin-LAP, using antibodies to GFP. We stained proliferating cell

nuclear antigen (PCNA) and Aurora B in the same cells as

markers for S and G2 phases, respectively. Cellular Sororin-

LAP levels were low in G1, accumulated between early S and

G2 phases in the nucleus, and became dispersed in the cyto-

plasm following nuclear envelope breakdown (Figures S1J–

S1L). When we analyzed cells from which soluble proteins had

been extracted before fixation, we observed that Sororin-LAP

accumulated on chromatin between early S phase and G2

phase, whereas most Sororin-LAP disappeared from chromo-

somes in prophase (Figures 1C–1E). At this stage, the cellular

levels of Sororin were still high (Figure S1L), indicating that the

removal of Sororin from prophase chromosomes is caused by

dissociation, not degradation. Biochemical fractionation experi-

ments confirmed this notion (Figure S1M). Importantly, however,

small amounts of Sororin-LAP could still be detected by IFM on

the trisomic tff1 locus on chromosome 21. BrdU-labeled nuclei (blue) with three pairs of FISH signals (red) are shown. Higher-magnification images are shown in

the insets. Bar: 5 mm.

(B) Quantification of the distance between paired FISH signals in (A) (mean ± standard deviation [SD]; n R 30 per condition, *p < 0.01).

(C) Sororin-LAP cells were pre-extracted prior to fixation and stained for Sor-LAP (GFP), PCNA, and Aurora B. DNA was counterstained with Hoechst. Bar: 10 mm.

(D) Quantification of chromatin-bound Sororin-LAP levels in (C) (mean ± SD; n R 50 per class).

(E) Sororin-LAP cells were synchronized in mitosis, pre-extracted prior to fixation, and stained for Sor-LAP (GFP), Scc1, and DNA (Hoechst). Bar: 10 mm.

(F) Sororin-LAP localizes to centromeres in mitosis. Sororin-LAP cells were pre-extracted prior to fixation and stained for Sor-LAP (GFP), kinetochores (CREST),

and DNA (DAPI). Insets show magnified views. Bar: 10 mm.

See also Figure S1.


chromosomes in prophase, prometaphase, and metaphase, but

not in anaphase or telophase (Figure 1E). Like cohesin (Waize-

negger et al., 2000), Sororin-LAP was enriched at centromeres

in prometa/metaphase (Figure 1F). Sororin therefore associates

with chromatin from S phase until metaphase, i.e., as long as

cohesion exists.

The Association of Sororin with Chromatin Dependson CohesinBecause Sororin binds to cohesin and, like cohesin, is removed

from mitotic chromosomes in two steps, during prophase and at

the metaphase-anaphase transition, we tested whether the

association of Sororin with chromatin depends on cohesin.

Scc1 depletion reduced the intensity of Sororin-LAP staining

on chromatin without affecting the cellular levels of Sororin-

LAP (Figures 2A–2D), indicating that Sororin can only efficiently

associate with chromatin in the presence of cohesin. Biochem-

ical experiments in Xenopus egg extracts confirmed this notion

(see Figure 2F below). The presence of Sororin on mitotic

chromosomes also depends on cohesin, as depletion of either

Scc1 or Shugoshin-like 1 (Sgo1) reduced chromosomal So-

rorin-LAP staining, whereas depletion of Wapl or inhibition of

Plk1 increased the amounts of Sororin on chromosome arms

(Figure S2A).

Although the intracellular distribution of Sororin and cohesin is

similar from prophase to anaphase, the two proteins behave

differently in telophase. Whereas cohesin reassociates with

chromatin at this stage, little if any Sororin-LAP could be de-

tected on chromatin in telophase (Figure 1E). This difference

was not due to lower sensitivity in the detection of Sororin than

cohesin because Sororin-LAP could easily be observed on early

mitotic chromosomes, where endogenous cohesin cannot be

detected (due to its low abundance; Waizenegger et al., 2000).

The absence of Sororin on telophase chromatin was also not

caused by APC/CCdh1-mediated degradation of all cellular So-

rorin because Sororin-LAP could be observed in fixed telophase

cells (Figure S1L). Time-lapse microscopy of living cells showed

that Sororin levels begin to decrease in anaphase when APC/

CCdh1 becomes active but revealed that most Sororin degrada-

tion occurs after telophase, i.e., during G1, as is typical for

APC/CCdh1 substrates (Figures S2B–S2E). The absence of So-

rorin on chromatin in telophase is therefore not simply due to

the complete degradation of Sororin.

Efficient Association of Sororin with ChromatinDepends on DNA ReplicationThe absence of Sororin on telophase chromatin could be caused

by local APC/CCdh1-mediated degradation on chromatin, or the

Figure 2. Association of Sororin with

Chromatin Depends on Cohesin and DNA

Replication

(A–D) Sororin-LAP cells were transfected with

siRNAs and synchronized in G2 phase. Cells

were fixed (C and D) or pre-extracted prior to

fixation (A and B) and stained for Sor-LAP (GFP),

Scc1, and DNA (Hoechst). Bar: 10 mm. Quantifica-

tion of Sororin-LAP levels in (A) and (C) is shown in

(B) and (D), respectively (mean ± SD; n R 110 (B)

and n R 130 (D) per condition).

(E) Sororin is stably present throughout the cell

cycle but associates with chromatin during S

phase in Xenopus egg extracts. CaCl2 and cyclo-

heximide were added to meiotic metaphase II

(MII) arrested CSF extract to induce meiotic exit.

At 90 min after CaCl2 addition, D90 Cyclin B was

added to induce mitosis. Samples were taken at

indicated time points after CaCl2 addition (release

from MII) or D90 Cyclin B addition (D90 Cyc B

addition). DNA replication (DNA repl.) was moni-

tored by incorporation of [a-32P]dCTP into sperm

chromatin. Chromatin-bound proteins in the

same extracts are also shown. Chromatin was

preincubated for 30 min in CSF extracts.

(F) Sororin association with chromatin depends

on cohesin. Xenopus interphase extracts were

subjected to mock or SA1/2 immunodepletion.

Two hours after sperm chromatin addition, chro-

matin fractions were analyzed by immunoblotting.

(G) Sororin association with chromatin depends

on DNA replication. Interphase extracts were

incubated for indicated times with sperm chro-

matin. DMSO, aphidicolin (Aph.), or actinomycin

D (ActD) was added to the extracts 25 min

after sperm addition. Chromatin fractions were

analyzed by immunoblotting.

See also Figure S2.


association of cohesin with chromatin could be required but not

sufficient for Sororin binding to chromatin. To distinguish

between these possibilities, we analyzed the chromatin associa-

tion of Sororin in Xenopus eggs, which do not contain Cdh1 and

where Sororin is therefore predicted to be stable during mitotic

exit. If cohesin was sufficient for recruiting Sororin to chromatin,

both proteins would be expected to associate with chromatin

simultaneously in Xenopus egg extracts. To test this possibility,

we isolated two Xenopus Sororin cDNAs (Sororin-A and -B),

which encode closely related 35 kDa proteins. Xenopus Sororin

antibodies recognized both Sororin isoforms in immunoblots

(visible as a doublet of bands; see for example Figure 2E) and

could deplete both proteins from egg extracts (see Figure 4A

below). Immunodepletion experiments also revealed that the

chromatin association of Xenopus Sororin proteins depends on

cohesin (Figure 2F) and that these proteins are required for cohe-

sion (see Figure 4B below), even though their amino acid

sequences are only 38% identical to the sequence of human So-

rorin. The two Xenopus proteins characterized here (hereafter

collectively called Xenopus Sororin) are therefore functionally

related to mammalian Sororin.

To address when Sororin and cohesin associate with chro-

matin, we released Xenopus egg extracts from a cytostatic

factor (CSF) arrest in metaphase of meiosis II into interphase

by addition of Ca2+, which leads to activation of APC/CCdc20,

degradation of mitotic Cyclins, and mitotic exit (Figure 2E). As

a source of chromatin, demembranated sperm nuclei were

added. DNA replication was monitored by incorporation of

[a-32P]dCTP into DNA and occurred within 60 min after Ca2+

addition. After 90 min, we added a recombinant form of nonde-

gradable Cyclin B (D90 Cyc B) to induce entry of the extract into

a mitotic state. At different time points, proteins in the chromatin

fraction or the total extract were analyzed by immunoblotting

(Figure 2E). As expected, Ca2+ addition led to rapid degradation

of Cyclin B2 (a substrate of APC/CCdc20), but the levels of the

APC/CCdh1 substrates Sororin and Plx1 remained largely

unchanged (only the electrophoretic mobility of Sororin was

reduced by phosphorylation in CSF and mitotic extracts). Impor-

tantly, even though Sororin was present throughout all stages of

the cell cycle, it began to associate with chromatin only 60 min

after addition of Ca2+, i.e., when DNA replication was initiated.

In contrast, the cohesin subunits Scc1 and Smc3 could be de-

tected on chromatin at least 30 min earlier. The association of

Sororin with chromatin was abolished by Geminin (Figure S2F),

a protein that inhibits cohesin loading onto DNA (Gillespie and

Hirano, 2004; Takahashi et al., 2004), indicating that our assay

reflected physiological binding of Sororin to chromatin. These

observations suggest that local APC/CCdh1-mediated degrada-

tion of Sororin on chromatin cannot explain why Sororin associ-

ates with chromatin later than cohesin. Instead, our results indi-

cate that the presence of cohesin on chromatin is not sufficient

for recruitment of Sororin.

Because Sororin associates with chromatin during S phase in

Xenopus extracts and in somatic cells (Figure 1C and Figure 2E),

we tested whether DNA replication is required for recruitment of

Sororin to chromatin. We prevented replication in Xenopus

extracts by addition of aphidicolin or actinomycin D. Aphidicolin

allows initiation of DNA replication but leads to the stalling of

replication forks from which the replicative MCM helicase is un-

coupled, whereas actinomycin D inhibits progression of both

DNA polymerase and helicase (Pacek and Walter, 2004). In our

assays, aphidicolin reduced association of Sororin with chro-

matin partially, and actinomycin D inhibited this process largely,

even though Smc3 levels on chromatin were not reduced (Fig-

ure 2G). DNA replication is therefore required for efficient recruit-

ment of Sororin to chromatin. However, because aphidicolin and

actinomycin D inhibited DNA replication more efficiently than So-

rorin binding, it is possible that some Sororin can associate with

chromatin in the absence of DNA replication. Similar observa-

tions were made in HeLa cells where inhibition of DNA replication

by thymidine also reduced the Sororin-LAP levels on chromatin

(Figures S2G and S2H).

Cohesin Acetylation Facilitates but Is Not Sufficient forthe Association of Sororin with ChromatinBecause Sororin associates with chromatin during DNA replica-

tion, i.e., when cohesin is known to be acetylated, we analyzed

whether Smc3 acetylation and Sororin binding depend on each

other. To detect Smc3 acetylation, we used a monoclonal anti-

body that specifically recognizes Smc3 singly acetylated on

K106 or doubly acetylated on K105 and K106 (Figure S3A). We

observed that Sororin binding to chromatin and Smc3 acetyla-

tion occurred at the same time (Figure 2E) and that inhibition of

DNA replication had similar effects on both events, supporting

the notion that the two events are linked (Figure 2G). However,

depletion of Sororin from Xenopus extracts or from HeLa cells

affected neither the kinetics nor the degree of Smc3 acetylation,

suggesting that Sororin is not required for cohesin acetylation

(Figures S3B and S3C).

To test whether Smc3 acetylation is required for the chromatin

association of Sororin, we depleted Esco1 and Esco2 from HeLa

cells. Only depletion of both enzymes reduced Smc3 acetylation,

indicating that Esco1 and Esco2 can both acetylate cohesin

(Figure 3A). To analyze whether depletion of Esco1 and Esco2

affects the association of Sororin with chromatin, we synchro-

nized cells in S phase by double thymidine arrest-release and

measured the amount of Sororin-LAP on chromatin by immuno-

blotting and IFM. We also depleted endogenous Sororin in these

experiments to ensure that Sororin-LAP was analyzed under

conditions where it is functional. To rule out that reduced chro-

matin binding of Sororin was caused indirectly by a delay in

DNA replication, we labeled cells with BrdU and quantified

Sororin-LAP IFM signals only in cells that had similar amounts

of BrdU incorporated. Both by immunoblotting and IFM we

observed a reduction in Sororin on chromatin (Figures 3B–3D).

Depletion of Esco1 and Esco2 also reduced the amount of

endogenous Sororin that was associated with chromatin-bound

cohesin (Figures S3D and S3E). Esco1 and Esco2 are therefore

required for efficient binding of Sororin to cohesin on chromatin.

It is possible that the residual amounts of Sororin on chromatin

that were seen in our assays were due to incomplete depletion

of Esco1 and Esco2.

To address whether Esco1 and Esco2 regulate Sororin by

acetylating Smc3, we mutated K105 and K106 in Smc3 to either

glutamine (Smc3QQ), arginine (Smc3RR), or alanine (Smc3AA)

residues. Smc3QQ has been proposed to mimic acetylated and


Smc3RR and Smc3AA to mimic nonacetylated Smc3. We

mutated a LAP-tagged version of the Smc3 gene on a BAC,

stably integrated the modified BACs into HeLa cells, purified

the wild-type and mutant forms of Smc3 either from soluble

extracts or from chromatin, and analyzed their interaction part-

ners by immunoblotting and mass spectrometry. For wild-type

Smc3-LAP, these experiments confirmed that Sororin only asso-

ciates with cohesin on chromatin but not, or only to a small

degree, with soluble cohesin (Figure S3G). However, when

Smc3QQ-LAP was purified, Sororin could also reproducibly be

found in association with soluble cohesin, consistent with the

possibility that Smc3 acetylation promotes binding of Sororin

to cohesin (Figure 3E and Figure S3G). This interaction was abol-

ished by depletion of Scc1, indicating that Smc3QQ does not

simply represent a misfolded protein to which Sororin binds

nonspecifically (Figure S3H). Unexpectedly, similar results

were also obtained when Smc3RR and Smc3AA were analyzed

(Figures 3E, Figure S3F, and Figure S3G). This suggests that So-

rorin-cohesin interactions can be stabilized not only by muta-

tions that might chemically mimic acetylation but also by other

mutations that alter K105 and K106 (for possible interpretations

of these results, see Discussion).

We also attempted to generate acetylated cohesin in vitro by

using recombinant purified Esco1 (Figure S3I). We observed

that Esco1 could acetylate Smc3 when cohesin was associated

with chromatin in a Xenopus extract, but not in extract lacking

chromatin or when Esco1 was incubated with purified soluble

cohesin (Figure 3F and data not shown). Esco1 may therefore

only be able to modify cohesin on chromatin. Consistent with

this possibility, endogenous acetylated forms of Smc3 could

only be detected by immunoblotting in chromatin fractions (Fig-

ure S3J), and quantitative mass spectrometry indicated that the

fraction of acetylated Smc3 relative to total Smc3 is 97-fold

higher for chromatin-bound than for soluble cohesin (data not

shown).

When we added Esco1 to Xenopus extract containing chro-

matin, we observed that Smc3 acetylation was advanced by at

least 30 min, but Esco1 had no effect on the chromatin associa-

tion of Sororin (Figure 3F), indicating that Smc3 acetylation is not

sufficient for recruitment of Sororin to chromatin. In support of

Figure 3. Acetylation of Smc3 Facilitates

but Is Not Sufficient for the Association of

Sororin with Chromatin

(A) RNAi against both Esco1 and Esco2 causes

a decrease in Smc3 acetylation. HeLa cells were

transfected with siRNAs and harvested at S

phase. Chromatin-bound proteins were analyzed

by immunoblotting. Asterisks indicate nonspecific

signals.

(B) Reduction of Smc3 acetylation causes a

decrease of Sororin on chromatin. Sororin-LAP

HeLa cells were synchronized at S phase and

chromatin fractions were analyzed by immuno-

blotting.

(C) Cells in (B) were treated with BrdU after the

second thymidine release, pre-extracted, and

costained for BrdU, Sor-LAP (GFP), and DNA

(DAPI). Bar: 10 mm.

(D) Quantification of chromatin-associated

Sororin-LAP signal in cells with similar levels of

BrdU incorporation. Cells described in (C) with

similar BrdU intensities (left) were analyzed for

Sor-LAP intensity (right) (mean ± confidence

interval; *p < 0.01).

(E) Soluble Smc3QQ and Smc3RR proteins stably

bind to Sororin in HeLa cells. HeLa cells express-

ing Smc3WT-, Smc3QQ-, or Smc3RR-LAP were

synchronized in G2 phase, Smc3-LAP was immu-

noprecipitated from the soluble fraction of cells,

and the coprecipitated proteins were analyzed

by immunoblotting using a 2-fold serial dilution.

(F) Acetylation of Smc3 is not sufficient for Sororin

binding to chromatin. Interphase Xenopus egg

extracts were incubated with sperm chromatin

in the presence (Esco1) or absence (buffer) of

Esco1 for indicated times. Chromatin fractions

were analyzed by immunoblotting (on chromatin).

Extracts without sperm chromatin were incubated

for 120 min in the presence or absence of Esco1

(extracts).

See also Figure S3.


this hypothesis, we found that the association of Sororin with

Smc3QQ was still partially dependent on DNA replication (Fig-

ure S3K). Taken together, these results indicate that Smc3 acet-

ylation is required but not sufficient for efficient recruitment of

Sororin to chromatin-bound cohesin.

Sororin Is Dispensable for Cohesion in the Absenceof WaplSeveral previous observations are consistent with the possibility

that Sororin and Wapl have antagonistic functions: depletion of

Sororin and Wapl has opposite effects on cohesion (increased

and decreased proximity between sister chromatids, respec-

tively) and on the stability of cohesin-DNA interactions

(decreased and increased residence times of cohesin on chro-

matin, respectively). Likewise, addition of excess Sororin to Xen-

opus extracts mimics the ‘‘overcohesion’’ phenotype caused by

depletion of Wapl, and overexpression of Wapl causes cohesion

defects, as does loss of Sororin (Gandhi et al., 2006; Kueng et al.,

Figure 4. Sororin Is Dispensable for Cohe-

sion in the Absence of Wapl

(A) Chromatin fractions from mock-, Sororin-,

Wapl-, and Wapl- and Sororin-depleted inter-

phase extracts were analyzed by immunoblotting.

(B) D90 Cyclin B was added to the extracts shown

in (A) and mitotic chromosomes were assembled.

Chromosomes were isolated 120 min after D90

Cyclin B addition and stained for XCAP-E

(magenta) and Bub1 (green). Higher-magnification

images are shown in lower panels. Distance

between two chromosome arms stained by

XCAP-E in each extract is shown in a histogram

as a comparison to the mock-depleted extract.

Depletion of SA1/2 is shown as an example of

cohesin depletion. Bar: 5 mm.

(C) Codepletion of Sororin and Wapl in HeLa cells.

Cells were transfected with the indicated siRNAs

and treated with nocodazole. After mitotic shake-

off for chromosome spreads (D and E), residual

cells were harvested for immunoblotting. See

also Figure S4A.

(D) Analysis of chromosome spreads after Sororin

and Wapl depletion. Mitotic cells harvested as in

(C) were examined by chromosome spreading

and Giemsa staining. Five hundred cells per RNAi

experiment were classified into three categories.

(E) Representative pictures of the most prominent

phenotype class upon RNAi depletion in the

Giemsa spread analysis. Color code is shown in

(D). Bar: 10 mm.

2006; Rankin et al., 2005; Schmitz et al.,

2007; Shintomi and Hirano, 2009).

To understand the functional relation-

ship between Sororin and Wapl we

depleted both proteins either individually

or simultaneously from Xenopus extracts

and analyzed cohesion in mitotic chro-

mosomes. We analyzed chromosome

morphology by staining the condensin

subunit XCAP-E and Bub1 as markers

for sister chromatid arms and kinetochores, respectively. Deple-

tion of Sororin alone increased the distance between sister chro-

matids, indicating a partial cohesion defect (Figures 4A and 4B).

This defect was similar in magnitude to the defect that was

observed after simultaneous depletion of the cohesin subunits

SA1 and SA2, suggesting that also in Xenopus extracts Sororin

is similarly important for cohesion as cohesin itself (Figure 4B).

As expected, depletion of Wapl had the opposite effect, i.e., re-

sulted in tightly connected chromatids. Remarkably, depletion of

both proteins caused a phenotype that was very similar to the

phenotype caused by depletion of Wapl alone. Similar results

were obtained when Sororin and Wapl were depleted singly or

simultaneously by RNAi from HeLa cells and mitotic chromo-

somes were analyzed by Giemsa staining (Figures 4C–4E).

Also in this case, the phenotype obtained after codepletion of

Sororin and Wapl was nearly identical to the phenotype obtained

after depletion of Wapl alone, i.e., in the majority of mitotic cells

sister chromatids were more tightly associated with each other


than normally. These observations indicate that Sororin is only

required for cohesion in the presence of Wapl, and they therefore

suggest that Sororin’s key function is to antagonize Wapl.

We also observed in these experiments that Wapl depletion

greatly increased the degree of Smc3 acetylation (Figure 4C

and Figure S4A). Wapl depletion could cause this effect by

increasing the residence time of cohesin on DNA, but it is also

possible that Wapl inhibits cohesin acetylation and that this func-

tion is required for Wapl’s ability to allow cohesin dissociation

from DNA.

Sororin Interacts with Pds5 via a Conserved FGF Motifand Can Displace Wapl from Pds5When we isolated Sororin-LAP via tandem affinity purification,

we identified cohesin core subunits and Pds5A and Pds5B, indi-

cating that Sororin can directly bind to these proteins (Figure S1I

and data not shown). Sororin antibodies also immunoprecipi-

tated Pds5A and Pds5B from solubilized chromatin of HeLa cells

(Figure S4B), and when we immunodepleted Pds5A and Pds5B

from Xenopus extracts the binding of Sororin to chromatin was

greatly reduced (Figure 5A). The latter effect was not caused

by a delay in DNA replication because [a-32P]dCTP incorporation

into sperm DNA was unaffected by depletion of Pds5 proteins

(Figure 5B). These observations are consistent with the possi-

bility that the association of Sororin with cohesin depends on

Pds5 proteins.

To address directly whether Sororin interacts with Pds5

proteins or Pds5-Wapl heterodimers, we purified recombinant

forms of human Sororin, Pds5A and Wapl. As predicted, Wapl

bound efficiently to Pds5A, either when expressed simulta-

neously in Baculovirus-infected insect cells or when incubated

with each other as individually purified proteins (Figure S4C

Figure 5. The FGF Motif of Sororin Is

Required for Cohesion

(A and B) Pds5 is required for Sororin association

with chromatin. Interphase Xenopus egg extracts

were subjected to mock or Pds5A and B immuno-

depletion. Two hours after sperm chromatin addi-

tion, chromatin fractions were analyzed by immu-

noblotting (A). DNA replication in the extracts

shown in (A) was monitored for 30 or 60 min by

incorporation of [a-32P]dCTP into sperm chro-

matin (B).

(C) Sequence comparison in the region including

FGF motifs of vertebrate Sororin and fly Dalmatian.

Identical and similar residues are shaded in black

and gray, respectively. In Xenopus, Sororin-A is

shown. In fruit fly, the latter two of three FGF motifs

are shown (see also Figure 7A).

(D) Anti-Pds5A antibody beads were incubated

with Sororin-WT or -AA mutant in the presence

or absence of Pds5A protein. Beads-bound pro-

teins were analyzed by immunoblotting.

(E) Anti-Pds5A antibody beads were incubated

with Sororin-WT or -AA mutant in the presence or

absence of the Pds5A-Wapl heterodimer. Beads-

bound proteins were separated from the superna-

tant and were analyzed by immunoblotting.

(F) Wapl removal activity of Sororin is increased in

a dose-dependent manner. Increasing amounts

(10–40 ng/ml) of Sororin-WT or -AA mutant were

used in the experiment shown in (E), supernatant

fractions were analyzed by immunoblotting (left),

and signal intensity of Wapl was quantified (right).

(G) Sororin-depleted interphase extracts were

combined with buffer, Sororin-WT, or -AA mutant.

Two hours after sperm chromatin addition, chro-

matin fractions were analyzed by immunoblotting.

(H) D90 Cyclin B was added to the extracts shown

in (G) and mitotic chromosomes were assembled.

Chromosomes were isolated 120 min after

D90 Cyclin B addition and stained for XCAP-E.

Magnified images are shown in lower panels.

Distance between two chromosome arms stained

by XCAP-E is shown in lower histogram as a

comparison to mock-depleted extract. Bar: 5 mm.

See also Figure S4.


and data not shown). The interaction between Pds5 and Wapl

depends on two sequence elements composed of phenylala-

nine-glycine-phenylalanine (FGF) residues in Wapl (Shintomi

and Hirano, 2009), and we noticed that a similar FGF motif is

also present at a conserved position in all known Sororin

sequences (Figure 5C and see Figure S5B). We therefore also

generated a Sororin mutant in which the two phenylalanine resi-

dues in this motif were changed to alanines (hereafter called

‘‘Sororin-AA’’). Wild-type Sororin associated with Pds5A,

whereas the AA mutant bound less well (Figure 5D). Also,

when added to Xenopus extracts, wild-type Sororin associated

with cohesin and Pds5B more efficiently than the AA mutant (Fig-

ure S4D). When we performed Sororin-binding experiments with

Pds5A-Wapl, we observed, remarkably, that Sororin displaced

some Wapl from the Pds5A-Wapl heterodimers. Also, this effect

was reduced when the AA mutant was used (Figures 5E and 5F).

These observations raised the possibility that Sororin regulates

cohesin by interacting with the Pds5-Wapl heterodimer.

The FGF Motif of Sororin Is Essential for Its CohesionFunctionTo address whether Sororin’s ability to displace Wapl from Pds5

is of functional relevance, we replaced Sororin in Xenopus

extracts by the Sororin-AA mutant and analyzed its effect on

cohesion. We immunodepleted Sororin from interphase egg

extracts, added either buffer, recombinant wild-type Sororin,

or the AA mutant, and analyzed mitotic chromosomes as above.

Importantly, the cohesion defect observed after Sororin deple-

tion could be restored by wild-type Sororin but not by the AA

mutant (Figures 4G and 4H). Similar results were obtained

when excess Sororin was added to Xenopus extracts from which

the endogenous protein had not been depleted: in this assay

wild-type Sororin causes an ‘‘overcohesion’’ phenotype (Rankin

et al., 2005), but the AA mutant had no effect (Figure S4E). These

results show that the FGF motif of Sororin is required for its func-

tion in cohesion, and they suggest that Sororin might execute

this function by displacing Wapl from Pds5.

However, we could not obtain evidence that the Sororin-

dependent displacement of Wapl from Pds5 results in the disso-

ciation of Wapl from chromatin. Addition of recombinant Sororin

to Xenopus extracts increased, and did not decrease, the

amount of Wapl and Pds5A on chromatin, as if Sororin stabilized

the interactions between Pds5A-Wapl and cohesin, rather than

dissociating Wapl from cohesin (Figure S4F). It is therefore

possible that the Sororin-mediated displacement of Wapl from

Pds5A causes a rearrangement in the topology of cohesin-asso-

ciated proteins and does not lead to dissociation of Wapl from

cohesin.

Sororin Is Inactivated by Phosphorylation in MitosisThe prophase pathway of cohesin dissociation depends on Wapl

(Gandhi et al., 2006; Kueng et al., 2006). It is therefore conceiv-

able that Sororin has to be inactivated at the onset of mitosis

to relieve Wapl from its inhibition by Sororin. We therefore

analyzed whether Sororin’s ability to dissociate Wapl from

Pds5 proteins is cell cycle regulated. Consistent with this possi-

bility, recombinant Sororin could associate with Pds5B in Xeno-

pus interphase extracts but not in mitotic extracts where Sororin

is phosphorylated (Figure 6A). Furthermore, we observed that

Sororin could bind to recombinant purified Wapl-Pds5A

Sororin

FGF

S/G2-phase M-phaseTelo/G1-phase

DNA replication

dynamic

Pds5

mitotic entry

stable

WaplPds5

SororinWap

l

Smc3 acetylation

dynamic

Pds5

WaplSororinP

I-S

or

M-S

or λ

PP

supernatant

Sororin

Wapl

Pds5A

M-S

or

I-S

or

M-S

or

M-S

or λ

PP

beads bound

buffe

r

buffe

r

A B

C

I MI M

Sororin

Pds5B

: human Sororin− ＋

*

Figure 6. Phosphorylated Sororin Is Unable

to Dissociate Wapl from Pds5

(A) Sororin is dissociated from Pds5 in mitosis.

Sororin-WT was incubated in either interphase (I)

or mitotic (M) Xenopus egg extracts and immuno-

precipitated, and the precipitates were analyzed

by immunoblotting. Asterisk indicates nonspecific

signal.

(B) Wapl removal activity is abolished by phosphor-

ylation of Sororin. Wapl-Pds5A heterodimer on anti-

Pds5A antibody beads was incubated with either

buffer, Sororin preincubated in interphase egg

extract (I-Sor) or mitotic egg extract (M-Sor), or

l-protein phosphatase-treated M-Sor (M-Sor

l-PP). Beads-bound proteins were separated from

the supernatant and analyzed by immunoblotting.

(C) Model for the role of Sororin in sister chromatid

cohesion. The cohesin complex is loaded onto

chromatin during telo/G1 phase, where Wapl-

Pds5 destabilizes cohesin binding to chromatin

in the absence of Sororin. During DNA replication

in S phase, Sororin associates with chromatin

depending on cohesin and this association is facil-

itated by acetylation of Smc3. Sororin binds to

Pds5 through its FGF motif and thereby can antag-

onize the function of Wapl by modulating the

topology of Wapl and Pds5 so that stable cohesion

is maintained. Upon entry into mitosis, phosphory-

lation of Sororin abolishes the ability to inhibit

Wapl, allowing cohesin removal in prophase.


heterodimers and dissociate Wapl from Pds5A when Sororin

was preincubated in Xenopus interphase extracts but not when

Sororin had been incubated in a mitotic extract (Figure 6B).

The Wapl dissociation activity of mitotic Sororin was fully

restored when Sororin was first dephosphorylated by l-protein

phosphatase. These results suggest that Sororin phosphoryla-

tion in mitosis relieves Wapl from inhibition by Sororin (Figure 6C;

for further discussion of this model see below).

Dalmatian Is a Drosophila Ortholog of SororinWapl orthologs exist in species from yeast to human (Kueng

et al., 2006), but Sororin has only been identified in vertebrates

(Rankin et al., 2005). To address whether inhibition of Wapl by

Sororin could also be required for cohesion in nonvertebrate

species, we searched for Sororin-related sequences in inverte-

brate genomes (Table S1). BLAST searches identified Sororin

Figure 7. Dalmatian Is an Ortholog of So-

rorin in Drosophila

(A) Schematic sequence comparison of human

and Xenopus Sororin and Drosophila Dalmatian.

The conserved regions are shaded in gray and

KEN-box and FGF motifs are depicted with white

and black boxes, respectively.

(B) Dalmatian (Dmt) RNAi causes premature sister

chromatid separation in S2 cells. Cells were trans-

fected with dsRNA against Dmt or BubR1 or were

left untransfected (control). Chromosome spreads

were stained with DAPI. Representative images

are shown. Bar: 5 mm.

(C) Cells described in (B) were quantified for loss of

cohesion. Error bars denote standard deviations

between three independent experiments.

(D) Mitotic defects in Dalmatian knockdown cells.

Cells were transfected with dsRNA against Dmt

or BubR1 or were untransfected (control) and

costained for a-tubulin and Cyclin B to define

mitotic stages, CID (Cenp-A in Drosophila) to

assess centromere pairing, and DAPI (upper

panel). The lower table summarizes the observed

phenotype over all mitotic cells (n > 59 per condi-

tion). Bar: 5 mm.

See also Figure S5 and Table S1.

sequences in vertebrates and one

distantly related protein in the mollusc

Lottia gigantea. We used the C-terminal

portion of these sequences, where the

highest degree of similarity is found, to

perform iterative rounds of similarity

searches in invertebrate proteome data-

bases. We identified a single sequence

with significant similarity to Sororin in 18

different metazoan species belonging to

different taxa, including cephalochor-

dates, echinoderms, insecta, cnidaria,

and placozoa. All of these proteins

contain sequences related to the C

terminus of Sororin, which we therefore

call the ‘‘Sororin domain’’ (Figure S5A). Furthermore, 17 of these

proteins also contain an FGF sequence motif (Figure S5B), or

sometimes several of these motifs (Figure 7A).

Of the 18 hypothetical proteins, only one has previously been

characterized. This is a 95 kDa protein called Dalmatian, which is

required for development of theDrosophila embryonic peripheral

nervous system (Prokopenko et al., 2000). Recent RNAi screens

have shown that depletion of Dalmatian causes defects in mitotic

spindle assembly, chromosome alignment, and cell division

(Goshima et al., 2007; Somma et al., 2008). Dalmatian inactiva-

tion also causes precocious sister chromatid separation in the

presence of colchicine, a compound that activates the SAC. It

has therefore been proposed that Dalmatian is required for the

SAC (Somma et al., 2008).

Because Dalmatian shares sequence similarity with Sororin,

we tested whether Dalmatian is required for cohesion. If this


were the case, Dalmatian depletion would be predicted to cause

precocious sister chromatid separation, to activate the SAC, and

thus to cause an increase in mitotic index, whereas inactivation

of a SAC protein would shorten mitosis and cause a decrease

in mitotic index. We observed a modest increase in mitotic index

from 3.2% in control Drosophila S2 cells to 5.3% in Dalmatian

RNAi cells, whereas depletion of BubR1, a protein required for

the SAC (Perez-Mongiovi et al., 2005), decreased the mitotic

index to 1.4% (data not shown). Chromosome spreading re-

vealed that cohesion had been lost in 82% of all mitotic Dalma-

tian RNAi cells, but only in less than 6% of mitotic control or

BubR1 RNAi cells (Figures 6B and 6C). In IFM experiments, we

observed that Dalmatian depletion caused chromosome con-

gression defects (‘‘scattered chromosomes’’) in 57.6% of prom-

eta/metaphase cells (Figure 6D). Many of the scattered chromo-

somes were single sister chromatids, as judged by staining of

the centromere protein centromere identifier (CID), and Cyclin

B levels were similarly high in cells with scattered chromatids

as in control prometaphase cells. Because SAC defects would

lead to precocious APC/CCdc20 activation and Cyclin B degrada-

tion, these results indicate that Dalmatian depletion does not

inactivate the SAC. Instead, our results suggest that Dalmatian

is a distant ortholog of Sororin that is required for cohesion.

DISCUSSION

Although establishment and maintenance of sister chromatid

cohesion are essential for chromosome segregation, it is poorly

understood how cohesin generates cohesive structures during

DNA replication and how these are maintained for hours, or in

the case of mammalian oocytes even for years. Recent studies

have revealed that both the stability of cohesin-DNA interactions

(Gerlich et al., 2006) and the acetylation state of cohesin change

during DNA replication (Ben-Shahar et al., 2008; Rowland et al.,

2009; Unal et al., 2008; Zhang et al., 2008), suggesting that cohe-

sion is not simply established by doubling the number of sister

chromatids inside otherwise unchanged cohesin rings. Our

results further extend this view by showing that also the compo-

sition of cohesin complexes changes during DNA replication

through the recruitment of Sororin, and importantly our data

suggest that only Sororin-associated cohesin complexes are

able to mediate cohesion. Consistent with this view, we find

that Sororin is the only known protein whose presence on chro-

matin coincides precisely with the periods of the cell cycle during

which cohesion exists (from initiation of DNA replication to meta-

phase), whereas cohesin binds to DNA long before cohesion is

established.

Based on our results, we propose the following model for how

Sororin enables cohesin to become ‘‘cohesive’’ (Figure 6C):

Smc3 acetylation and possibly other unidentified events during

DNA replication promote the recruitment of Sororin to chro-

matin-bound cohesin. These events might occur directly at repli-

cation forks because Eco1 has been detected at these sites

(Lengronne et al., 2006), Smc3 can only be acetylated on chro-

matin (Unal et al., 2008; this study), and actinomycin D,

a compound that inhibits DNA polymerase and MCM helicase

progression (Pacek and Walter, 2004), prevents both Smc3 acet-

ylation and Sororin recruitment. Because Smc3 acetylation and

Sororin recruitment are blocked less efficiently by aphidicolin

and thymidine, in whose presence helicase progression can still

occur, it is possible that Smc3 acetylation and Sororin binding

are coupled to helicase progression. Within the cohesin

complex, Sororin binds to Pds5 via an FGF sequence motif

that is shared between Sororin and Wapl. Sororin displaces

Wapl from Pds5, but not from cohesin, suggesting that Sororin

induces a rearrangement in the topology of these cohesin-

associated proteins. We propose that these changes inhibit

Wapl’s ability to dissociate cohesin from DNA, and that the re-

sulting stable interaction of cohesin with DNA enables cohesin

to mediate cohesion. Our data further indicate that in prophase,

Sororin is inactivated by phosphorylation, enabling Wapl to

dissociate cohesin from mitotic chromosomes. Later in telo-

phase and G1, APC/CCdh1 targets Sororin for degradation. The

function of this process remains to be understood, but it might

ensure that Sororin associates with cohesin only after the initia-

tion of DNA replication once APC/CCdh1 has been inactivated.

This model makes a number of important predictions: (1) If So-

rorin is an antagonist of Wapl, one would expect that Sororin or-

thologs can be identified in species where Wapl exists. We show

that this is indeed the case for many metazoans, including

species from evolutionarily old taxa such as cnidaria (jellyfish)

and placozoa, the simplest known metazoa. Our observation

that depletion of the Drosophila member of this protein family

(Dalmatian) causes cohesion defects suggests that these

proteins are also functionally related to Sororin. We have so far

not been able to identify Sororin-related proteins in worms or

yeast. It therefore remains to be seen whether Sororin is required

for cohesion in all eukaryotes, or whether some species have

evolved cohesion mechanisms that are independent of Sororin.

(2) If the key function of Sororin is to inhibit Wapl, then Sororin

is expected to be dispensable in the absence of Wapl. Our

results indicate that this is indeed the case. An interesting impli-

cation of this result is that Sororin might not be essential for the

initial entrapment of sister chromatids by cohesin rings, i.e., for

cohesion establishment, at least in the absence of Wapl. It is

therefore possible that Sororin’s main function is to prevent

dissociation of cohesin from DNA, rather than enabling opening

and closure of the ring around DNA. However, the situation could

be different in yeast where deletion of WAPL/RAD61 does not

result in accumulation of cohesin on DNA but has the opposite

effect, a reduction of cohesin on chromatin (Rowland et al.,

2009; Sutani et al., 2009). If a Sororin-related Wapl/Rad61 antag-

onist exists in yeast, such a protein (or protein domain) might

therefore instead be needed for cohesion establishment by

having to overcome the proposed ‘‘anti-establishment’’ activity

of Wapl/Rad61 (Rowland et al., 2009; Sutani et al., 2009).

(3) If the stable postreplicative association of cohesin with

DNA was due to inhibition of Wapl by Sororin, depletion of

Wapl should enable cohesin to bind to DNA also stably before

Sororin has been recruited to cohesin, i.e., in G1 phase. At vari-

ance with this prediction, we observed previously that depletion

of Wapl from HeLa cells increased the residence time of dynam-

ically bound cohesin complexes only modestly, from 8 min in

control cells to 18 min (Kueng et al., 2006), and not to many

hours, as is normally seen for cohesin complexes in G2 phase

(Gerlich et al., 2006). However, we have in the meantime


measured the residence time of cohesin on chromatin in mouse

embryonic fibroblasts from which the Wapl gene has been

deleted, and in which therefore a more complete depletion of

Wapl can be achieved than by RNAi. In these cells the residence

time of cohesin on chromatin is increased from minutes to

several hours even before S phase (A. Tedeschi, personal

communication), indicating that it is indeed the presence of

Wapl that enables cohesin to interact with DNA dynamically

before replication. This result supports the hypothesis that inhi-

bition of Wapl by Sororin enables stable binding of cohesin to

DNA in postreplicative cells.

Our model also raises several important new questions. One of

them is whether the essential function of Smc3 acetylation is to

recruit Sororin, or whether this modification has other important

effects, for example on the ATPase activity of Smc3. The

absence of Sororin in yeast would suggest that cohesin acetyla-

tion must have other essential functions, but given the low

sequence similarity among Sororin orthologs it cannot be

excluded that Sororin-related proteins also exist in yeast.

A related important question is how Smc3 acetylation might

promote recruitment of Sororin. As Pds5 proteins are required

for the recruitment of Sororin to cohesin, and Sororin binds to

Pds5 proteins, we suspect that Smc3 acetylation promotes So-

rorin binding indirectly, possibly by causing changes in how

Pds5 or Wapl interact with cohesin or each other. Likewise, it

is unclear why replacement of K105/106 to not only glutamine

(which is believed to mimic acetylated lysine) but also to arginine

or alanine residues can stabilize cohesin-Sororin interactions. It

is possible that it is not the presence of acetyl residues on

K105/106 that creates a binding site, for example for a cohesin

subunit, but that any mutation that removes lysines at these posi-

tions will destroy a binding site or pocket, which would lead to

subunit rearrangements that would facilitate Sororin recruitment.

A more detailed characterization of how cohesin interacts with

Wapl, Pds5, and Sororin will be required to address these

questions.


Immunodepletion and Monitoring of DNA Replication in Xenopus

Egg Extracts

For immunodepletion of Xenopus egg extracts, affinity-purified antibody (70

mg anti-Sororin, mixture of 40 mg anti-Pds5A and 25 mg anti-Pds5B, 200 mg

anti-Wapl, or 250 mg anti-SA1/2) was conjugated to 30 ml Affi-Prep Protein A

Matrix (Bio-Rad), mixed with 100 ml interphase extracts, incubated for 30

min for Sororin depletion or 1 hr for Pds5A/B, Wapl, and SA1/2 depletions

on ice, and beads were removed by centrifugation. For add-back experiments,

Sororin wild-type or AA mutant (F166A, F168A) was added to Sororin-depleted

extracts at 6.5 nM.

DNA replication was monitored by the incorporation of [a-32P]dCTP into

DNA. Demembranated sperm nuclei (2000 nuclei/ml) were added to egg

extract containing [a-32P]dCTP (3.7 kBq/ml), incubated at 22�C, and the reac-

tion stopped by addition of 2 volumes of stop solution (8 mM EDTA, 0.13%

phosphoric acid, 10% Ficoll, 5% SDS, 0.2% bromophenol blue, 80 mM Tris-

HCl pH 8.0). The mixture was incubated with 2 mg/ml Proteinase K for 30

min at 37�C and analyzed by agarose gel electrophoresis followed by

autoradiography.

Preparation of Xenopus Chromatin Fractions

Sperm nuclei were incubated in extracts at a concentration of 2000 nuclei/ml.

Thirty microliters of extract was diluted 10-fold with ice-cold extract buffer (EB;

5 mM MgCl2, 100 mM KCl, HEPES-KOH pH 7.5) containing 0.25% Triton X-

100, overlaid onto a 30% sucrose/EB cushion, and spun at 15,000 g for 10

min. The pellets were washed with EB containing 0.25% Triton X-100 and re-

suspended in SDS sample buffer.


Supplemental Information includes Extended Experimental Procedures, five

figures, and one table and can be found with this article online at doi:10.

1016/j.cell.2010.10.031.

ACKNOWLEDGMENTS

We are grateful to O. Hudecz, P. Huis in’t Veld, I. Poser, and M. Sykora for

assistance and reagents; to G. Karpen, J. Knoblich, C. Lehner, and C. Sunkel

for reagents and advice on Drosophila experiments; and to N. Kraut for BI

2536. T.N. is supported by the European Molecular Biology Organization

(EMBO) and the Japanese Society for the Promotion of Science (JSPS). K.S.

is supported by Grant-in-Aid for Scientific Research (S). Research in the

groups of J.-M.P. and K.M. is supported by Boehringer Ingelheim and the

Austrian Science Fund via the special research program ‘‘Chromosome

Dynamics’’ (F34-B03). Work in the groups of J.-M.P., K.M., and A.A.H. was

also supported by the EC via the Integrated Project ‘‘MitoCheck.’’





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Nonenzymatic Rapid Controlof GIRK Channel Functionby a G Protein-Coupled Receptor KinaseAdi Raveh,1 Ayelet Cooper,1 Liora Guy-David,1 and Eitan Reuveny1,*1Department Biological Chemistry Weizmann Institute of Science, Rehovot 76100, Israel


DOI 10.1016/j.cell.2010.10.018

SUMMARY

G protein-coupled receptors (GPCRs) respond toagonists to activate downstream enzymatic path-ways or to gate ion channel function. Turning offGPCR signaling is known to involve phosphorylationof the GPCR by GPCR kinases (GRKs) to initiate theirinternalization. The process, however, is relativelyslow and cannot account for the faster desensitiza-tion responses required to regulate channel gating.Here, we show that GRKs enable rapid desensitiza-tion of the G protein-coupled potassium channel(GIRK/Kir3.x) through a mechanism independent oftheir kinase activity. On GPCR activation, GRKstranslocate to the membrane and quench channelactivation by competitively binding and titrating Gprotein bg subunits away from the channel. Ofinterest, the ability of GRKs to effect this rapid desen-sitization depends on the receptor type. The findingsthus reveal a stimulus-specific, phosphorylation-independent mechanism for rapidly downregulatingGPCR activity at the effector level.

INTRODUCTION

G protein-coupled receptors (GPCR) modulate the activity of

enzymes and ion channels to fine tune cellular activity (Pierce

et al., 2002). To avoid abnormal cellular activity, GPCR-mediated

G protein cycles should be temporally precise. Several mecha-

nisms guarantee the precise length of GPCR activation by

controlling the levels of agonist. For example, the level of free

neurotransmitters present in the synapse are limited by fast

neurotransmitter reuptake at the presynaptic site (Torres et al.,

2003), or degradation at the synaptic cleft (Massoulie et al.,

1993). These processes are specific for specific types of ligands.

For regulation at a longer time scale, additional mechanisms

control GPCR signaling efficacy. These mechanisms control

the robustness of the activation signals by regulating receptor

number at the plasma membrane, in a process termed downre-

gulation (Bunemann et al., 1999; Tsao and von Zastrow, 2000).

This mechanism involves a receptor-mediated signaling

cascade, where activated receptors are initially phosphorylated

by GPCR kinases (GRKs), to initiate intracellular events leading

to a clathrin-mediated endocytosis of the GPCRs. This process

occurs over a time scale of many minutes to hours.

In the context of GPCR-mediated regulation of ion channel

activity, short-term desensitization to an activating signal has

been observed. For instance, regulation of GPCR-controlled

excitability through the activation of the G protein-coupled

potassium channels (GIRK/Kir3.x), displays short-term desensi-

tization characterized by a reduction in channel currents in the

presence of the receptor agonist in a time scale of few seconds

(Sickmann and Alzheimer, 2003). This short-term reduction in

postsynaptic GIRK channel activity is independent of elements

that are known to affect the G protein cycle and PtdIns(4,5)P2

hydrolysis. It is, therefore, of great interest to identify the molec-

ular mechanism that mediates this process.

We set out to identify the mechanism responsible for short-

term desensitization of GIRK channels. We found that for some

GPCRs, continued activation of their receptors leads to GIRK

current desensitization (GCD). This current desensitization is

enhanced in the presence of GRK2 and, surprisingly, does not

involve its kinase activity, but rather depends on its ability to

bind the Gbg subunits of the G protein. This binding appears

to compete for the available pool of the G protein subunits that

activate the channel and hence to effectively quench channel

activity. These findings assign a new role for the GRK proteins

in providing negative feedback control of GPCR function at the

effector level.

RESULTS

GRK2 Accelerates Desensitization of GIRK CurrentsInduced by A1R and mOR, but Not by mGluR2 and M4RWe set out to test the involvement of GRK2 in mediating short-

term desensitization of GIRK channels. GRK2 is involved in the

desensitization of GPCRs after exposure to their agonists. For

this purpose we expressed GIRK1, GIRK4 (for now on referred

as GIRK channels) and adenosine type 1 receptor (A1R) with or

without (control) GRK2 in HEK293 cells, and used whole cell

patch-clamp recordings to measure various channel current

parameters after receptor activation by adenosine (Figure 1A).

After A1R activation by adenosine (100 mM), GIRK channel

currents desensitize (GCD) as evident from the monoexponential


decay curve of the current traces with a time constant of 24.9 ±

11.1 s, n = 8 (Figures 1A, upper trace, and 1C). Interestingly, in

cells cotransfected with GRK2, GCD rates were accelerated

�10-fold, to 2.6 ± 0.0 s, n = 9 (p < 0.05). To assess whether

the enhancement of current desensitization was a general

phenomena to all PTX-sensitive GPCRs, we also tested GCD

rates induced by m-opioid receptor (mOR). Similar to the effect

of GRK2 on A1R-mediated GCD, mOR activation (methionine

enkephalin, ME, 100 nM) accelerated GCD in the presence of

GRK2 compared to control cells, with a time constant of 38.9 ±

5.9 s, n = 10 and 64.4 ± 6.18 s, n = 7, respectively (see Figures

S1A and S1C available online). In contrast, activation of GIRK

channels in the absence or presence of GRK2 by metabotropic

glutamate type 2 receptor (mGluR2) (Figures 1B and 1C) or

muscarinic acetylcholine type 4 receptor (M4R) activation

(Figures S1B and S1C) did not show any acceleration in GCD,

with time constants 41.7 ± 8.6 s, n = 9 and 41.7 ± 9.5 s, n = 9,

0

20

40

60

80

100

NT siGRK2 #1 siGRK2 #2 siGRK2 #1 + smGRK2GFP

Rescue

% C

urre

nt (@

2 m

)

0

20

40

60

0

50

100

shGRK2 NT

GK2

(%)

A

B

shRNA NTGRK2

GFP

A1R

mG

luR

2

C

D

10s1000pA

mGluR2A1R

+GR

K2+G

RK2

NTsiGRK2 #1

F

G

E

0

20

40

60

80

100

NT siGRK2 #1 siGRK2 #2

GR

K2 R

NA

(%, n

orm

aliz

ed)

10s100 pA

Figure 1. GRK2 Accelerates the Desensiti-

zation of GIRK Currents Induced by A1R,

but Not by mGluR2

(A) GIRK channel currents induced by the activa-

tion of A1R rapidly desensitize in the presence of

GRK2.

(B) GIRK channel currents induced by mGluR2

activation are insensitive to GRK2.

(C) Bar plot that depicts GCD rates of cells acti-

vated with A1R or mGluR2 without or with GRK2,

GRK2 shRNA, or nontarget (NT) shRNA.

(D) Bar plot compares the normalized expression

levels of GRK2 in silenced and NT cells as de-

picted from western blot for GRK2 (inset).

(E) GIRK current traces induced by adenosine in

control HL-1 cell (black) and of siRNA#1 silenced

cell (gray).

(F) Bar plot depicting GCD in HL-1 cells trans-

fected with two independent siRNAs, NT, and

siRNA#1 transfected cells rescued by the expres-

sion of silently mutated GRK2GFP (smGRK2GFP).

(G) GRK2 mRNA quantification in HL-1 cells trans-

fected with two independent siRNAs or NT control.

See also Figure S1.

respectively for mGluR2, and 37.7 ±

10.7 s, n = 7 and 33.4 ± 11.7 s, n = 6,

respectively, for M4R. Like in the case

shown above for GRK2, GRK3, but not

GRK6, also accelerated GCD in a similar

receptor-specific manner (data not

shown).

Because GRK2 is endogenously ex-

pressed in HEK293 cells (Violin et al.,

2006), we were interested to know

whether there is a contribution of the

endogenous protein to current desensiti-

zation in cells not transfected with GRK2.

To address this question we silenced

endogenous GRK2 levels using shRNA

specific for the human GRK2 (shGRK2).

GRK2 expression levels were reduced

by 58%, as determined using western blot (Figure 1D). A1R-in-

duced GIRK currents were significantly slower in GRK2-silenced

cells (42.9 ± 6.8 s, n = 12) in comparison with cells cotransfected

with nontarget (NT) shRNA (26.0 ± 4.5 s, n = 12) (Figure 1C), con-

firming that endogenous levels of GRK2 are sufficient to enhance

GCD rate after A1R simulation. The above results suggest that

GRK2 has a role in modulating current desensitization rates of

GIRK currents in a receptor-selective manner.

To study whether GRK is also involved in GCD in cells that

natively express GIRK, A1R and the kinase, we measured

GIRK currents in HL-1 cells. HL-1 is a mouse cardiac muscle

cell-line that maintains the characteristics of adult cardiac myo-

cytes, including contraction (Claycomb et al., 1998). These cells

express both GIRK channels and the necessary components for

their activation (Nobles et al., 2010). GIRK currents of HL-1 cells,

where GRK2 was silenced using two independent siRNAs,

(siGRK2#1 and siGRK2#2) displayed significantly smaller


desensitizations compared to cells transfected with NT (Figures

1E and 1F). After continuous application of adenosine, the

induced currents were reduced to 79.2 ± 11.0% (n = 6), 86.3 ±

7.3% (n = 5) and 24.7 ± 7.4% (n = 6) at 2 min, for both silenced

and NT cells, respectively. Expression of silently mutated

GRK2-GFP (smGRK2-GFP) in cells silenced with siGRK2#1

rescued the reduction in current desensitization (31.5 ± 12.5%,

n = 4) to levels comparable to NT cells (Figure 1F). Similarly,

GRK2 mRNA levels were reduced in cells transfected with either

siGRK2#1 or siGRK2#2 compared to NT control cells with 54.0 ±

2.4% and 57.1 ± 0.6%, respectively (Figure 1G). Qualitatively

similar results were obtained using primary mouse hippocampal

neurons (Figure S1). These experiments suggest that, qualita-

tively, the effect of GRK in HEK cells is relevant at physiological

expression levels, and is not due to overexpression of GRK, the

receptors or the channels.

A1R Activation Recruits GRK2-GFP to the MembraneSimultaneously with GIRK Current Desensitization,but Not mGluR2GRK2 is mainly cytosolic and translocates to the membrane to

phosphorylate active receptors (Pitcher et al., 1998). We wanted

to detect these translocations and to test whether there is a

correlation between the acceleration of GIRK desensitization

rates and GRK translocations. For this purpose, we C-terminally

tagged GRK2 with EGFP (GRK2-GFP) and used total internal

reflection fluorescence (TIRF) microscopy to detect exclusively

the membrane-associated fluorescence (Riven et al., 2003).

Cells transfected with GRK2-GFP and A1R showed a significant

GRK2-GFP basal membrane associated fluorescence (Fig-

ure 2A), as previously reported (Garcia-Higuera et al., 1994).

On A1R activation (Figures 2B and 2C) the membrane-associ-

ated fluorescent signal increased by 22.2 ± 6.2% with a t of

1.5 ± 0.4 s (Figures 2D and 2F). mOR also increased membrane

associated fluorescence on activation by 10.8 ± 2.8% with a t

of 23.4 ± 3.9 s (n = 11), temporally correlated with GCD for this

receptor (Figure S1D). Similar to the inability of mGluR2 to

accelerate GCD, membrane associated fluorescence also did

not significantly increase after mGluR2 activation (Figure 2D).

Similarly, M4R activation by carbachol did not induce GRK2

translocation to the membrane (data not shown). The transloca-

tions of GRK2-GFP to the membrane were reversible, as

membrane fluorescence returned to its basal level after washing

out the agonist (Figure S2). These results may indicate a strong

correlation between GRK2 translocation to the plasma

membrane and the acceleration in GCD rates. To further

strengthen this idea, we recorded A1R induced GIRK currents

and measured GRK-GFP translocation simultaneously, using

whole cell recording of the patch clamp technique, and quantita-

tive fluorescence under TIRF, respectively (Figure 2E). In cells

measured this way, GIRK desensitization and GRK2 recruit-

ments to the membrane occurred simultaneously, with change

of currents and membrane-associated fluorescence displaying t

of 2.4 ± 0.5 s and 4.6 ± 0.9 s, n = 5, respectively. Additional

independent observations of GCD rates and membrane-associ-

ated fluorescence increase of GRK2-GFP were also temporally

correlated with t of 1.3 ± 0.3 s, n = 20 and 1.5 ± 0.4 s, n = 11,

respectively (Figure 2F).

GPCR Phosphorylation and Receptor DownregulationAre Not Required for GRK2-Mediated GIRK CurrentDesensitizationIn the traditional view, after translocation to the membrane,

GRKs are responsible for the phosphorylation of activated

GPCRs. This event initiates the process of receptor downregula-

tion by clathrin-mediated endocytosis (Tsao and von Zastrow,

2000). To examine the relationship between this process and

the apparent GRK2-mediated acceleration in GCD as shown

above, we tested the ability of GRK2/K220R (dnGRK2), a domi-

nant negative mutant that lacks kinase catalytic activity (Kong

et al., 1994), in accelerating GCD rates (Figure 3A). The GCD

rates of cell cotransfected with GIRK, A1R, and dnGRK2 (5.5 ±

1.1 s, n = 9) were not different from cells expressing GRK2, the

receptor and channel components, with t of 2.6 ± 0.0 s (n = 9),

and significantly faster than in cells that were not cotransfected

with the kinase (24.9 ± 11.1 s; n = 8). These results suggest that

the enhancement of GCD rates is not mediated via the kinase

activity of GRK2.

Another possible mechanism for enhancing GCD might be

a change in receptor number, independent of GRK2-mediated

phosphorylation, or channel number, at the plasma membrane.

To test for these two possibilities, we C-terminally tagged the

A1R with GFP (A1R-GFP) or C-terminally tagged GIRK4 with

GFP (GIRK4-GFP) and measured plasma membrane-associated

fluorescence under TIRF. A1R-GFP and GIRK4-GFP plasma

membrane levels remained constant in the first minute after

agonist application both in control cells and in cells cotrans-

fected with GRK2, with DF/F of 96.3 ± 1.0%; n = 6 and 97.7 ±

0.3%; n = 12, for A1R-GFP and 96.4 ± 0.6%; n = 5 and

106.5 ± 1.4%; n = 9, for GIRK4-GFP, respectively (Figure 3B).

These results suggest that GRK2-mediated acceleration of the

GCD is neither due to a loss of receptors nor due to a loss of

GIRK channels from the plasma membrane.

Pertussis Toxin-Insensitive Pathways Are Sufficientto Induce GRK2 Translocations and Accelerationof GIRK Current DesensitizationThe sensitivity of A1R and mOR to GRK2-mediated desensitiza-

tion was distinct in comparison to mGluR2 and M4R, GPCRs that

display pure Gi/o activation. However, whereas A1R and mOR

primarily activate the Gi/o pathway, they may have also a

secondary transduction mechanism through different G protein

subsets (Cordeaux et al., 2004). We therefore tested whether

other minor secondary G protein activation mechanisms might

explain the selectivity of only a subset of receptors to induce

GRK2-mediated GCD. To inactivate the Gai/o pathway, we

coexpressed the catalytic subunit of pertussis toxin, PTX-S1,

that been shown to effectively abolish GPCR-mediated GIRK

activation (Sadja and Reuveny, 2009). In cells cotransfected

with PTX-S1, A1R, and GIRK channels, A1R activation did not

induce GIRK currents, in agreement with Gai/o sensitivity to

PTX (Figure 3C, middle). In contrast, when cells cotransfected

with both GRK2 and PTX-S1 were activated, the basal activity

of the GIRK channels, assessed by barium sensitivity of the

inward K+ currents at �80 mV, was rapidly reduced, in agree-

ment with the observation that a major part of GIRK basal activity

is Gbg-dependent (Rishal et al., 2005). Along the same line,


GRK2 translocation to the plasma membrane remained intact,

demonstrating that GRK2 membrane recruitment is not depen-

dent on the Gai/o pathway (Figures 3D and 3E). DF/F values

without or with PTX were 7.8 ± 0.6%, n = 13 and 8.0 ± 1.0%,

n = 7, respectively. As shown above, PTX-insensitive pathways

were sufficient to induce GRK2 translocations. The involvement

of other G protein signaling pathways, Gaq and Gas, were also

tested and were found not to be involved in GRK2 action on

GCD (Figures S3).

The Effects of Mutations in GRK2 that Impair ItsInteraction with Various Auxiliary MoleculesGRK2 is known to form a quaternary complex with Gaq and Gbg

(Tesmer et al., 2005). We set out to test whether impairing its

ability to interact with these auxiliary proteins may affect the

ability of GRK2 to accelerate GCD rates. GRK2 mutations that

disrupt GRK2-Gaq interaction, GRK2/R106A;D110A (Day et al.,

2004; Sterne-Marr et al., 2003) were tested. These mutations

are located in the RGS homology domain that is known to bind

Gaq but not Gai/o (Carman et al., 1999). GRK2/R106A;D110A

also accelerated GCD, similar to wt GRK2 (Figure S4A), with t

of 1.3 ± 0.4 s, n = 6 and 1.3 ± 0.3 s, n = 20, respectively.

GRK2D97-140, a GRK2 mutant that lacks the two helices that

are involved in GRK2-Gaq interaction, was also able enhance

GCD with t of 3.2 ± 0.8 s; n = 8. These results indicate that

A B

C

E

D

Figure 3. Kinase Catalytic Activity Is Not Required for GRK2 Effect

on GCD

A1R-GFP or GIRK1/GIRK4-GFP plasma membrane levels are not affected by

A1R stimulation. PTX treatment is not affecting basal GCD and membrane

recruitment.

(A) A bar graph summarizing measurements of GCD rates (t, s) from cells

cotransfected with GRK2/K220R (dnGRK2), GIRK, and A1R.

(B) The relative change of membrane fluorescence under TIRF (DF/F, %) asso-

ciated with either A1R-GFP or GIRK1/GIRK4-GFP before and during A1R

activation (1 min after adenosine application).

(C) Typical current traces of cells expressing GIRK and A1R (control); GIRK,

A1R and PTX (+PTX); and GIRK, A1R, PTX and GRK2 (+PTX +GRK2).

(D) A bar plot summarizing DF/F of GRK2-GFP signal after A1R activation,

measured under TIRF in cells expressing GIRK, PTX, and GRK2.

(E) A typical TIRF data of the membrane fluorescence change of GRK2-GFP

overtime of a cell expressing PTX after A1R activation.

See also Figure S3.

60

100

140

180

0 20 40 6

Fluo

resc

ence

Time (s)

0.0

1.0

2.0

Current desensitization

GRK2 translocation

t 0.6

7 (s

)

t=0 t=60s +AdeA

C

B

0

10

20

30

A1R mGluR2

dF/F

(%)

D

1s

Ade

20pA

FluorescenceCurrent

(s)

F

Ade

E

dF/F

(%)

Figure 2. A1R Activation Recruits GFP-Tagged GRK2 to the

Membrane Simultaneously with GCD as Revealed under TIRF

(A) A TIRF image of HEK293 cell transfected with GRK2-GFP. Basal membra-

nous fluorescence can be detected before stimulation by A1R.

(B) Image of the same cell in the presence of adenosine.

(C) Time course of fluorescence increase seen on receptor activation.

(D) A bar plot comparing the relative membrane-associated fluorescent

change (DF/F, in %) of GRK2-GFP after activation of A1R or mGluR2.

(E) Typical trace of whole-cell GIRK currents (black) and TIRF signal (green)

recorded simultaneously from the same cell.

(F) Bar graph depicting the similarity between GRK2-GFP translocation and

GCD rates.

See also Figure S2.


GRK2 interaction with Gaq is not required for GRK2 action on

GIRK currents.

The interactions between GRK2 and Gbg or phosphatidylino-

sitol 4,5-bisphosphate (PtdIns(4,5)P2) have also been thoroughly

studied in vitro, with different point mutations in GRK2

PH-domain (Carman et al., 2000; Sterne-Marr et al., 2003;

Touhara et al., 1995). Because both Gbg and PtdIns(4,5)P2 are

key players in the activation of GIRK channels (Huang et al.,

1998; Logothetis et al., 1987; Reuveny et al., 1994; Sui et al.,

1998), the GRK2-mediated enhancement of GCD might involve

interference of the interactions with these two molecules. We

thus compared GCD rates of control and GRK2 transfected cells,

and compared them with cells coexpressing the various GRK2

mutants (Figure 4A): GRK2/R587Q (Carman et al., 2000) and

GRK2/K663E;K665E;K667E (Touhara et al., 1995), that disrupt

the interactions of the kinase with Gbg, and GRK2/

K567E;R578E mutant that disrupts GRK2-PtdIns(4,5)P2 interac-

tions. Disrupting GRK2 interactions with Gbg abolished the

GRK2-mediated enhancement of GCD with t of 15.6 ± 1.9 s,

n = 32 and 12.6 ± 1.8 s, n = 15 for the GRK2/R587Q and

GRK2/K663E;K665E;K667E, respectively (Figure 4B). These

rates are comparable with cells that do not coexpress GRK2,

(t of 19.3 ± 2.1 s, n = 37). Furthermore, mutations that interrupt

GRK2 interactions with PtdIns(4,5)P2, GRK2/K567E;R578E

partially reduced the enhancement of GCD with t of 5.8 ±

0.6 s, n = 13. When the ability of membrane translocation after

receptor activation was tested for both PtdIns(4,5)P2 and Gbg

interaction mutants, using GRK2/K567E;R578E-GFP, GRK2/

K663E;K665E;K667E-GFP or GRK2/R587Q-GFP, respectively,

translocations to the membrane could be seen, but were

reduced in comparison to the wt GRK2 (Figure S4B). On the

contrary, a triple mutant GRK2/K567E;R578E;R587Q-GFP, in

which mutations that disrupt both Gbg and PtdIns(4,5)P2 binding

were introduced, no translocations were observed (Figure S4B).

These results are in agreement with the observations of coordi-

nated interactions of GRK2 with Gbg and PtdIns(4,5)P2 in

mediating GRK2 membrane recruitment (Pitcher et al., 1995).

To address whether the inability of GRK2/R587Q to accelerate

GCD is due to its reduced membrane translocation, we tethered

wild-type GRK2-GFP and GRK2/R587Q-GFP to the membrane

by fusing them with Src-myristoylation signal (myrGRK2-GFP

and myrGRK2/R587Q-GFP, respectively) (Figure 4C). GCD rates

were 1.3 ± 0.5 s (n = 8) and 23.9 ± 5.4 s (n = 5), for myrGRK2-GFP

and myrGRK2/R587Q-GFP, respectively (p < 0.05). Moreover,

five cells expressing myrGRK2/R587Q-GFP did not display

GCD at all. This supports the idea that failure of myrGRK2/

R587Q to accelerate GCD is due to its inability to chelate Gbg,

and not due to its impaired membrane targeting.

GRK2 Does Not Cause Desensitizationof Constituently Active GIRK MutantsBecause Gbg-GRK2 interactions seem to play an important role

in mediating the enhancement of GCD, one possible scenario is

that GRK2 is competing with the GIRK channel for Gbg on A1R-

activated release. To test this possibility, we examined the effect

of GRK on constituently active, Gbg independent GIRK mutant

channels (Sadja et al., 2001), GIRK1/S170P;GIRK4/S176P (Fig-

ure 5A). To avoid saturation and to ensure high quality voltage

clamp, we recorded currents in 5.6 mM external K+ solution.

Whole cell recordings of GIRK1/S170P;GIRK4/S176P show

high basal activity regardless of receptor activation (Figure S5)

(Sadja et al., 2001), with only a minor current induction on aden-

osine application. In contrast to wt GIRK recordings, GRK2 failed

to accelerate the GCD rates of the mutant channels (Figure 5B).

Currents flowing through GIRK1/S170P;GIRK4/S176P channel

mutants without or with GRK2 cotransfection showed current

levels of 95 ± 2%, n = 8 and 81 ± 4%, n = 10 (at 5 s of agonist

application), respectively. This is in contrast to the significant

GCD observed for the wild-type channel that had a reduction

of the residual current from 94 ± 14%, n = 7 to only 21 ± 4%,

n = 10 with GRK2 cotransfection at the same time point

(Figure 5B). These findings further point toward the possibility

that GRK2-mediated GCD involves the competition between

the channel and GRK2 for Gbg subunit.

B

A

C

Figure 4. GRK2 Mutants with Impaired GbgBinding Capability Fail to

Accelerate GIRK Desensitization

(A) A cartoon that displays the structure of the complex of GRK2 with Gbg

(Tesmer et al., 2005). The locations of the different point-mutations that were

used in (B) are marked in red.

(B) A bar plot summarizing the desensitization rates (t, s) of GIRK currents,

measured from cells transfected with GIRK channel, A1R, and the various

GRK2 mutants.

(C) A bar plot comparing the effect of myristoylated GRK2 (myr-GRK2) and

myrGRK2/R587Q mutant on GIRK desensitization rate.

See also Figure S4.


In light of the results described above, we were interested to

test whether PtdIns(4,5)P2 depletion from the channel may also

account for GRK2-mediated GCD. Therefore, we took an

advantage of the previously described GIRK mutants that

display enhanced PtdIns(4,5)P2 affinity, GIRK1/M223L;GIRK4/

I229L (Koike-Tani et al., 2005; Zhang et al., 1999) (Figure 5C).

Increasing GIRK channel affinity to PtdIns(4,5)P2 did not inhibit

the action of GRK2 on GCD rates, where GIRK1/M223L;

GIRK4/I229L without or with GRK2 showed (at 5 s during agonist

application) residual currents of 75 ± 4%, n = 13 and 31 ± 7%,

n = 16, respectively. Wild-type GIRK without or with GRK2

showed residual currents of 88 ± 5%, n = 7 and 14 ± 4%, n = 10,

respectively (Figure 5D). These results demonstrate that

GRK2-mediated acceleration of GCD does not occur by PtdIns

(4,5)P2 depletion from the channel.

A1R Activation Increases the Fraction of GRK2-BoundGbg PopulationAs shown above, mutations that impair GRK2-Gbg interaction

abolish the ability of GRK2 to accelerate GCD. To obtain further

evidence that indeed GRK2 binds Gbg in the context of the

plasma membrane, we recorded dynamic FRET using fluores-

cence lifetime approach (FRET-FLIM), under TIRF microscopy.

In this method donor fluorescence lifetime is recorded continu-

ously and shortening in donor lifetime is indicative of FRET. For

this purpose we used YFP and mCherry as donor and acceptor,

respectively. This pair has the advantage of a significant overlap

between donor emission and acceptor absorption, yet leaving an

acceptor-free donor fluorescence bandwidth for detection,

resulting in high FRET efficiencies (Goedhart et al., 2007) (Fig-

ure S6A). YFP has a nearly monoexponential lifetime decay

(Figures S6A and S6B) (Kremers et al., 2006), making it suitable

for use as a donor for FLIM measurements. Although cytosolic

+GRK2

Ade

Ade

50pA10s

Ba++

Ba++CONTROL

200pA10s

Ade

Ade

Ba++

Ba++

0

20

40

60

80

100

WT GIRK1/M223L; GIRK4/I229L

% C

urre

nt (

@ 5

s)

CONTROL GRK2

+GRK2

CONTROL

0

20

40

60

80

100

120

WT GIRK1/S170P; GIRK4/S176P

% C

urre

nt (@

5 s

)

CONTROL GRK2

A C

B D

Figure 5. Constituently Active, Gbg-Inde-

pendent, but Not GIRK Mutants that Have

Higher Affinity to PtdIns(4,5)P2, Are Insensi-

tive to GRK2

(A) Typical traces of GIRK1/S170P;GIRK4/S176P

channel mutants, without (upper trace) or in the

presence of GRK2 (lower trace).

(B) A bar plot summarizing the residual current

(in % of total current) after agonist application

without (dark gray) and in the presence of GRK2

(light gray).

(C) Typical current traces of GIRK1/M223L;GIRK4/

I229L channel mutants, without (upper trace) or in

the presence of GRK2 (lower trace).

(D) A bar plot summarizing the residual current (in

% of induced current) after agonist application

from cell without (dark gray) and with GRK2

(light gray). In both case, desensitization was

measure 5 s after agonist application.

See also Figure S5.

YFP showed a t of 2.6 ± 0.0 ns, n = 10,

in a fused dimer of YFP and mCherry

a subpopulation (92.9 ± 0.5%) of the

donor molecules displayed a much

shorter lifetime (0.6 ± 0.0 ns) corresponding to a FRET efficiency

of 76.7 ± 0.2%, n = 10 (Figure S6A). We set out to measure the

changes in FRET between N-terminally fused Gb1 with YFP

(YFP-Gb1) (Riven et al., 2006) and C-terminally fused GRK2

with mCherry (GRK2-Cherry) (Figure 6A). On A1R activation

YFP-Gb1 fluorescence decreased in the presence of GRK2-

Cherry, in agreement with YFP fluorescence quenching by

mCherry due to FRET (Figure 6B). Fitting the fluorescence life-

time decays of the donor over time revealed that, at rest, two

donor subpopulations exist (Figure 6C). One subpopulation

(22.6 ± 0.9%, n = 8) contains YFP-Gb1 proteins that interact

with GRK2-Cherry and hence result in shorter fluorescent life-

times of 0.6 ± 0.1 ns, n = 8. The remaining fraction consists of

free YFP-Gb1 proteins that display the characteristic monoexpo-

nential lifetime of YFP-Gb1 monomers (t-3.04 ns; see Fig-

ure S6B). After A1R activation, the relative fraction of YFP-Gb1

subunits that interact with GRK2-Cherry increases, seen as an

increase in the relative fraction of the shorter lifetime constants

(to 29.4 ± 1.6%, n = 8, p < 0.05) and as a decrease in the fraction

displaying long lifetime of the YFP (Figure 6C, D). The time

course of the shift in relative fraction of short and long lifetimes

(4.2 ± 0.7 s) resembles GCD rates and GRK2 translocations.

Similar correlation was seen when mOR was used, the rates of

YFP-Gb1 association with GRK2-Cherry was similar to the

GCD and to the GRK2-GFP translocation rates, with t average

for binding increase of 69.5 ± 15.3 s (n = 6) (Figure S6C). These

findings support the above observations that GRK2 action on

GCD is mediated through the binding of Gbg to GRK2.

DISCUSSION

Desensitization is an important cellular mechanism that allows

cells to adapt to long-term external stimuli. In the case of


GPCR signaling pathways, desensitization is mediated by a

decrease in the cellular response to a continuous GPCR stimula-

tion by agonists, resulting in a decrease in receptor number at

the plasma membrane. This process, that takes minutes to

hours, is mediated by phosphorylation of the receptor by GRK,

leading to clathrin-mediated endocytosis, in a process termed

downregulation (Bunemann et al., 1999; Tsao and von Zastrow,

2000). In addition to this well characterized process, other

mechanisms are necessary for a more rapid control of GPCR-

mediated signaling, specifically when the signal is intended to

control changes in electrical responsiveness of cells. In this

study we have described a mechanism that is responsible for

the termination of GPCR-mediated activation of GIRK channels,

which occurs within seconds.

In locus ceruleus neurons, Blanchet and Luscher (2002)

showed that prolonged activation of the mOR leads to inhibition

of GIRK function. It was shown that whereas mOR-mediated

presynaptic inhibition remained constant over time, postsyn-

aptic inhibition, mediated by GIRK activation, showed strong

desensitization of the response, indicating control over the

GIRK currents downstream of the receptors. This decrease in

GIRK currents could be overcome by additional activation of

G protein pathways. As a possible model for their results, it

was suggested that the receptor might activate Gbg scaven-

gers such as GRK2 and GRK3, to induce competitive inhibition

on GIRK activation. In a separate study using the same

neurons, it was shown that GCD was dependent on two molec-

ular pathways, the b-arrestin/GRK2 and the ERK1/2 pathways

(Dang et al., 2009). These findings suggested that GCD might

involve modifications of the G protein pathway that serves to

translate receptor activation to GIRK gating. In contrast, GCD

by muscarinic receptor stimulation has been attributed to

a mechanism solely involved the GPCR phosphorylation-

dependent and independent mechanisms by GRK2, and not

the G protein subunits (Shui et al., 1998). Here, using electro-

physiological and fluorescence resonance energy transfer

techniques, we unequivocally demonstrate that GRK2 is the

component of the G protein pathway that mediates this

short-term current decrease in the presence of the receptor

agonist. The molecular mechanism of this action will be

discussed below.

Based on our results, we suggest the following mechanism for

GRK2-mediated GCD (Figure 7): at rest, trimeric G-proteins are

bound to the nonactivated Gi/o-coupled GPCR and the channel

(Riven et al., 2006). After receptor activation by an agonist, the

Gbg subunits dissociate from the Ga subunit to interact with

the Gbg-binding domains on the channel, and promote channel

gating (opening). At the same time, GRK2 is recruited, either

within the two-dimensional space of the membrane (within

100 nm of the membrane space), or through the classical

cytosolic-to-plasma membrane translocation (Pitcher et al.,

1998). The former possibility may be aided by PtdIns(4,5)P2 or

by other membrane associated proteins, including the GIRK1

channel subunit (Dhami et al., 2004; Li et al., 2003; Palczewski,

1997; Rishal et al., 2005). This recruitment of GRK2, which is in

our case a receptor-specific event, promotes the binding of

the Gbg subunit to GRK2 or GRK3, but not GRK6 that lacks

Gbg binding capability, and thus reduces the availability of the

Gbg subunits to the channel. To have this chelation capacity,

GRK2 has to have a higher or comparable affinity for Gbg than

does the channel. Indeed, from binding studies it has been

shown that Gbg subunits bind recombinant GIRK1 or GIRK4

subunits with dissociation constants of �125 nM and �50 nM,

respectively (Krapivinsky et al., 1995), whereas Gbg affinity for

GRK2 is �20 nM (Pitcher et al., 1992; Wu et al., 1998). Further

evidence to support the idea that differential affinity to Gbg

may mediate this action comes from experiments where

GIRK4 was overexpressed in atrial myocytes (Bender et al.,

20

30

40

50

60

70

80

0 50 100 150 200 250 300

Rel

ativ

e fra

ctio

n (%

)

Time (s)

Tau donor

Tau fret

Ade

Ade

C

BA

-40-20

020406080

100120

0 20 40 60 80

Incr

ease

bin

ding

(Nor

mal

ized

, %)

Time (s)

DAde

600650700750800850900950

1000

0 50 100 150 200 250 300

Cou

nts

Time (s)

Figure 6. FLIM-FRET under TIRF Reveals

that A1R Activation Increases the Fraction

of GRK2-Bound Gbg

(A) A cartoon showing the experimental scheme

used in the FLIM-FRET experiments.

(B) Time course of YFP-Gb1 emission after the

activation of A1R in the presence of GRK2-Cherry,

in agreement with YFP quenching by mCherry on

increase of FRET.

(C) YFP-Gb1 lifetime changes after A1R activation.

Yellow symbol depicts the FRET-free YFP-Gb1

(t of 3.0 ns fraction) and red symbol depicts the

faster lifetime component (0.9 ns) that corre-

sponds to a FRET interaction between YFP-Gb1

and GRK2-Cherry. This FRET interaction corre-

sponds to a FRET efficiency of 0.7.

(D) YFP-Gb1 GRK2-Cherry binding increases

after A1R activation (n = 9). Black line depicts

the fitting to a monoexponential function with a

t = 4.2 ± 0.7 s.

See also Figure S6.


2001). In these experiments, GCD rates were greatly reduced, in

comparison to the GCD of GIRK1/4 heterotetramer, supporting

the idea that high affinity binding of Gbg may determine the

extent of channel current desensitization. Removal of Gbg from

the channel by GRK to affect channel function may not require

the removal of all four Gbg subunits, due to the steep depen-

dence of channel function on Gbg binding (Sadja et al., 2002).

Removing only one Gbg dimer reduces the efficacy of gating

by �70%. Finally, by using other means to chelate Gbg on the

membrane, such as coexpression of phosducin, similar effects

on GCD can be achieved (Riven et al., 2006). In conclusion, the

evidence provided above strongly points toward the possibility

that the acceleration of GCD by GRK2 is due to competition

for Gbg dimers with the channel.

How may GRK2-mediated GCD be interpreted in light of

previous suggested mechanisms? Few other mechanisms

have been proposed in the past to explain GCD. It has been

proposed that GIRK desensitization in cardiac cells might result

from simultaneous activation of M2R and M3R of the Gi/o and

the Gq pathways by acetylcholine, respectively (Keselman

et al., 2007; Kobrinsky et al., 2000; Meyer et al., 2001). Whereas

the former leads to GIRK opening, the latter leads to GCD by

PLC-mediated PtdIns(4,5)P2 depletion. Evidently, GCD occurs

also in simpler cases, where cross-talk between different GPCRs

pathways are probably not involved, and can be independent of

PtdIns(4,5)P2 depletion as showed by the use of PLC inhibitors

or activators (Meyer et al., 2001; Sickmann and Alzheimer,

2003). This was also true for our observations using NCDC,

a PLC inhibitor that does not block GIRK channel function (Sick-

mann et al., 2008). Furthermore, as shown above, mutations that

affect the affinity of the channel to PtdIns(4,5)P2 (Koike-Tani

et al., 2005; Zhang et al., 1999), are not affecting GRK2-mediated

channel desensitization. We thus suggest that changes in PtdIns

(4,5)P2 may only be an additional form of a much slower regula-

tion of channel function, mediated by the enzymatic activity of

PLC (Kobrinsky et al., 2000).

Our observations show that among four different receptors

described in this study, GCD was tightly regulated by GRK2 in

currents induced by A1R and mOR, showing a very robust

acceleration of GCD. On the contrary two other receptors,

namely mGluR2 and M4R were not able to induce GCD in the

presence of GRK2. How might this receptor selectivity be

addressed? It is interesting to note that receptors that were not

able to support GRK2-mediated GCD, were also not able to

recruit GRK2 to the plasma membrane, even though they all

release Gbg on activation to gate GIRK channels. This may

suggest that different receptors have differential mechanisms

to recruit GRK2 to the plasma membrane. The process of

membrane recruitment of GRK proteins has been ascribed to

a Gbg subunit-dependent mechanism (Pitcher et al., 1998;

Pitcher et al., 1992). It is therefore not clear how only a subset

of receptors have the ability to recruit the kinase, where others,

that also release Gbg to activate the GIRK channels, do not.

We have tried to address this issue and found that neither PLC

inhibition by NCDC, treatment with pertussis toxin, or using

dominant negative Gas mutant (Berlot, 2002) affected the ability

of the receptor to recruit GRK2 to the membrane (see Figure S3).

This may suggest of other still unknown mechanisms that

mediate this process by selective type of GPCRs, probably by

a specific direct interaction of the intracellular loops of the

receptor with GRK2.

How might the immediate desensitization be achieved? In

addition to cytosolic GRK that is recruited to the membrane on

receptor activation, a basal membranous subpopulation of

GRK2 is observed by us and by others (Aragay et al., 1998;

Garcia-Higuera et al., 1994; Murga et al., 1998). This subpopula-

tion can enable the immediate negative feedback of GIRK

activation. We cannot rule out also the possibility that GRK is

precoupled to GIRK (Rishal et al., 2005) and undergoes an

orientation/conformation change on activation, enabling its

immediate competition with the channels for Gbg subunits.

There are many studies suggesting the existence of signaling

complex between GIRK and Gbg (Clancy et al., 2005; Doupnik,

2008; Nikolov and Ivanova-Nikolova, 2004; Riven et al., 2006).

The GIRK-Gbg precoupling, before GPCR activation, might

enable the specificity of GPCR signaling cascade in an environ-

ment that may be populated by receptors of different types. Gbg

precoupled to GIRK undergo local rearrangement on GPCR

activation to immediately transduce GIRK gating independent

of diffusion rates (Riven et al., 2006). So if indeed the effector

(GIRK) is a module precoupled to its ‘‘switch-on,’’ could it be

that it is also precoupled to its ‘‘switch-off’’? There is evidence

that GRK2 and GIRK channel encompass a common signaling

complex (Nikolov and Ivanova-Nikolova, 2004).

Figure 7. A Cartoon Describing the Mechanism by

Which GRK2 Is Negatively Regulating GIRK Channel

Function

On receptor stimulation by GPCR, the G protein trimer

undergoes activation characterized by the exchange of GDP

for GTP on the Ga subunit. This in turn leads to the dissociation

of the Gbg subunits to freely bind and activate the GIRK

channel. Concomitantly, the GPCR induces the recruitment

of GRK2 to the plasma membrane making it available to bind

Gbg subunits of the G protein. Due to the relative higher affinity

of GRK2 for Gbg and to the larger mass action, GRK2 is now

able to effectively compete for the available pool of Gbg with

the GIRK channel, leading to a gradual removal of the Gbg

subunits and to a channel closure (desensitization), still in the

presence of the receptor agonist. Channel activation precedes

the action of GRK2 mainly due to the preexisting trimeric G

proteins in the vicinity of the channels (Riven et al., 2006).


Our results add a unique aspect to emerging evidence for

phosphorylation-independent activity of the GRK family, from

the regulation of receptor numbers or uncoupling of the GPCR

from the G protein at the plasma membrane, to regulation of

intracellular enzymes (for reviews see Ferguson [2007] and Reiter

and Lefkowitz [2006]). In all of these cases, there is no indication

of a direct involvement of the Gbg subunits of the G protein in

GRK action. GPCR/GRK2-dependent action on channel activity,

or other effectors, forms a new mechanism for a short-term

negative feedback for GPCR function, that selectively regulate

effector activity in the continued presence of receptor agonists.

This mechanism may not exclusively pertain to GIRK channels,

but can be relevant to all membrane associated Gbg regulated

effectors (Dupre et al., 2009). Because drug therapies for many

diseases are targeted to the receptor, a better understanding

of the pathway that links receptor to effector activation and

regulation (in this case the GIRK channel), and finding new

means to regulate these steps, might lead to therapies with

better resistance to complications such as tolerance and

side-effects.


Patch-Clamp Recordings

Membrane currents were recorded under voltage-clamp conditions using

whole-cell patch-clamp configuration with an Axopatch 200B (Axon Instru-

ments) patch-clamp amplifier. Patch pipettes were fabricated from borosili-

cate glass capillaries (2–5 MU). Signals were analog filtered using a 1 kHz

low-pass Bessel filter. After patch formation in a low K+ bath solution, the

bath solution was changed to high K+ solution. Adenosine (100 mM), glutamate

(100 mM), methionine enkephalin (ME, 100 nM), carbachol (100 mM), and Ba+2

(3 mM) were used to study induced and basal GIRK currents. GIRK currents

were measured as inward currents at a holding potential of �80 mV at room

temperature. Data acquisition and analysis were done using pCLAMP 9

software (Axon Instruments). To determine GCD kinetics, current traces

were fitted to a monoexponential decay function using Chebyshev method.

Results are expressed as average ± standard error of the mean (SEM). Signif-

icant differences were considered when p < 0.05 using Student’s t test.

TIRF Microscopy

Fluorescence was measured using through the objective TIRF microscopy

(Riven et al., 2003) with a 60 3 1.45 N.A. TIRFM objective (Olympus, Japan)

and TIRF condenser (TILL Photonics, Germany). Images were acquired with

Ixon+ EMCCD camera (Andor, Ireland) using Imaging Workbench 6 software

(Indec, USA). DF/F (%) was calculated from ROI that contained the whole

cell membrane area and was background subtracted. Time constant (t) for

GRK2 translocations, was calculated by determining the time after agonist

application when fluorescence reached 63% of maximum.

Fluorescence Lifetime Measurements

For fluorescence lifetime measurements (FLIM), 470 nm ps diode laser

(FWHM < 90 ps) was used, driven by a 40 MHz pulse controller, PDL 800-B.

Single photons were collected using PMA-165P photon counter and

processed using TimeHarp 200 PC-board. Data was acquired and analyzed

using SymPhoTime software (PicoQuant, Germany). Donor fluorescence

was collected from single cells under TIRF configuration (Riven et al., 2003).

For all measurements, laser intensities were set such that signal count rate

will be <1% of laser pulse rate. IRF was reconstructed from lifetime measure-

ment of YFP-Gb1 under TIRF using laser powers comparable to those used in

the experiment. YFP-Gb1 monomer lifetime was monoexponential with t of

3.0 ns (Figure S6B). To extract lifetimes and relative intensities, donor fluores-

cence traces were binned to 1-s segments and IRF reconvoluted trace

was fitted to double-exponential fitting model. One t parameter, td, was

constrained to 3.0 ns (YFP-Gb1), and tda as well as the relative size for

each exponential term was extracted from fitting result (Lleres et al., 2007;

Peter et al., 2005; Wallrabe and Periasamy, 2005; Yasuda et al., 2006).

Maximum likelihood estimation (MLE) method was used for fitting. Fit quality

was examined both by c2 values and by the absence of systematic variations

of fit residuals.

Molecular Biology and Cell Culture

Fusions to fluorescent proteins (EGFP, YFP and mCherry) were based on

commercially available pCMV-XFP vectors (Clontech). In EGFP A206K point

mutation was made to eliminate its week dimerization tendency (Zacharias

et al., 2002). Point mutations and deletion done in GIRK and GRK2 were

carried out by polymerase chain reaction (PCR) and verified by sequencing.

Nonfused GIRK and PTX-S1 subunits (Sadja and Reuveny, 2009) were all in

pcDNA3.1 (Invitrogen). C-terminal fusion of fluorescent proteins to GRK2 did

not affect its function. HEK293 cells were transiently transfected using

Metafectene (Biontex, Germany) with cDNAs encoding for the channel

subunits, the receptor of choice and GRK (wt, GFP-fused or mutant). In

GRK2 silencing experiments GRK2 shRNA (0.1 mg) or nontarget control

(0.1 mg) was cotransfected with the channel and the receptor. Currents were

measured 24–48 hr posttransfection according to Raveh et al. (2008). The

HL-1 cells, a gift from Dr. William C. Claycomb, were maintained using the

recommended protocols (Claycomb et al., 1998). For electrophysiology

experiments, cells were transferred to uncoated 24-mm glass coverslips on

the day of the recording.


Supplemental Information includes Extended Experimental Procedures

and six figures and can be found with this article online at doi:10.1016/

j.cell.2010.10.018.

ACKNOWLEDGMENTS

The authors like to thank Ruth Meller and Elisha Shalgi for technical help,

and the Reuveny laboratory for helpful comments. We are grateful to

Drs. J.L. Benovic for GRK2 and GRK6, W.C. Claycomb for HL-1 cells,

D.E. Logothetis for PtdIns(4,5)P2 GIRK mutants, C. Barlot for Gas mutant,

S. Nakanishi for the mGluR2, Z. Vogel for mOR, and R. Tsien for mCherry

cDNAs. The work was supported in part by the Josef Cohn Center for

Biomembrane Research, The Israeli Science Foundation (ISF grant 207/09),

The Minerva Foundation, and the Human Frontier Science Program.

Received: April 12, 2010




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Sequence-Dependent Sortingof Recycling Proteins by Actin-StabilizedEndosomal MicrodomainsManojkumar A. Puthenveedu,1,* Benjamin Lauffer,2 Paul Temkin,2 Rachel Vistein,1 Peter Carlton,3 Kurt Thorn,4

Jack Taunton,5 Orion D. Weiner,4 Robert G. Parton,6 and Mark von Zastrow2,5

1Department of Biological Sciences, Carnegie Mellon University, Pittsburgh, PA, USA2Department of Psychiatry3Department of Physiology4Department of Biochemistry and Biophysics5Department of Cellular and Molecular Pharmacology

University of California at San Francisco, San Francisco, CA 94158, USA6The University of Queensland, Institute for Molecular Bioscience and Centre for Microscopy and Microanalysis, St. Lucia,

Queensland 4072, Australia 8


DOI 10.1016/j.cell.2010.10.003

SUMMARY

The functional consequences of signaling receptorendocytosis are determined by the endosomal sort-ing of receptors between degradation and recyclingpathways. How receptors recycle efficiently, ina sequence-dependent manner that is distinct frombulk membrane recycling, is not known. Here, inlive cells, we visualize the sorting of a prototypicalsequence-dependent recycling receptor, the beta-2adrenergic receptor, from bulk recycling proteinsand the degrading delta-opioid receptor. Our resultsreveal a remarkable diversity in recycling routes atthe level of individual endosomes, and indicate thatsequence-dependent recycling is an active processmediated by distinct endosomal subdomainsdistinct from those mediating bulk recycling. Weidentify a specialized subset of tubular microdo-mains on endosomes, stabilized by a highly localizedbut dynamic actin machinery, that mediate this sort-ing, and provide evidence that these actin-stabilizeddomains provide the physical basis for a two-stepkinetic and affinity-based model for protein sortinginto the sequence-dependent recycling pathway.

INTRODUCTION

Cells constantly internalize a large fraction of proteins from their

surface and the extracellular environment. The fates of these

internalized proteins in the endosome have a direct impact on

several critical functions of the cell, including its response to

environmental signals (Lefkowitz et al., 1998; Marchese et al.,

2008; Sorkin and von Zastrow, 2009).

Internalized proteins have three main fates in the endosome.

First, many membrane proteins, such as the transferrin receptor

(TfR), are sorted away from soluble proteins, largely by bulk

membrane flow back to the cell surface. This occurs via the

formation and fission of narrow tubules that have a high ratio

of membrane surface area (and therefore membrane proteins)

to volume (soluble contents) (Mayor et al., 1993). Several

proteins have been implicated in the formation of these tubules

(Shinozaki-Narikawa et al., 2006; Cullen, 2008; Traer et al.,

2007), which provide a geometric basis to bulk recycling and

explain how nutrient receptors can recycle leaving soluble

nutrients behind to be utilized in the lysosome (Dunn and

Maxfield, 1992; Mayor et al., 1993; Maxfield and McGraw,

2004). Second, many membrane proteins are transported to

the lysosome to be degraded. This involves a process called

involution, where proteins are packaged into vesicles that bud

off to the interior of the endosome and, in essence, converts

these proteins into being a part of the soluble contents (Piper

and Katzmann, 2007). Involution has also been studied exten-

sively, and the machinery responsible, termed ESCRT complex,

identified (Hurley, 2008; Saksena et al., 2007; Williams and Urbe,

2007). Third, several other membrane proteins, such as many

signaling receptors, escape the bulk recycling and degradation

pathways, and are instead recycled in a regulated manner (Ha-

nyaloglu and von Zastrow, 2008; Yudowski et al., 2009). This

requires a specific cis-acting sorting sequence present on the

receptor’s cytoplasmic surface (Cao et al., 1999; Hanyaloglu

and von Zastrow, 2008). How receptors use these sequences

to escape the involution pathway and recycle, though they are

excluded from the default recycling pathway (Maxfield and

McGraw, 2004; Hanyaloglu et al., 2005), is a fundamental cell

biological question that is still unanswered.

Although it is clear that different recycling cargo can travel

through discrete endosomal populations (Maxfield and McGraw,

2004), endosome-to-plasma membrane recycling from a single

endosome is generally thought to occur via a uniform population

of tubules. Contrary to this traditional view, we identify special-

ized endosomal tubular domains mediating sequence-depen-

dent recycling that are kinetically and biochemically distinct


from the domains that mediate bulk recycling. These domains

are stabilized by a local actin cytoskeleton that is required and

sufficient for receptor recycling. We propose that such special-

ized actin-stabilized domains provide the physical basis for over-

coming a kinetic barrier for receptor entry into endosomal

tubules and for affinity-based concentration of proteins in the

sequence-dependent recycling pathway.

RESULTS

Visualization of Receptor Sorting in the Endosomesof Living CellsThe beta 2-adrenergic receptor (B2AR) and the delta opioid

receptor (DOR) provide excellent models for physiologically rele-

vant proteins that are sorted from each other in the endosome.

Although they share endocytic pathways, B2AR is recycled effi-

ciently in a sequence-dependent manner while DOR is selec-

tively degraded in the lysosome (Cao et al., 1999; Whistler

et al., 2002). To study the endosomal sorting of these cargo

molecules, we started by testing whether tubulation was

involved in this process. Because such sorting has not been

observed in vivo, we first attempted to visualize the dynamics

of receptor sorting in live HEK293 cells expressing fluorescently

labeled B2AR or DOR receptors, using high-resolution confocal

microscopy. Both receptors were observed mostly on the cell

surface before isoproterenol or DADLE, their respective

agonists, were added. After agonist addition, both B2AR

(Figure 1A) and DOR (data not shown) were robustly internalized,

and appeared in endosomes within 5 min (Figure 1A and Movie

S1 available online). As a control, receptors did not internalize

in cells not treated with agonists, but imaged for the same period

of time (Figure S1A). The B2AR-containing endosomes colocal-

ized with the early endosome markers Rab5 (Figure S1B) and

EEA1 (data not shown), consistent with previous data.

Internalized B2AR (Figure 1B), but not DOR (Figure 1C), also

labeled tubules that extended from the main body of the

receptor. When receptor fluorescence was quantified across

multiple B2AR-containing tubules, we saw that receptors were

enriched in these tubules compared to the rest of the endosomal

limiting membrane (Figure 1D). The bulk recycling protein TfR, in

contrast, was not enriched in endosomal tubules (Figure 1D).

This suggests that sequence-dependent recycling receptors

are enriched by an active mechanism in these endosomal

tubules.

These endosomal tubules were preferentially enriched for

B2AR over DOR on the same endosome. In cells coexpressing

FLAG-tagged B2AR and GFP-tagged DOR, we observed endo-

somes that contained both receptors within 5 min after coapply-

ing isoproterenol and DADLE. Notably, these endosomes

extruded tubules that contained B2AR but not detectable DOR

Figure 1. B2AR Is Enriched in Endosomal Tubular Domains Devoid of DOR

(A) HEK293 cells stably expressing FLAG-B2AR, labeled with fluorescently-tagged anti-FLAG antibodies, were followed by live confocal imaging before (left) and

after 5 min (right) of isoproterenol treatment. Arrows show internal endosomes.

(B) Example endosomes showing tubular domains enriched in B2AR (arrowheads) with one enlarged in the inset.

(C) Examples of DOR endosomes. DOR is smoothly distributed on the endosomal membrane and is not detected in tubules.

(D) Average fluorescence of B2AR (red circles) and TfR (green diamonds) calculated across multiple tubules (n = 123 for B2AR, 100 for TfR). B2AR shows a 50%

enrichment over the endosomal membrane, while TfR is not enriched. Each point denotes an individual tubule, the bar denotes the mean, and the gray dotted line

denotes the fluorescence of the endosomal membrane.

(E) An endosome containing both internalized B2AR and DOR, showing a tubule containing B2AR but no detectable DOR (arrowheads).

(F) Trace of linear pixel values across the same endosome, normalized to the maximum, confirms that the tubule is enriched for B2AR but not DOR.

(G) Linear pixel values of endosomal tubules averaged across 11 endosomes show specific enrichment of B2AR in tubules.

Error bars are SEM. See also Figure S1 and Movie S1 and Movie S2.


(e.g., in Figure 1E and in Movie S2). Fluorescence traces across

the endosome and the tubule confirmed that DOR was not

detectable in these B2AR tubules, suggesting that B2AR was

specifically sorted into these tubular domains (e.g., in Figure 1F).

When linear pixel values from multiple sorting events were quan-

tified, B2AR was enriched�50% in the endosomal domains from

which tubules originate, compared to the endosomal membrane

outside these domains (Figure 1G). Thus, these experiments

resolve, for the first time, individual events that mediate sorting

of two signaling receptors in the endosomes of live cells.

B2AR-ContainingEndosomal TubulesDeliver Receptorsto the Cell SurfaceTo test whether these tubules mediated recycling of B2AR, we

visualized direct delivery of receptors from these tubules to the

cell surface. In endosomes containing internalized B2AR and

DOR, these tubular domains pinched off vesicles that contained

B2AR but not detectable levels of DOR (Figure 2A and Movie S3).

To reliably assess if these vesicles traveled to the surface and

fused with the plasma membrane, we combined our current

imaging with a method that we have used previously to visualize

individual vesicle fusion events mediating surface receptor

delivery (Yudowski et al., 2006). Briefly, we attached the pH-

sensitive GFP variant superecliptic pHluorin to the extracellular

domain of B2AR (SpH-B2AR) (Miesenbock et al., 1998). SpH-

B2AR is highly fluorescent when exposed to the neutral pH at

the cell surface, but is quenched in the acidic environments of

endosomes and intracellular vesicles. This allows the detection

of individual fusion events of vesicles containing B2AR at the

cell surface (Yudowski et al., 2009). In cells coexpressing SpH-

B2AR and B2AR labeled with a pH-insensitive fluorescent dye

(Alexa-555), vesicles derived from the endosomal tubules traf-

ficked to the cell surface and fused, as seen by a sudden

increase in SpH fluorescence followed by loss of fluorescence

due to diffusion (Figure 2B, and Movie S4). A fluorescence trace

from movie S4 confirmed the fusion and loss of B2AR fluores-

cence (Figure 2C). Also, Rab4 and Rab11, which function in

endosome-to-plasma membrane recycling (Zerial and McBride,

2001; Maxfield and McGraw, 2004), were localized to the

domains containing B2AR (Figure S1). Together, this indicates

that the B2AR-containing endosomal tubules mediate delivery

of B2AR to the cell surface.

B2AR-Containing Tubules Are Marked by a HighlyLocalized Actin CytoskeletonWe next examined whether the B2AR-containing microdomains

were biochemically distinct from the rest of the endosomal

membrane. We first focused on actin, as the actin cytoskeleton

Figure 2. Membranes Derived from Endosomal Tubules Deliver B2AR to the Cell Surface

(A) Frames from a representative time lapse series showing scission of a vesicle that contains B2AR but not detectable DOR, from an endosomal tubule.

(B) An image plane close to the plasma membrane in cells coexpressing SpH-B2AR and FLAG-B2AR (labeled with Alexa555), exposed to isoproterenol for 5 min,

and imaged by fast dual-color confocal microscopy. Arrows denote the FLAG-B2AR-containing membrane derived from the endosomal tubule that fuses.

(C) Fluorescence trace of the B2AR-containing membranes from the endosome in movie S4, showing the spike in SpH-B2AR fluorescence (fusion) followed by

rapid loss of fluorescence.

Scale bars represent 1mm. See also Figure S1 and Movie S3 and Movie S4.


is required for efficient recycling of B2AR but not of TfR (Cao

et al., 1999; Gage et al., 2005), and as it has been implicated in

endosome motility (Stamnes, 2002; Girao et al., 2008) and

vesicle scission at the cell surface (Yarar et al., 2005; Perrais

and Merrifield, 2005; Kaksonen et al., 2005). Strikingly, in cells

coexpressing B2AR and actin-GFP, actin was concentrated on

the endosome specifically on the tubular domains containing

B2AR (Figure 3A). Virtually every B2AR tubule observed showed

this specific actin concentration on the tubule (n = 350). As with

actin, coronin-GFP (Uetrecht and Bear, 2006), an F-actin binding

protein, also localized specifically to the B2AR-containing

tubules on endosomes (Figure 3B), confirming that this was

a polymerized actin cytoskeleton. Coronin was also observed

on the B2AR-containing vesicle that was generated by dynamic

scission of the B2AR tubule (Figure 3B and Movie S5). Fluores-

cence traces of the linear pixels across the tubule and the vesicle

Figure 3. B2AR Tubules Are Marked by a Highly Localized Actin Cytoskeleton

(A) Cells coexpressing fluorescently labeled B2AR and actin-GFP exposed to isoproterenol for 5 min. The boxed area is enlarged in the inset, with arrowheads

indicating specific concentration of actin on B2AR endosomal tubules.

(B) Time lapse series from an example endosome with B2AR and coronin-GFP. Coronin is detectable on the endosomal tubule (arrows) and on the vesicle (arrow-

heads) that buds off the endosome.

(C) A trace of linear pixel values across the same endosome, normalized to maximum fluorescence, shows coronin on the endosomal domain and the vesicle.

(D) Example structured illumination image of a B2AR endosome showing specific localization of coronin to a B2AR tubule (arrowheads).

(E) Electron micrograph of an HRP-positive endosome (arrow) showing actin filaments (labeled with 9 nm gold, arrowheads) along a tubule. The right panel shows

an enlarged view.

See also Movie S5 and Movie S6.


confirmed that coronin pinched off with the B2AR vesicle

(Figure 3C).

We also used two separate techniques to characterize actin

localization on these tubules beyond the �250 nm resolution

offered by conventional microscopy. First, we first imaged the

localization of coronin on endosomes containing B2AR tubules

using structured illumination microscopy (Gustafsson et al.,

2008), which resolves structures at �100 nm spatial resolution.

3D stacks obtained using this high-resolution technique

confirmed that coronin was specifically localized on the endoso-

mal tubule that contained B2AR (Figure 3D and Movie S6).

Second, we examined the morphology of actin on endosomal

tubules at the ultrastructural level by pre-embedding immunoe-

lectron microscopy. Actin was clearly labeled as filaments lying

along tubules extruded from endosomal structures (Figure 3E).

Actin Is Dynamically Turned over on the B2AR-Containing Endosomal TubulesWe then tested whether the actin filaments on these tubules

were a stable structure or were dynamically turned over. When

cells expressing actin-GFP were exposed to latrunculin, a drug

that prevents actin polymerization, endosomal actin fluores-

cence became indistinguishable from the ‘‘background’’ cyto-

plasmic fluorescence within 16–18 s after drug exposure (e.g.,

in Figure 4A). When quantified across multiple cells, endosomal

actin fluorescence showed an exponential loss after latrunculin

exposure, with a t1/2 of 3.5 s (99% Confidence Interval = 3.0 to

4.1 s) (Figure 4B), indicating that endosomal actin turned over

quite rapidly. As a control, stress fibers, which are composed

of relatively stable capped actin filaments, were turned over

more slowly in these same cells (e.g., in Figure S2A). Endosomal

actin was lost in >98% of cells within 30 s after latrunculin, in

contrast to stress fibers, which persisted for over 2 min in

>98% of cells (Figure S2B). Rapid turnover of endosomal actin

was also independently confirmed by fluorescence recovery

after photobleaching (FRAP) studies. When a single endosomal

actin spot was bleached, the fluorescence recovered rapidly

within 20 s (Figure 4C). As a control for more stable actin fila-

ments, stress fibers showed little recovery of fluorescence after

bleaching in this interval (Figure 4C). Exponential curve fits

yielding a t1/2 of 8.26 s (99% CI = 7.65 to 8.97 s), consistent

with rapid actin turnover (Figure 4D). In contrast, only part of

the fluorescence (�30%) was recovered in stress fibers in the

same cells by 20 s, with curve fits yielding a t1/2 of 50.35 s

(99% CI = 46.05 to 55.54 s). These results indicate that actin is

dynamically assembled on the B2AR recycling tubules.

Considering the rapid turnover of actin, we next explored the

machinery responsible for localizing actin at the tubule.

The Arp2/3 complex is a major nucleator of dynamic actin poly-

merization that has been implicated in polymerization-based en-

dosome motility (Stamnes, 2002; Girao et al., 2008; Pollard,

2007). Arp3, an integral part of the Arp2/3 complex useful for

visualizing this complex in intact cells (Merrifield et al., 2004),

was specifically concentrated at the base of the B2AR tubules

on the endosome (e.g., in Figure 4E and fluorescence trace in

Figure 4F, Movie S7). Every B2AR tubule observed had a corre-

sponding Arp3 spot at its base (n = 200). Surprisingly, however,

we did not see N-WASP and WAVE-2, canonical members of the

two main families of Arp2/3 activators (Millard et al., 2004), on the

endosome (Figure 4G). Similarly, we did not see endosomal

recruitment of activated Cdc42, as assessed by a previously

characterized GFP-fusion reporter consisting of the GTPase

binding domain of N-WASP (Benink and Bement, 2005)

(data not shown). All three proteins were readily detected at

lamellipodia and filopodia as expected, indicating that the

proteins were functional in these cells. While we cannot rule

out a weak or transitory interaction of these activators with

Arp2/3 at the endosome, the lack of enrichment prompted us

to test for alternate Arp2/3 activators. Cortactin, an Arp- and

actin- binding protein present on endosomes, has been

proposed to be such an activator (Kaksonen et al., 2000; Millard

et al., 2004; Daly, 2004). Cortactin-GFP was clearly concentrated

at the base of the B2AR tubule on the endosome (Figure 4G), in

a pattern identical to Arp2/3. When quantified (>200 endosomes

each), every B2AR tubule was marked by cortactin, while none of

the endosomes showed detectable N-WASP, WAVE-2, or

Cdc42. Similarly, the WASH protein complex, which has been

recently implicated in trafficking from the endosome (Derivery

et al., 2009; Gomez and Billadeau, 2009; Duleh and Welch,

2010), was also clearly localized to B2AR tubules (Figure 4G).

Together, these data suggest that an Arp2/3-, cortactin- and

WASH-based machinery mediates dynamic actin assembly on

the endosome.

B2AR-Containing Tubules Are a Specialized Subsetof Recycling Tubules on the EndosomeSince the traditional view is that the endosomal tubules that

mediate direct recycling to the plasma membrane are a uniform

population, we next tested whether these tubules were the same

as those that recycle bulk cargo. When B2AR recycling was visu-

alized along with bulk recycling of TfR, endosomes containing

both cargo typically extruded three to four tubules containing

TfR. Strikingly, however, only one of these contained detectable

amounts of B2AR (Example in Figure 5A, quantified in Figure 5B).

This was consistent with fast 3D confocal live cell imaging of

B2AR in endosomes, which showed that most endosomes

extruded only one B2AR containing tubule, with a small fraction

containing two. When quantified, only 24.4% of all TfR tubules

contained detectable B2AR (n = 358 tubules).

B2AR Tubules Are a Kinetically and BiochemicallyDistinct from Bulk Recycling TubulesWhen the lifetimes of tubules were quantified, the majority

(>80%) of B2AR tubules lasted more than 30 s. In contrast, the

majority of TfR tubules devoid of B2AR lasted less than 30 s

(Figures 5B and 5C, Movie S8). Each endosome extruded

several tubules containing TfR, only a subset (�30%) of which

were marked by actin, coronin, or cortactin (Figures 5D and

5E, arrows). Time-lapse movies indicated that the highly tran-

sient TfR-containing tubules were extruded from endosomal

domains that were lacking cortactin (Figure 5E, arrows), while

the relatively stable B2AR containing tubules were marked by

cortactin (Figure 5E, arrowheads). Importantly, the relative

stability of the subset of tubules was conferred by the actin cyto-

skeleton, as disruption of actin using latrunculin virtually abol-

ished the stable fraction of TfR tubules (Figures 5B and 5C).


Figure 4. Actin on B2AR Tubules Is Dynamic and Arp2/3-Nucleated

(A) Cells expressing actin-GFP imaged live after treatment with 10 mM latrunculin for the indicated times, show rapid loss of endosomal actin. A time series of the

boxed area, showing several endosomal actin loci, is shown at the lower panel.

(B) The change in endosomal and cytoplasmic actin fluorescence over time after latrunculin normalized to initial endosomal actin fluorescence (n = 10). One-

phase exponential curve fits (solid lines) show a t1/2 of 3.5 s for actin loss (R2 = 0.984, d.f = 23, Sy.x = 2.1 for endosomal actin, R2 = 0.960, d.f = 23, Sy.x =

1.9 for cytoplasmic). Endosomal and cytoplasmc actin fluorescence becomes statistically identical within 15 s after latrunculin. Error bars denote SEM.

(C) Time series showing FRAP of representative examples of endosomal actin (top) and stress fibers (bottom).

(D) Kinetics of FRAP of actin (mean ± s.e.m) quantified from 14 endosomes and 17 stress fibers. One-phase exponential curve fits (lines), show a t1/2 of 8.26 s for

endosomal actin (R2 = 0.973, d.f = 34, Sy.x = 4.8) and 50.35 s for stress fibers (R2 = 0.801, d.f = 34, Sy.x = 3.9).


Together, these results suggest that sequence-dependent

recycling of B2AR is mediated by specialized tubules that are

kinetically and biochemically distinct from the bulk recycling

tubules containing only TfR.

A Kinetic Model for Sorting of B2AR into a Subsetof Endosomal TubulesThe relative stability of B2AR tubules suggested a simple model,

based on kinetic sorting, for how sequence-dependent cargo

was sorted into a specific subset of tubules and excluded from

the transient TfR-containing bulk-recycling tubules. We hypoth-

esized that B2AR diffuses more slowly on the endosomal

membrane relative to bulk recycling cargo. The short lifetimes

of the bulk-recycling tubules would then create a kinetic barrier

for B2AR entry, while this barrier would be overcome in the

subset of tubules stabilized by actin.

To test the key prediction of this model, that B2AR diffuses

more slowly than TfR on the endosomal membrane, we directly

measured the diffusion rates of B2AR and TfR using FRAP. When

B2AR or TfR was bleached on a small part of the endosomal

membrane, B2AR fluorescence took significantly longer to

recover than TfR (Figure 5F). When quantified, the rate of

recovery of fluorescence of B2AR (t1/2 = 25.77 s, 99% CI 23.45

to 28.6 s) was �4 times slower than that of TfR (t1/2 = 6.21 s,

99% CI 5.49 to 7.17 s), indicating that B2AR diffuses significantly

slower on the endosomal membrane than TfR (Figures 5F and

5G). Neither B2AR or TfR recovered within the time analyzed

when the whole endosome was bleached (Figure 5H), confirming

that the recovery of fluorescence was due to diffusion from the

unbleached part of the endosome and not due to delivery of

new receptors via trafficking. Further, B2AR on the plasma

membrane diffused much faster than on the endosome (t1/2 =

6.45 s, 99% CI 5.62 to 7.66 s), comparable to TfR, suggesting

that B2AR diffusion was slower specifically on the endosome

(Figure 5H).

We next tested whether the diffusion of B2AR into endosomal

tubules was slower than that of TfR, by using the rate of increase

of B2AR fluorescence as an index of receptor entry into tubules.

B2AR fluorescence continuously increased throughout the dura-

tion of the tubule lifetimes (Figure S3A). Further, in a single tubule

containing TfR and B2AR, TfR fluorescence reached its

maximum at a markedly faster rate than that of B2AR (Fig-

ure S3B). Together, these results suggest that slow diffusion of

B2AR on the endosome and stabilization of recycling tubules

by actin can provide a kinetic basis for specific sorting of

sequence-dependent cargo into subsets of endosomal tubules.

Local Actin Assembly Is Required for B2AR Entryinto the Subset of TubulesBecause actin stabilizes the B2AR-containing subset of tubules,

the model predicts that endosomal actin would be required for

sequence-dependent concentration of B2AR into these tubules.

Consistent with this, B2AR was no longer concentrated in endo-

somal tubules when endosomal actin was acutely removed

using latrunculin (e.g., in Figure 6A). When the pixel fluorescence

along the limiting membrane of multiple endosomes was quanti-

fied, B2AR was distributed more uniformly along the endosomal

membrane in the absence of actin (Figures 6B and 6C). We

further confirmed this by comparing the variance in B2AR fluo-

rescence along the endosomal perimeter, irrespective of their

orientation. B2AR fluorescence was significantly more uniform

in endosomes without actin (Figure 6D), indicating that actin

was required for endosomes to concentrate B2AR in microdo-

mains. Less than 20% of endosomes showed B2AR-containing

tubules in the absence of endosomal actin, in contrast to control

cells where over 75% of endosomes showed B2AR-containing

tubules (Figure 6E). Further, cytochalasin D, a barbed-end

capping drug that prevents further actin polymerization but

does not actively cause depolymerization, also inhibited B2AR

entry into tubules (Figure 6E) and B2AR surface recycling

(Figure S4A). Neither TfR tubules on endosomes (Figure 6E)

nor TfR recycling (Figure S4B) was inhibited by actin depolymer-

ization, consistent with a role for actin specifically in sequence-

dependent recycling of B2AR (Cao et al., 1999). Further, deple-

tion of cortactin using siRNA (Figure 6F) also inhibited B2AR

entry into tubules (Figures 6G and 6H). This inhibition was

specific to cortactin depletion, as it was rescued by exogenous

expression of cortactin (Figure 6H). Together, these results indi-

cate that a localized actin cytoskeleton concentrates sequence-

dependent recycling cargo into a specific subset of recycling

tubules on the endosome.

B2AR Sorting into the Recycling SubdomainsIs Mediated by Its C-Terminal PDZ-Interacting DomainWe next asked whether this actin-dependent concentration of

receptors into endosomal tubules depended on the PDZ-inter-

acting sequence present in the B2AR cytoplasmic tail that medi-

ates sequence-dependent recycling (Cao et al., 1999; Gage

et al., 2005). To test if the sequence was required, we used

a mutant B2AR (B2AR-ala) in which the recycling sequence

was specifically disrupted by the addition of a single alanine

(Cao et al., 1999). Unlike B2AR, internalized B2AR-ala was not

able to enter the tubular domains in the endosome (e.g., in

Figure 6I, quantified in Figure 6J), or recycle to the cell surface

(Figure S4). To test if this sequence was sufficient, we used

a chimeric DOR construct with the B2AR-derived recycling

sequence fused to its cytoplasmic tail, termed DOR-B2 (Gage

et al., 2005), which recycles much more efficiently than DOR

(Figure S4). In contrast to DOR, which showed little concentra-

tion in endosomal tubules, DOR-B2 entered tubules (Figures 6I

and 6J) and recycled in an actin-dependent manner similar to

B2AR (Figure S4D). Together, these results indicate that the

(E) Example endosomes in live cells coexpressing B2AR and Arp3-GFP showing Arp3 at the base of B2AR tubules (arrowhead in the inset).

(F) Trace of linear pixel fluorescence of B2AR and Arp3 shows Arp3 specifically on the endosomal tubule.

(G) Example endosomes from cells coexpressing B2AR and N-WASP-, WAVE2-, cortactin-, or WASH-GFP. N-WASP and WAVE2 were not detected on endo-

somes, while cortactin and WASH were concentrated at the B2AR tubules (arrowheads).

Scale bars represent 1 mm. See also Figure S2 and Movie S7.


Figure 5. B2AR Is Enriched Specifically in a Subset of Endosomal Tubules that Are Stabilized by Actin

(A) A representative example of an endosome with two tubules containing TfR, only one of which is enriched for B2AR.

(B) The number of tubules with B2AR, TfR, and TfR in the presence of 10 mM latrunculin, per endosome per min, binned into lifetimes less than or more than 30 s,

quantified across 28 endosomes and 281 tubules.

(C) The percentages of B2AR, TfR, and TfR + latrunculin tubules with lifetimes less than or more than 30 s, normalized to total number of tubules in each case.

(D) An example endosome containing TfR and coronin, showing that coronin is present on a subset of the TfR tubules. Arrowheads indicate a TfR tubule that is

marked by coronin, and arrows show a TfR tubule that is not.

(E) Time lapse series showing TfR-containing tubules extruding from endosomal domains without detectable cortactin. Arrowheads indicate a relatively stable

TfR tubule that is marked by coronin, and arrows denote rapid transient TfR tubules without detectable cortactin.

(F) Frames from a representative time lapse movie showing FRAP of B2AR (top row) or TfR (bottom row). The circles mark the bleached area of the endosome. TfR

fluorescence recovers rapidly, while B2AR fluorescence recovers slowly.

(G) Fluorescence recovery of B2AR (red circles) and TfR (green diamonds) on endosomes quantified from 11 experiments. Exponential fits (solid lines) show that

B2AR fluorescence recovers with a t1/2 of 25.77 s (R2 = 0.83, d.f = 37, Sy.x = 6.3), while TfR fluorescence recovers with a t1/2 of 6.21 s (R2 = 0.91, d.f = 30, Sy.x = 7.1).


PDZ-interacting recycling sequence on B2AR was both required

and sufficient to mediate concentration of receptors in the actin-

stabilized endosomal tubular domains.

As PDZ-domain interactions have been established to indi-

rectly link various integral membrane proteins to cortical actin

(Fehon et al., 2010), we tested whether linking DOR to actin

was sufficient to drive receptor entry into endosomal tubules.

Remarkably, fusion of the actin-binding domain of the ERM

protein ezrin (Turunen et al., 1994) to the C terminus of DOR

was sufficient to localize the receptor (termed DOR-ABD) to

endosomal tubules (Figure 6J). The surface recycling of B2AR,

DOR-B2, and DOR-ABD were dependent on the presence of

an intact actin cytoskeleton (Figure S4), consistent with previous

publications (Cao et al., 1999; Gage et al., 2005; Lauffer et al.,

2009). Further, transplantation of the actin-binding domain was

also sufficient to specifically confer recycling to a version of

B2AR lacking its native recycling signal (Figure S4F). These

results indicate that the concentration of B2AR in the actin-stabi-

lized recycling tubules is mediated by linking receptors to the

local actin cytoskeleton through PDZ interactions.

DISCUSSION

Even though endocytic receptor sorting was first appreciated

over two decades ago (e.g., Brown et al., 1983; Farquhar,

1983; Steinman et al., 1983), our understanding of the principles

of this process has been limited. A major reason for this has been

the lack of direct assays to visualize signaling receptor sorting in

the endosome. Here we directly visualized, in living cells, endo-

somal sorting between two prototypic members of the largest

known family of signaling receptors for which sequence-specific

recycling is critical for physiological regulation of cell signaling

(Pippig et al., 1995; Lefkowitz et al., 1998; Xiang and Kobilka,

2003). We resolve sorting at the level of single trafficking events

on individual endosomes, and define a kinetic and affinity-based

model for how sequence-dependent receptors are sorted away

from bulk-recycling and degrading proteins.

By analyzing individual sorting and recycling events on single

endosomes, we demonstrate a remarkable diversity in recycling

pathways emanating from the same organelle (Scita and Di

Fiore, 2010). The traditional view has been that recycling to the

plasma membrane is mediated by a uniform set of endosomal

tubules from a single endosome. In contrast to this view, we

demonstrate that the recycling pathway is highly specialized,

and that specific cargo can segregate into specialized subsets

of tubules that are biochemically, biophysically, and functionally

distinct. Receptor recycling plays a critical role in controlling the

rate of cellular re-sensitization to signals (Lefkowitz et al., 1998;

Sorkin and von Zastrow, 2009), and recent data suggest that

the sequence-dependent recycling of signaling receptors is

selectively controlled by signaling pathways (Yudowski et al.,

2009). The physical separation between bulk and sequence-

dependent recycling that we demonstrate here allows for such

selective control without affecting the recycling of constitutively

cycling nutrient receptors. Further, such physical separation

might also reflect the differences in molecular requirements

that have been observed between bulk and sequence-depen-

dent recycling (Hanyaloglu and von Zastrow, 2007).

Endosome-associated actin likely plays a dual role in endoso-

mal sorting, both of which contribute to sequence-dependent

entry of cargo selectively into special domains. First, by stabi-

lizing the specialized endosomal tubules relative to the much

more dynamic tubules that mediate bulk recycling, the local actin

cytoskeleton could allow sequence-dependent cargo to

overcome a kinetic barrier that limits their entry into the bulk

pathway. Supporting this, we show that most endosomal tubules

are highly transient, lasting less than a few seconds (Figures 5B

and 5C), which allows enough time for entry of the fast-diffusing

bulk recycling cargo, but not the slow-diffusing sequence-

dependent cargo (Figures 5F and 5G), into these tubules.

A subset of these tubules representing the sequence-dependent

recycling pathway is stabilized by the presence of an actin cyto-

skeleton (Figures 5B and 5C). This stabilization allows time for

B2AR to diffuse into these tubules (Figure S3), which eventually

pinch off membranes that can directly fuse with the plasma

membrane (Figure 2). Interestingly, inhibition of actin caused

a decrease in the total number of tubules by approximately

25% (Figure 5B), suggesting that the actin cytoskeleton plays

a role in maintaining the B2AR-containing subset of tubules,

and not just in the sorting of B2AR into these tubules.

Second, a local actin cytoskeleton could provide the

machinery for active concentration of recycling proteins like

the B2AR, which interact with actin-associated sorting proteins

(ERM and ERM-binding proteins) through C-terminal sequences

(Weinman et al., 2006; Wheeler et al., 2007; Lauffer et al., 2009;

Fehon et al., 2010), in specialized recycling tubules. Consistent

with this, the C-terminal sequence on B2AR was both required

and sufficient for sorting to the endosome and for recycling,

and a distinct actin-binding sequence was sufficient for both

receptor entry into tubules and recycling (Figure 6 and Figure S4).

PDZ-interacting sequences have been identified on several

signaling receptors, including multiple GPCRs, with different

specificities for distinct PDZ-domain proteins (Weinman et al.,

2006). Further, actin-stabilized subsets of tubules were present

even in the absence of B2AR in the endosome. We propose

that, using a combination of kinetic and affinity-based sorting

principles, discrete Actin-Stabilized SEquence-dependent

Recycling Tubule (ASSERT) domains could thus mediate effi-

cient sorting of sequence-dependent recycling cargo away

from both degradation and bulk recycling pathways that diverge

from the same endosomes.

Our results, therefore, uncover an additional role for actin poly-

merization in endocytic sorting, separate from its role in endo-

some motility. It will be interesting to investigate the mechanism

and signals that control the nucleation of such a spatially local-

ized actin cytoskeleton on the endosome. The lack of obvious

(H) Fluorescence recovery of B2AR (blue triangles) and TfR (green diamonds) on endosomes when the whole endosome was bleached, or of B2AR on the cell

surface (red circles) quantified from 12 experiments. B2AR fluorescence on the surface recovers with a t1/2 of 6.49 s (R2 = 0.94, d.f = 27, Sy.x = 8.1).

Error bars denote SEM. Scale bars represent 1 mm. See also Figure S3 and Movie S8.


Figure 6. B2AR Enrichment in Tubules Depends on Endosomal Actin and a PDZ-Interacting Sequence on the B2AR Cytoplasmic Domain

(A) Representative fields from B2AR-expressing cells exposed to isoproterenol showing B2AR endosomes before (top panel) or after (bottom panel) exposure to

10 mM latrunculin for 5 min. Tubular endosomal domains enriched in B2AR (arrowheads) are lost upon exposure to latrunculin.

(B) Schematic of measurement of endosomal B2AR fluorescence profiles in the limiting membrane. The profile was measured in a clockwise manner starting from

the area diametrically opposite the tubule (an angle of 0�).(C) B2AR concentration along the endosomal membrane, calculated from fluorescence profiles of 20 endosomes, normalized to the average endosomal B2AR

fluorescence. In the presence of latrunculin, B2AR enrichment in tubules is abolished, and B2AR fluorescence shows little variation along the endosomal

membrane.

(D) Variance in endosomal B2AR fluorescence values measured before and after latrunculin. B2AR distribution becomes more uniform after latrunculin.

(E) The percentages of endosomes extruding B2AR-containing tubules, calculated before (n = 246) and after (n = 106) treatment with latrunculin, or before

(n = 141) and after (n = 168) cytochalasin-D, show a significant reduction after treatment with either drug. As a control, the percentages of endosomes extruding

TfR-containing tubules before (n = 317) and after (n = 286), respectively, are shown.

(F) Cortactin immunoblot showing reduction in protein levels after siRNA.

(G) Representative fields from B2AR-containing endosomes in cells treated with control and cortactin siRNA. Arrowheads denote endosomal tubules in the

control siRNA-treated cells.

(H) Percentages of endosomes extruding B2AR tubules calculated in control siRNA-treated cells (n = 210), cortactin siRNA-treated cells (n = 269), and cortactin

siRNA-treated cells expressing an siRNA-resistant cortactin (n = 250).

(I) Representative examples of endosomes from agonist-exposed cells expressing B2AR, B2AR-ala, DOR, or DOR-B2. Arrowheads denote receptor-containing

tubules on B2AR and DOR-B2 endosomes.

(J) The percentage of endosomes with tubular domains containing B2AR, B2AR-ala, DOR, DOR-B2, or DOR-ABD (n = 246, 302, 137, 200, and 245, respectively)

were quantified.

Scale bars represent 1 mm; and error bars represent SEM. See also Figure S4.


concentration of the canonical Arp2/3 activators, WASP and

WAVE, suggests a novel mode of actin nucleation involving cor-

tactin. Cortactin can act as a nucleation-promoting factor for

Arp2/3, at least in vitro (Ammer and Weed, 2008), and can

interact with dynamin (Schafer et al., 2002; McNiven et al.,

2000), which makes it an attractive candidate for coordinating

actin dynamics on membranes. Interestingly, inhibition of

WASH, a recently described Arp regulator that is present on

B2AR tubules, has been reported to result in an increase in endo-

somal tubules (Derivery et al., 2009). Although its role in

sequence-dependent recycling remains to be tested, this

suggests the presence of multiple actin-associated proteins

with distinct functions on the endosome.

The simple kinetic and affinity-based principle that we

propose likely provides a physical basis for sequence-depen-

dent sorting of internalized membrane proteins between essen-

tially opposite fates in distinct endosomal domains. Proteins that

bind sequence-dependent degrading receptors and are required

for their degradation (Whistler et al., 2002; Marley and von

Zastrow, 2010) might act as scaffolds and provide a similar

kinetic barrier to prevent them from accessing the rapid bulk-re-

cycling tubules. Entry of these receptors into the involution

pathway might then be accelerated by their association with

the well-characterized ESCRT-associated domains on the vacu-

olar portion of endosomes (Hurley, 2008; Saksena et al., 2007;

Williams and Urbe, 2007), complementary to the presently iden-

tified ASSERT domains on a subset of endosomal tubules.

Such diversity at the level of individual trafficking events to the

same destination from the same organelle raises the possibility

that there exists yet further specialization among the pathways

that mediate exit out of the endosome, including in the degrada-

tive pathway and the retromer-based pathway to the trans-Golgi

network. Importantly, the physical separation in pathways that

we report here potentially allows for cargo-mediated regulation

as a mode for controlling receptor recycling to the plasma

membrane. Such a mechanism can provide virtually an unlimited

level of selectivity in the post-endocytic system using minimal

core trafficking machineries, as has been observed for endocy-

tosis at the cell surface (Puthenveedu and von Zastrow, 2006).

As the principles of such sorting depend critically on kinetics,

the high-resolution imaging used here to analyze domain kinetics

and biochemistry, and to achieve single-event resolution in living

cells, provides a powerful method to elucidate biologically

important sorting processes in the future.


Constructs and Reagents

Receptor constructs and stably transfected HEK293 cell lines are described

previously (Gage et al., 2005; Lauffer et al., 2009) Transfections were per-

formed using Effectene (QIAGEN) according to manufacturer’s instructions.

For visualizing receptors, FLAG-tagged receptors were labeled with M1 anti-

bodies (Sigma) conjugated with Alexa-555 (Invitrogen) as described (Gage

et al., 2005), or fusion constructs were generated where receptors were

tagged on the N-terminus with GFP. Latrunculin and Cytochalasin D (Sigma)

were used at 10 mM final concentration.

Live-Cell and Fluorescence Imaging

Cells were imaged using a Nikon TE-2000E inverted microscope with a 1003

1.49 NA TIRF objective (Nikon) and a Yokagawa CSU22 confocal head (Sola-

mere), or an Andor Revolution XD Spinning disk system on a Nikon Ti micro-

scope. A 488 nm Ar laser and a 568 nm Ar/Kr laser (Melles Griot), or 488 nm

and 561 nm solid-state lasers (Coherent) were used as light sources. Cells

were imaged in Opti-MEM (GIBCO) with 2% serum and 30 mM HEPES

(pH 7.4), maintained at 37�C using a temperature-controlled incubation

chamber. Time lapse images were acquired with a Cascade II EM-CCD

camera (Photometrics) driven by MicroManager (www.micro-manager.org)

or an Andor iXon+ EM-CCD camera using iQ (Andor). The same lasers were

used as sources for bleaching in FRAP experiments. Structured illumination

microscopy was performed as described earlier (Gustafsson et al., 2008).

Electron Microscopy

EM studies were carried out using MDCK cells because they are amenable to

a previously described pre-embedding processing that facilitates detection of

cytoplasmic actin filaments (Ikonen et al., 1996; Parton et al., 1991), and

because they contain morphologically similar endosomes to HEK293 cells.

Cells were grown on polycarbonate filters (Transwell 3412; Costar, Cam-

bridge, MA) for 4 days as described previously (Parton et al., 1991). To allow

visualization of early endosomes and any associated filaments a pre-embed-

ding approach was employed. Cells were incubated with HRP (Sigma type II,

10mg/ml) in the apical and basolateral medium for 10min at 37�C and then

washed, perforated, and immunogold labeled with a rabbit anti- actin anti-

body, a gift of Professor Jan de Mey (Strasbourg), followed by 9nm protein

A-gold. HRP visualization and epon embedding was as described previously

(Parton et al., 1991; Ikonen et al., 1996).

Image and Data Analysis

Acquired image sequences were saved as 16-bit tiff stacks, and quantified

using ImageJ (http://rsb.info.nih.gov/ij/). For estimating receptor enrichment,

a circular mask 5 px in diameter was used to manually select the membrane

at the base of the tubule or membranes derived from endosomes. Fluores-

cence values measured were normalized to that of the endosomal membrane

devoid of tubules. An area of the coverslip lacking cells was used to estimate

background fluorescence. For estimating linear pixel values along the tubules,

a line selection was drawn along the tubule and across the endosome, and the

Plot Profile function used to measure pixel values. For obtaining the average

value plot across multiple sorting events, the linear pixels were first normalized

to the diameter of the endosome and then averaged. To generate pixel values

along the endosomal limiting membranes, the Oval Profile plugin, with 60

segments, was used after manually selecting the endosomal membrane using

an oval ROI. Lifetimes of tubules were calculated by manually tracking the

extension and retraction of tubules over time-lapse series. Microsoft Excel

was used for simple data analyses and graphing. Curve fits of data were per-

formed using GraphPad Prism. All P-values are from two-tailed Mann-Whitney

tests unless otherwise noted.


Supplemental Information includes four figures and eight movies and can be

found with this article online at doi:10.1016/j.cell.2010.10.003.

ACKNOWLEDGMENTS

The majority of the imaging was performed at the Nikon Imaging Center at

UCSF. We thank David Drubin, Matt Welch, John Sedat, Aylin Hanyaloglu,

Aaron Marley, and James Hislop for essential reagents and valuable help.

M.A.P. was supported by a K99/R00 grant DA024698, M.v.Z. by an R37 grant

DA010711, and O.D.W. by an RO1 grant GM084040, all from the NIH. J.T. is an

investigator of the Howard Hughes Medical Institute.

Received: October 31, 2009

Revised: April 7, 2010

Accepted: September 27, 2010



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Mechanisms Determining the Morphologyof the Peripheral ERYoko Shibata,1,2 Tom Shemesh,3 William A. Prinz,4 Alexander F. Palazzo,1,5 Michael M. Kozlov,3,*and Tom A. Rapoport1,2,*1Howard Hughes Medical Institute2Department of Cell Biology

Harvard Medical School, 240 Longwood Avenue, Boston, MA 02115, USA3Department of Physiology and Pharmacology, Sackler Faculty of Medicine, Tel Aviv University, Ramat Aviv, 69978 Tel Aviv, Israel4Laboratory of Cell Biochemistry and Biology, National Institute of Diabetes and Digestive and Kidney Disorders,

National Institute of Health, Bethesda, MD 02892, USA5Department of Biochemistry, University of Toronto, 1 King’s College Circle, Toronto, ON M5S 1A8, Canada*Correspondence: [email protected] (M.M.K.), [email protected] (T.A.R.)

DOI 10.1016/j.cell.2010.11.007

SUMMARY

The endoplasmic reticulum (ER) consists of thenuclear envelope and a peripheral network of tubulesand membrane sheets. The tubules are shaped bythe curvature-stabilizing proteins reticulons andDP1/Yop1p, but how the sheets are formed isunclear. Here, we identify several sheet-enrichedmembrane proteins in the mammalian ER, includingproteins that translocate and modify newly synthe-sized polypeptides, as well as coiled-coil membraneproteins that are highly upregulated in cells withproliferated ER sheets, all of which are localized bymembrane-bound polysomes. These results indicatethat sheets and tubules correspond to rough andsmooth ER, respectively. One of the coiled-coilproteins, Climp63, serves as a ‘‘luminal ER spacer’’and forms sheets when overexpressed. More univer-sally, however, sheet formation appears to involvethe reticulons and DP1/Yop1p, which localize tosheet edges and whose abundance determines theratio of sheets to tubules. These proteins maygenerate sheets by stabilizing the high curvature ofedges.

INTRODUCTION

How the characteristic shape of a membrane-bound organelle is

generated is a fundamental question in cell biology. We have

started to address this question for the endoplasmic reticulum

(ER), an organelle that has a particularly intriguing morphology.

It is a continuous membrane system that is comprised of the

nuclear envelope as well as of a peripheral network of tubules

and sheets (Baumann and Walz, 2001; Shibata et al., 2009;

Voeltz et al., 2002). Both the tubules and sheets are dynamic,

i.e., they are continuously forming and collapsing. Previous

work has identified proteins that are responsible for shaping

the tubular ER network (Hu et al., 2008, 2009; Shibata et al.,

2008; Voeltz et al., 2006), but essentially nothing is known about

how ER sheets are generated. In addition, it is unknown whether

proteins specifically segregate into ER sheets and whether there

is a functional significance to the existence of different ER

morphologies.

ER tubules are characterized by high membrane curvature in

cross-section and are shaped by two families of curvature-stabi-

lizing proteins, the reticulons and DP1/Yop1p (Voeltz et al.,

2006). Members of both families are ubiquitously expressed in

all eukaryotic cells. These proteins localize to the ER tubules,

and their depletion leads to the loss of tubules. Conversely, the

overexpression of certain isoforms results in long, unbranched

tubules. Purified members of the two families deform reconsti-

tuted proteoliposomes into tubules (Hu et al., 2008). Together,

these results indicate that the reticulons and DP1/Yop1p are

both necessary and sufficient for ER tubule formation. These

two protein families do not share sequence homology, but

both have a conserved domain containing two long hydrophobic

segments that sit in the membrane as hairpins (Voeltz et al.,

2006). These hairpins may stabilize the high curvature of tubules

in cross-section by forming a wedge in the lipid bilayer. In addi-

tion, oligomerization of these proteins may generate arc-like

scaffolds around the tubules (Shibata et al., 2008).

The peripheral ER sheets vary in size but always consist of two

closely apposed membranes whose distance is approximately

the same as the diameter of the tubules (�30 nm in yeast

[Bernales et al., 2006] and �50 nm in mammals). Consequently,

the edges of sheets have a similarly high curvature as the cross-

section of tubules. In ‘‘professional’’ secretory cells, such as

plasma B cells or pancreatic cells, the ER sheets extend

throughout the entire cell and are studded with membrane-

bound ribosomes. They are stacked tightly with regular

distances between the membranes on both the cytoplasmic

and luminal sides (Fawcett, 1981). By contrast, cells that do

not secrete many proteins contain mostly tubular ER. These

observations have led to the idea that ER sheets correspond to

rough ER (Shibata et al., 2006), the region of the ER that contains

membrane-bound ribosomes, i.e., ribosomes associated with


the translocons, the sites of translocation and modification of

newly synthesized secretory and membrane proteins. On the

other hand, ER tubules would correspond to smooth ER (Shibata

et al., 2006), the ER region devoid of ribosomes, which may be

specialized in lipid metabolism or Ca2+ signaling. While these

ideas are attractive, the tubular ER clearly contains membrane-

bound ribosomes, and a segregation of rough ER proteins into

sheets has not yet been demonstrated.

Several mechanisms of ER sheet formation have been consid-

ered. One possibility is that integral membrane proteins would

form bridges across the luminal space of the ER (Senda and

Yoshinaga-Hirabayashi, 1998; Shibata et al., 2009). A second

possibility is that proteins form flat cytoplasmic or luminal

scaffolds, as suggested for the formation of flat Golgi cisternae

(Short et al., 2005). It has also been proposed that the membrane

association of ribosomes could directly be responsible for the

generation of ER sheets (Puhka et al., 2007). Finally, given that

the reticulons and DP1/Yop1p generate high curvature

membranes, one might imagine that they generate sheets by

stabilizing the sheet edges, bringing the apposing membranes

in close proximity (Shibata et al., 2009).

Here, we show that rough ER proteins partition into ER sheets.

This includes both proteins involved in translocation and modifi-

cation of newly synthesized polypeptides, as well as coiled-coil

membrane proteins that are highly upregulated in cells contain-

ing proliferated ER sheets. Membrane-bound polysomes are

required for the segregation of these rough ER proteins into

sheets, and one of the coiled-coil proteins, Climp63, serves as

a luminal ER spacer. However, neither the polysomes nor the

coiled-coil proteins are essential for sheet formation per se.

Instead, a major mechanism of sheet formation appears to

involve the reticulons and DP1/Yop1p proteins, which can

stabilize the high membrane curvature at sheet edges. Our

results suggest that, in many cells, their abundance is the major

determinant of ER morphology.

RESULTS

Segregation of Proteins into ER SheetsThe different morphologies of the ER imply that, despite the

continuity of the membrane system, some proteins are likely

enriched in certain domains. So far, the only proteins known

with a specific localization are the tubule-preferring reticulons,

DP1/Yop1p, and atlastins/Sey1p (Hu et al., 2009; Shibata

et al., 2008; Voeltz et al., 2006). These proteins localize to tubules

even when highly overexpressed. By contrast, other overex-

pressed ER proteins distribute indiscriminately throughout the

entire ER, making it impossible to draw conclusions about their

endogenous localizations. We therefore first tested whether

several endogenous ER proteins segregate into different ER

domains using immunofluorescence and confocal microscopy

in BSC1 cells. As expected, the luminal ER protein calreticulin,

which is involved in the folding of glycoproteins, was found in

peripheral ER sheets, which are mostly located close to the

nucleus, as well as in the tubular ER network and the nuclear

envelope (Figure 1A). Calreticulin almost perfectly colocalized

with GFP-tagged Sec61b, stably overexpressed in the same

cell. Endogenous Sec61b is part of the Sec61 complex, the

component forming the protein-conducting channel in the ER,

but due to its tagging with GFP and overexpression,

GFP-Sec61b is not associated with the translocon and distrib-

utes throughout the ER (Shibata et al., 2008). Antibodies recog-

nizing the luminal chaperones BiP and Grp94 (anti-KDEL) also

stained the entire ER (Figure 1C, middle). The integral membrane

proteins calnexin and Bap31 showed a similar ubiquitous local-

ization as overexpressed GFP-Sec61b (Figure 1B and Figure S1

available online). These results suggest that many luminal and

membrane ER proteins do not localize to a specific ER domain,

consistent with the continuity of the membrane system.

Next, we tested the endogenous localization of components of

the translocon. In contrast to overexpressed GFP-Sec61b,

endogenous Sec61b was found concentrated in ER sheets

when compared to the localization of the luminal ER proteins

BiP and GRP94 (Figure 1C), although some weak staining of

the tubular network and nonspecific staining of the cytoplasm

were also seen. Because endogenous Sec61b is contained in

the Sec61 complex, these data suggest that translocons are

enriched in ER sheets. This is supported by the localization of

endogenous TRAPa, a component tightly associated with the

ribosome-bound Sec61 complex (Menetret et al., 2008); TRAPa

was strongly enriched in the peripheral ER sheets (Figure 1D).

Finally, Dad1, a component of the translocon-associated

oligosaccharyl transferase complex that glycosylates nascent

secretory and membrane proteins, also showed a similar locali-

zation; GFP-tagged Dad1 that was stably expressed in Dad1-

deficient cells at a level just sufficient to sustain viability (Nikonov

et al., 2002) showed a clear preference for ER sheets, in contrast

to calreticulin in the same cell (Figure 1E). Together, these data

indicate that translocon components are enriched in ER sheets.

To identify additional sheet-segregating proteins that could

potentially be required for sheet formation, we reasoned that

such proteins would be abundant in highly secretory cells that

contain proliferated ER sheets. We therefore identified by mass

spectrometry the most abundant, integral ER membrane

proteins in dog pancreatic rough microsomes. The 25 most

abundant proteins include translocon components, such as

subunits of the oligosaccharyl transferase complex, signal

peptidase, SRP receptor, components of the TRAP complex,

and the Sec61 complex (Table S1). Of interest, the list also

includes p180 and Climp63. Kinectin, which is sequence related

to p180, is somewhat less abundant. All of these proteins have

a single transmembrane segment and an extended coiled-coil

domain, which is located on the luminal side of the ER membrane

in the case of Climp63 and on the cytoplasmic side in the case of

p180 and kinectin (Figure S2A). The molecular function of these

coiled-coil proteins is not well understood. Climp63 has been

implicated in the interaction of ER membranes with microtubules

(Klopfenstein et al., 1998). P180 was originally proposed to be

a ribosome receptor (Savitz and Meyer, 1990); it also interacts

with microtubules (Ogawa-Goto et al., 2007) and is now thought

to play a role in the differentiation of certain monocytic cells (Be-

nyamini et al., 2009). Kinectin was initially identified as a receptor

for the molecular motor kinesin (Toyoshima et al., 1992).

Another way to identify potential sheet-segregating proteins is

to analyze components that are upregulated during the differen-

tiation of immature B cells to IgG-secreting plasma cells, which


involves massive ER sheet proliferation. To identify mRNAs

whose abundance is greatly increased, we sorted through

published microarray data (Luckey et al., 2006). The list of the

25 most upregulated mRNAs coding for ER membrane proteins

(Table S2) includes components of the translocon, of the

unfolded protein response, and of the ER protein degradation

Figure 1. Localization of Proteins to Different ER Domains

(A) The localization of endogenous luminal ER protein calreticulin is compared with that of the stably overexpressed membrane protein GFP-Sec61b using

confocal microscopy in BSC1 cells. Calreticulin was detected with specific antibodies by indirect immunofluorescence (left) and Sec61b by GFP fluorescence

(middle). The right panel shows a merged image. Scale bar, 10 mm.

(B) As in (A) but comparing the localization of the ER membrane protein calnexin with that of GFP-Sec61b.

(C) The localization of endogenous Sec61b is compared to that of the endogenous ER luminal proteins BiP and GRP94 (anti-KDEL), using indirect immunoflu-

orescence with specific antibodies and confocal microscopy.

(D) As in (A) but comparing the localization of the translocon membrane protein TRAPawith that of GFP-Sec61b. Also note that TRAPa is noticeably depleted from

the nuclear envelope.

(E) The localization of stably expressed GFP-Dad1 in a BHK cell line lacking endogenous Dad1 is compared with that of endogenous calreticulin.

All insets show a magnified view of the boxed areas highlighting the junctions between ER sheets and tubules. See also Figure S1.


machinery. It also includes Climp63 and p180 (their mRNAs are

upregulated by a factor of 19–26; kinectin mRNA was not

analyzed). Together with the mass spectrometry data, these

results raise the possibility that the coiled-coil membrane

proteins Climp63, p180, and kinectin localize to ER sheets.

Because these proteins have no known function in protein trans-

location or modification, they are also candidates for being

involved in sheet formation.

Next, we tested whether the coiled-coil proteins are enriched

in ER sheets, using immunofluorescence and confocal micros-

copy. At endogenous levels, all three proteins indeed segregated

to ER sheets, whereas in the same cells, calreticulin distributed

throughout the entire ER (Figures 2A–2C). P180-GFP overex-

pressed at moderate levels was also enriched in ER sheets (Fig-

ure S2B). Thus, in addition to the translocon proteins, at least

three other abundant integral membrane proteins are enriched

in ER sheets. All three proteins were noticeably depleted from

the nuclear envelope (Figure 2 and Figure S2), as reported previ-

ously for Climp63 (Klopfenstein et al., 1998).

A Role for Polysomes in Protein Enrichmentin ER SheetsBecause translocon-associated proteins were found enriched in

ER sheets and are also generally associated with ribosomes, we

tested whether the sheet-preferring proteins are localized by

their association with membrane-bound translating ribosomes.

We treated tissue culture cells with puromycin, a drug that

releases nascent polypeptide chains from ribosomes and

disassembles polysomes; the localization of endogenous

sheet-preferring proteins was subsequently analyzed by immu-

nofluorescence. TRAPa moved into the tubular network (Fig-

ure 3A). Quantification shows that, in untreated cells, TRAPa is

enriched in sheets, as compared to the general ER marker

GFP-Sec61b, but 15 min after puromycin addition, TRAPa was

almost equally abundant in sheets and tubules (Figure 3E). The

disassembly of the polysomes did not abolish the ER sheets,

which in fact occupied a larger surface in many cells (Figure S3A).

To rule out the possibility that inhibition of translation causes the

redistribution of TRAPa, we performed control experiments with

cycloheximide, a drug that inhibits the elongation of polypeptide

chains but leaves polysomes intact. Cycloheximide inhibited

protein synthesis as effectively as puromycin (Figure S4), but

TRAPa stayed in ER sheets (Figures 3B and 3E). All of the other

tested ER sheet-preferring proteins behaved in the same way as

TRAPa (Figures 3C–3E). On the other hand, the localization of

calnexin and Bap31, membrane proteins that did not segregate

into ER sheets, remained unchanged after treatment with either

puromycin or cycloheximide, as was also the case for the luminal

protein calreticulin (Figure 3E and Figures S3B and S3C). Pacta-

mycin, an inhibitor of translation initiation, which allows ribo-

somes to run off the mRNAs, had a similar effect as puromycin

on sheet-segregating proteins, i.e., they were no longer concen-

trated in sheets (Figure S3D). Again, the sheets did not disappear

but often occupied a larger area of the cell (Figure S3E). These

results indicate that polysomes concentrate sheet domains

and localize certain membrane proteins to ER sheets, likely

because these proteins have a direct or indirect affinity for

membrane-bound polysomes.

Climp63 Serves as a ‘‘Luminal ER Spacer’’To test for a possible role of the coiled-coil membrane proteins in

ER morphology, we performed RNAi experiments. The depletion

of Climp63, p180, and kinectin (Figure S5A) either individually or

together did not abolish the existence of ER sheets (Figure 2D

versus 2E). Nevertheless, these proteins have an effect on ER

morphology, as the sheets in depleted cells spread throughout

the cytoplasm (Figures S5B and S5C), similarly to what is

observed when cells are treated with puromycin or pactamycin

(Figure S3). It thus seems that the coiled-coil membrane proteins

are not required for sheet formation per se may but function in

segregating sheet domains close to the cell nucleus.

Thin-section electron microscopy of COS7 cells confirmed

that peripheral ER sheets persist after puromycin treatment or

depletion of Climp63, p180, and kinectin (compare Figures 4B

and 4C with 4A). No bulging of the two membrane sheets was

observed, but of interest, the luminal width was significantly

reduced in triple knockdown cells (from 45–50 nm to 25–30 nm;

Figure 4E). A similar effect was seen when Climp63 alone was

depleted (Figures 4D and 4E), whereas single or double knock-

down of p180 and kinectin had no obvious phenotype (Figure 4E

and data not shown). These results indicate that Climp63 serves

to maintain the normal luminal width of peripheral ER sheets,

likely by forming bridges through their luminal coiled-coil

domains (Klopfenstein et al., 2001). Consistent with a luminal

spacer function, organisms that lack Climp63, including

Drosophila S2 cells (Figure 4E), silkworm (Senda and Yoshi-

naga-Hirabayashi, 1998), and S. cerevisiae (Bernales et al.,

2006), all appear to have narrower ER sheets than mammals. It

should be noted that the distance between the inner and outer

nuclear membranes was unaffected by Climp63 depletion and

was the same in mammalian and insect cells (Figure 4E), consis-

tent with the absence of this protein from the nuclear envelope.

Linking the Formation of ER Sheets and TubulesThe overexpression of Climp63 led to a dramatic proliferation of

ER sheets; we observed a good correlation between the expres-

sion level of a FLAG-tagged version of Climp63 in COS7 cells

and the generation of sheets, an effect that is most strikingly

seen in three-dimensional (3D) reconstructions of the ER (Figures

5A and 5B; quantification in 5C). In thin-section electron micros-

copy, prominent membrane structures were seen that consisted

of anastomosing sheets containing membrane-bound ribo-

somes (Figure 5D). The sheets had a constant luminal width of

�50 nm, and at the highest expression levels, the luminal protein

calreticulin was displaced from areas of Climp63 localization

(Figure S6), consistent with Climp63 filling the luminal space.

We also observed organized smooth ER (OSER) structures in

which the membranes were tightly stacked and the internal

membranes were devoid of ribosomes (Figure S7). Although

these structures are likely caused by oligomerization of the

cytoplasmic GFP tag (Snapp et al., 2003), they differ from normal

OSERs by having a constant luminal spacing of �50 nm.

Given that ER sheet proliferation was also observed when the

curvature-stabilizing reticulons are depleted in mammalian cells

by RNAi (Anderson and Hetzer, 2008) or when the reticulons and

Yop1p are lacking in S. cerevisiae (Voeltz et al., 2006), we tested

whether Climp63 and the reticulons have opposing effects on ER


sheet formation. Indeed, when the reticulon Rtn4b was overex-

pressed in COS7 cells, peripheral ER sheets became diminished

with increasing expression levels (Figures 5E and 5F; quantifica-

tion in Figures 5G and 5H). Concomitant with the decrease in

sheet structures, the normal tubular network was gradually re-

placed with long, unbranched tubules (quantification in Figure 5I).

Figure 2. Membrane Proteins Enriched in ER Sheets

(A) The endogenous localization of the membrane protein Climp63 is compared with that of the luminal ER protein calreticulin in COS7 cells, using indirect

immunofluorescence with specific antibodies. The far-right panel shows a merged image. Junctions between peripheral ER sheets and tubules are highlighted

in the magnified view of the boxed area (inset). Scale bar, 10 mm.

(B) As in (A) but comparing the localization of kinectin (KTN) and calreticulin.

(C) As in (A) but comparing the localization of p180 and calreticulin.

(D) Climp63, p180, and kinectin were depleted in COS7 cells by RNAi (C/P/K siRNA), and Climp63, TRAPa, and calreticulin were visualized using indirect

immunofluorescence with specific antibodies. Scale bar, 10 mm.

(E) As in (D) but with cells transfected with control siRNA oligonucleotides.



When Climp63 and Rtn4a were both highly overexpressed, the

normal ER morphology was almost restored (Figure 5J). Taken

together, these results indicate that Climp63 and the curva-

ture-promoting proteins undergo a ‘‘tug-of-war’’ that determines

the amount of membrane partitioning into these domains.

Curvature-Stabilizing Proteins Localize to Sheet EdgesBecause the reticulons and DP1/Yop1p localize to tubules, one

might expect that they are also found at sheets edges because

these have a similarly high membrane curvature as tubules in

cross-section. Indeed, in many cells, the endogenous reticulons

localized to the edges of sheets, as demonstrated by immunoflu-

orescence using antibodies recognizing both Rtn4a and 4b (Fig-

ure 6A). Similar observations were made in plant cells (Sparkes

et al., 2010). In Climp63-overexpressing cells with proliferated

sheets, Rtn4a/b lined the edges of essentially all sheets in an

even more striking manner (Figure 6B).

To test whether the curvature-stabilizing proteins generally

localize to sheet edges, we tested the localization of a reticulon

in S. cerevisiae. We expressed Rtn1p-GFP from the chromo-

some together with ssRFP-HDEL, a general, luminal ER marker.

Indeed, peripheral ER sheets were generally lined by Rtn1p-GFP

(Figure 6C). The edge localization of Rtn1p-GFP was even more

obvious in cells where ER sheet proliferation was induced by

deletion of the genes encoding the tubule-shaping protein

Yop1p and the GTPase Sey1p (Figure 6D) (Hu et al., 2009).

Similar results were obtained when ER sheets were induced by

deletion of OPI1 (Figure 6E) (Schuck et al., 2009). Thus, as in

mammalian cells, the reticulons localize to the edges of periph-

eral ER sheets. These results indicate that the reticulons stabilize

the high curvature of both tubules in cross-section and of sheet

edges.

A Role for Curvature-Stabilizing Proteinsin Sheet FormationGiven the localization of the curvature-generating proteins to

sheet edges, we considered the possibility that they can

generate sheets by bringing the apposing membranes into close

proximity. In this model, the ratio of sheets and tubules would be

determined by the relative amounts of lipids and curvature-

stabilizing proteins. Indeed, the sheet proliferation seen upon

OPI1 deletion in S. cerevisiae (Figure 6E) is likely caused by an

increase in phospholipid synthesis; Opi1p normally inhibits the

transcription factors Ino2p and Ino4p, which control many

phospholipid synthesis enzymes (Ambroziak and Henry, 1994;

Carman and Henry, 2007). To test whether expression of a curva-

ture-stabilizing protein would convert the sheets into tubules, we

used opi1D cells that express Rtn1p-GFP from the chromosome

as well as the luminal ER marker ssRFP-HDEL. The overexpres-

sion of untagged Rtn1p from a CEN plasmid led to a partial

conversion of sheets into tubules (Figure 6F; quantification in

Figure 6H). When untagged Rtn1p was expressed at a still higher

level from a 2 m plasmid, the sheet-to-tubule ratio converted

back to about the level seen in wild-type cells (Figures 6G

and 6H). These data support the idea that the abundance of

the reticulons determines the relative amounts of sheets and

tubules in the cell.

A Model for the Generation of ER Sheets and TubulesTo test whether the curvature-stabilizing proteins alone could

explain the relative amounts of sheets and tubules in a cell, we

developed a simple theoretical model. We assume that the

reticulons and DP1/Yop1p localize exclusively to tubules and

sheet edges, generating and stabilizing these high curvature

membranes by forming oligomeric scaffolds that are shaped

as rigid arcs. Based on previous estimates, the energetically

optimal distance between the arcs is assumed to be 40 nm (Hu

et al., 2008). The edge membrane can be seen as a half-cylinder,

whose axis bends in the sheet plane forming the sheet circumfer-

ence. The protein-driven formation of a sheet edge enables the

two membranes of a sheet to adopt planar shapes (Figure 7A).

A tubule forms by self-folding of a part of the edge into

a complete cylinder and therefore represents an edge extension

(Figure 7A). We assume negligible bulging between the arc-like

scaffolds, as supported by previous results (Hu et al., 2008),

and a diameter of 30 nm for both sheet edges and tubules (Fig-

ure 4) (Bernales et al., 2006).

Our model calculates for a given membrane surface area the

total length of the tubules and the shape and dimensions of the

sheets in dependence of the number of curvature-stabilizing

proteins, Nc. We characterize the edge length by a parameter

G = Le/ Le0, wherein Le

0 is the circumference of a flat circular

disc with the same overall membrane area (i.e., G = 1 for a flat

disc). G is proportional to Nc (Supplemental Information). In our

calculations, we assume that Nc is at least large enough to

generate a circular sheet (G R 1).

For each G value, we computed the overall membrane shape

by minimizing the energy of the edge bending in the sheet plane

(see Experimental Procedures and Supplemental Information).

The top view of the shapes is presented in Figure 7B. Starting

from the circular disc configuration at G = 1 (Figure 7B, blue

line), the sheet shape elongates with increasing G (and Nc) (light

blue line) and then acquires a flattened dumbbell appearance

with a narrowing neck (aqua and yellow lines) and, finally, at

G�2, splits into two droplet-like sheets with a short tubule

between them (orange line). Further increase of G results in

tubule elongation and a decrease in the sizes of the two sheets

(Figure 7B, dark red line). Eventually, the whole membrane

converts into a tubule (not shown in Figure 7B). Thus, the curva-

ture-producing proteins alone can generate both sheets, and

tubules and their abundance determines the relative amounts

of the two membrane domains.

Next, we extended the model to include the effect of proteins

enriched in sheets. We assume that polysome-bound Climp63,

p180, and kinectin, as well as translocons, can diffuse

throughout the sheets but cannot move into high curvature

membrane areas, i.e., sheet edges and tubules. The number of

all of these ‘‘sheet proteins’’ together is denoted byNs. The sheet

proteins may be considered as generating an ‘‘osmotic pres-

sure’’ on the sheet edges, a force that resists the shrinkage of

a sheet domain. The magnitude of this effect is determined by

the interplay between the effective ‘‘osmotic pressure’’

produced by the sheet proteins and the effective stretching

elasticity of the edge, the latter being determined by the curva-

ture-stabilizing proteins (see Experimental Procedures and

Supplemental Information). Our model does not take into


Figure 3. Polysome-Dependent Membrane Protein Enrichment in ER Sheets

(A) The localization of the translocon component TRAPa is compared with that of stably expressed GFP-Sec61b after 15 min of treatment with puromycin (PURO).

The far-right panel shows a merged image. Junctions between peripheral ER sheets and tubules are highlighted in the magnified view of the boxed area (inset).

Scale bar, 10 mm.

(B) As in (A) but after 15 min of treatment with cycloheximide (CHX).

(C) As in (A) but comparing the localization of Climp63 with calreticulin after puromycin treatment.

(D) As in (C) but after cycloheximide treatment.

(E) Quantification of sheet enrichment of different ER proteins in untreated cells (blue bars) and in cells treated with puromycin (PURO; green) or cycloheximide

(CHX; red). The ratio of the average fluorescence intensity in sheets versus tubules was determined for calnexin (CNX), Bap31, calreticulin (CRT), TRAPa, and

kinectin and was divided by the sheet-to-tubule fluorescence ratio for stably expressed GFP-Sec61b, a protein that shows no preference for either ER domain.


account that Climp63 affects sheet formation by serving as a

luminal spacer, and it does not make any assumptions about

the specific roles of p180 and kinectin.

We computed the G values and membrane configurations for

different values of Nc and Ns (Figure 7C). The colored lines on the

bottom plane of the diagram represent the relationship between

Nc and Ns for a given shape of the system, with the colors

corresponding to the shapes as in Figure 7B. Figure 7C demon-

strates that an increase of Nc at a given Ns results in larger G

(blue to red transition) and thus in more tubules, whereas an

increase of Ns at a given Nc results in lower G, i.e., more sheets.

This is further illustrated in Figure 7D, in which the increase of

Ns at a constant Nc converts two small sheet areas connected

by a narrow tubule into a larger sheet area. Thus, the model reca-

pitulates the experimental observation of a tug-of-war between

sheet-promoting Climp63 and curvature-stabilizing proteins.

DISCUSSION

Our results indicate that several mechanisms shape peripheral

ER sheets. The most basic and universal mechanism appears

to involve the previously identified curvature-stabilizing proteins,

the reticulons and DP1/Yop1p. These proteins would stabilize

not only the high curvature of narrow tubules, but also the

Figure 4. Climp63 Affects the Luminal Width of Peripheral ER Sheets

(A) Rough ER sheets in a COS7 cell visualized by thin-section electron microscopy. Scale bar, 0.5 mm.

(B) As in (A) but after treatment with puromycin (PURO) for 15 min.

(C) As in (A) but after RNAi-depletion of Climp63, p180, and kinectin (C/P/K siRNA).

(D) As in (A) but after RNAi-depletion of Climp63.

(E) Quantification of the luminal width of peripheral ER sheets and the nuclear envelope (NE) in differently treated COS7 cells. For comparison, Drosophila S2R+

cells were also analyzed. Shown are the means and standard errors of n cells analyzed for each sample. Kinectin, KTN.

A similar analysis was done for GFP-Dad1 and Climp63 but with calreticulin as reference. Shown are the means and standard errors of data obtained from 7 to

30 cells for each condition.



Figure 5. Climp63 and Reticulon Overexpression Change the Abundance of Sheets and Tubules

(A) FLAG-Climp63 overexpressed at relatively high levels in a COS7 cell was visualized by indirect immunofluorescence using FLAG antibodies. A 3D image was

generated from a complete series of z sections (step size 0.25 mm) taken with a confocal microscope. Scale bar, 10 mm.

(B) As in (A) but in a cell expressing FLAG-Climp63 at the highest observed levels.

(C) Quantification of the effect of Climp63 overexpression on ER sheet abundance. Shown are the percentages of cells with normal reticular ER (blue bars), of cells

with both large sheets and reticular ER (red), and of cells with large ER sheets lacking reticular ER (green) at different expression levels of FLAG-Climp63. The cells

were divided into five groups according to their expression levels, as determined by overall average fluorescence intensity.

(D) Thin-section electron micrograph of a COS7 cell overexpressing GFP-Climp63. The inset shows an enlargement of the boxed region. Scale bar, 0.5 mm.

(E) HA-Rtn4b (red) was expressed in COS7 cells at relatively low levels and was localized with HA antibodies by indirect immunofluorescence and confocal

microscopy. Endogenous Climp63 (green) was localized in the same cells with specific antibodies.

(F) As in (G) but with the highest observed expression level of HA-Rtn4b. Note that Climp63 appears in bright punctae and in the nuclear envelope.


curvature of sheet edges, a mechanism that is sufficient to keep

the two membranes of a sheet closely apposed. The reticulons

and DP1/Yop1p probably stabilize high curvature by two mech-

anisms, ‘‘hydrophobic insertion/wedging’’ and ‘‘scaffolding’’

(Shibata et al., 2009). The conserved segments of these proteins

may form a wedge in the lipid bilayer that occupies more space in

the cytoplasmic leaflet than in the luminal leaflet. Oligomerization

of these proteins may generate scaffolds around curved

membranes, which may take the shape of open arcs, given

that they can localize to sheet edges. Our theoretical model

demonstrates that the reticulons and DP1/Yop1p alone can

generate both tubules and sheets, with their abundance deter-

mining the ratio of these domains. Consistent with the proposed

dual role of the reticulons and DP1/Yop1p in tubule and sheet

formation, they localize to both tubules and sheet edges, their

depletion leads to increased sheet areas, and their overexpres-

sion converts sheets into tubules.

In S. cerevisiae, the amount of membrane surface and the

abundance of the reticulons and Yop1p appear to be the deci-

sive factors determining the ratio of peripheral ER sheets and

tubules. Generating more lipid increases the sheet area, whereas

increasing the abundance of the curvature-stabilizing proteins

increases the number of tubules. The observation of sheets in

cells lacking the reticulons and Yop1p may be explained by

the presence of other low-abundance curvature-promoting

proteins or by the association of the cortical ER with the plasma

membrane. Although we cannot exclude the existence of sheet-

promoting proteins in yeast, the current data are consistent with

a model in which curvature-stabilizing proteins are the major

determinant of peripheral ER morphology.

Our data suggest that, in mammalian cells, there are several

additional factors that determine the morphology of peripheral

ER sheets. This includes the coiled-coil membrane protein

Climp63, which serves as a luminal spacer. After its depletion,

the luminal width of the sheets decreases from �50 to �30 nm,

a spacing that is also seen in organisms that lack the protein.

Climp63 is highly upregulated in mammalian cells with prolifer-

ated ER sheets, and it induces sheets at the expense of tubules

when overexpressed in tissue culture cells. Thus, at high

concentrations, Climp63 appears to generate sheets all by itself,

and the lack of extensive sheet edges may make the contribution

of the curvature-stabilizing proteins less important. However,

with luminal spacers alone, one would expect bulging of the

sheet edges, in contrast to our observations (Figure 4), indicating

that the curvature-stabilizing proteins may have a role even in

cells with proliferated ER sheets. Climp63’s function may be to

optimize the size of the luminal space of peripheral ER sheets,

such that sufficient luminal chaperones can be accommodated

and the sheets are packed into a minimal space.

Our analysis also identified two other coiled-coil membrane

proteins, p180 and kinectin, with a potential role in shaping ER

sheets. These proteins are enriched in sheets and abundant

in cells with proliferated ER sheets. Overexpression of p180

has been reported to induce sheets in S. cerevisiae and in a

monocytic cell line (Becker et al., 1999; Benyamini et al., 2009),

although in our own experiments and those of others, the effects

were smaller (Ueno et al., 2010 and data not shown). The deple-

tion of p180 and kinectin had no effect on ER sheet morphology.

Although the precise role of these proteins remains to be estab-

lished, all coiled-coil membrane proteins could stabilize sheets

simply by being excluded from high-curvature regions, as shown

by our theoretical considerations. They may be considered as

generating an ‘‘osmotic pressure,’’ a force that counteracts the

shrinkage of sheet domains. Consistent with experimental

observations for Climp63, the coiled-coil proteins are predicted

to be in a tug-of-war with the reticulons and DP1/Yop1p, with

the former shifting the balance toward sheets and the latter

toward tubules. In this model, it does not actually matter how

proteins are excluded from tubules and sheet edges. Given

that all identified sheet-promoting proteins contain extended

coiled-coil domains, they all have the propensity to oligomerize,

which may contribute to their exclusion from high-curvature

regions.

The coiled-coil membrane proteins are not essential for sheet

formation per se, as is obvious from our observation that their

depletion by RNAi does not abolish ER sheets. This suggests

that, like in yeast, the reticulons and DP1/Yop1p may provide

the basic mechanism by which both sheets and tubules are

generated. Consistent with this hypothesis, Climp63, p180,

and kinectin are not known in lower organisms, in contrast to

the reticulons and DP1/Yop1p, which are present in all

eukaryotes.

All sheet-enriched proteins tested, including translocon

components and the coiled-coil membrane proteins, appear

to be concentrated by membrane-bound polysomes; upon

polysome disassembly, all of these proteins distribute equally

between sheets and tubules throughout the cell. Thus, these

proteins must have a direct or indirect affinity for membrane-

bound polysomes. Indeed, several of the tested sheet-prefer-

ring proteins are known to be associated with membrane-

bound translating ribosomes, including components of the

Sec61 complex, the TRAP complex, the oligosaccharyl

(G) Quantification of the peripheral ER sheet area relative to the total ER area for different expression levels of HA-Rtn4b. The areas of ER sheets and tubules were

determined from the fluorescence of Climp63 and Rtn4b, respectively, after subtraction of background. The cells were divided into five groups according to their

expression levels of HA-Rtn4b, as determined by overall average fluorescence intensity, and the mean and standard error were calculated for each group.

(H) Quantification of the effect of Rtn4b overexpression on ER sheet morphology, as determined by Climp63 staining. Shown are the percentages of cells with

normal ER sheets (blue bars), of cells with disc-like ER sheets (red), and of cells with punctae (green) at different expression levels of Rtn4b. The cells were divided

into five groups according to their expression levels.

(I) Quantification of the effect of Rtn4b overexpression on ER tubule morphology, as determined by HA-Rtn4b staining. Shown are the percentages of cells with

normal reticular ER (blue bars), of cells with an abnormally dense ER network (red), and of cells with unbranched, long tubules (green) at different expression levels

of Rtn4b. The cells were divided into five groups according to their expression levels.

(J) Myc-Rtn4a and FLAG-Climp63 were both highly expressed in COS7 cells. The far-right panel shows a merged image. Note that the ER morphology is

almost normal.



Figure 6. The Reticulons Localize to the Edges of ER Sheets

(A) The localization of endogenous Rtn4a and 4b is compared with that of Climp63 using indirect immunofluorescence with specific antibodies in COS7 cells. The

insets show enlargements of the boxed region. Arrows point to reticulons lining the sheets. The far-right panel shows merged images. Scale bar, 10 mm.

(B) As in (A) but with cells overexpressing FLAG-Climp63.

(C) Rtn1p-GFP (green) and ssRFP-HDEL (red) were coexpressed in wild-type S. cerevisiae cells, and the cortical ER was visualized by fluorescence microscopy.

Scale bar, 5 mm.

(D) As in (C) except that the cells had proliferated ER sheets caused by deletion of SEY1 and YOP1 (sey1Dyop1D).

(E) As in (C) except the cells had proliferated ER sheets caused by deletion ofOPI1 (opi1D). The cells also contained an empty vector as a control for panels (F) and (G).

(F) As in (E) except that untagged Rtn1p was expressed under the endogenous promoter from a CEN plasmid.

(G) As in (E) except that untagged Rtn1p was expressed under the endogenous promoter from a 2 m plasmid.

(H) Quantification of the experiments in (C) and (E–G). The relative area of ER sheets was determined from the area of ssRFP-HDEL fluorescence that did not coloc-

alize with Rtn1p-GFP fluorescence and was divided by the total area of ssRFP-HDEL fluorescence. 14 to 38 cells were analyzed per condition, and the means and

standard errors were calculated.


transferase complex, and p180 (Gorlich and Rapoport, 1993).

These proteins stay bound to ribosomes upon detergent solu-

bilization of rough ER membranes, but they can be released

from the ribosomes by puromycin/high salt treatment. Climp63

and kinectin are not bound to detergent-solubilized translocons

(data not shown), so how they are recruited remains to be

clarified.

Our results indicate that ER sheets correspond to rough

ER and tubules to smooth ER. We propose that the assembly

of translating membrane-bound ribosomes into polysomes

concentrates the associated membrane-proteins, including

Climp63, p180, and kinectin. Their concentration might facilitate

their higher-order oligomerization, which may be required for

their exclusion from high-curvature areas and thus for their

Figure 7. Modeling of the Effect of Curvature-Stabilizing and Sheet-Promoting Proteins on ER Morphology

(A) The reticulons and DP1/Yop1p (yellow arcs) are assumed to localize exclusively to tubules and sheet edges, generating and stabilizing these high-curvature

membranes. Stabilization of sheet edges enables the upper and lower membranes of the sheet to adopt planar shapes.

(B) Top view of membrane shapes computed by the theoretical model for increasing G values. The computation was performed for a total membrane area cor-

responding to 1 mm radius of the initial disc-like shape, a 15 nm cross-section radius of the tubules and edges, and a 40 nm optimal distance between the arc-like

proteins at the edge (see Supplemental Information). Change ofG from 1 to 2.1(blue to red) corresponds to increasing the number of curvature-stabilizing proteins

Nc from 140 to 290.

(C) G values and membrane shapes were calculated for different numbers of curvature-stabilizing and sheet-promoting proteins, Nc and Ns. The colors corre-

spond to the membrane shapes shown in Figure 7B. The colored lines on the bottom plane of the diagram represent the relationship betweenNc andNs for a given

shape of the system.

(D) G values and membrane shapes were computed for different Ns values at Nc = 290. The shapes refer to Ns = 0, 500, and 1000.


sheet-promoting function. Once sheets are formed, the

membrane binding of polysomes would be facilitated. Poly-

somes often form spirals that could have an inherent preference

for associating with ER sheets (Christensen and Bourne, 1999);

whereas individual ribosomes or small polysomes can bind

to narrow tubules, it is unlikely that each ribosome of a large

polysome could be efficiently arranged on a narrow tubule. The

binding of large polysomes could therefore be restricted to

membrane sheets. The assembly of membrane-bound poly-

somes would concentrate more coiled-coil membrane proteins,

and these in turn would generate more sheet area by the

‘‘osmotic effect,’’ allowing more polysomes to bind, and so on,

a mechanism that would ultimately lead to a segregated rough

ER domain. This model is consistent with the observation that

the disassembly of polysomes or the depletion of Climp63

increases the mobility of translocons in the plane of the

membrane (Nikonov et al., 2007; Nikonov et al., 2002). It also

agrees with our results showing that the disassembly of poly-

somes leads to the spreading of ER sheets similar to that

seen upon depletion of the sheet-promoting proteins. Our model

explains the classic observation that, in many cells, membrane-

bound ribosomes are not randomly distributed throughout the

ER but, rather, concentrated in a separate membrane domain,

the rough ER. An active sorting of proteins into the rough ER is

consistent with previous cell fractionation experiments, which

demonstrated that general ER proteins indiscriminately

distribute throughout the ER, whereas translocon-associated

proteins are enriched in the rough ER (Hinman and Phillips,

1970; Kreibich et al., 1978; Vogel et al., 1990).

The nuclear envelope is a prominent ER domain whose struc-

ture is determined independently of the peripheral ER. Although

the reticulons have been implicated in the assembly of the

nuclear envelope and in the insertion of nuclear pores (Anderson

and Hetzer, 2008; Dawson et al., 2009), they are nearly absent

from the nuclear envelope, and their depletion or overexpression

has no significant effect on this domain’s morphology. Similarly,

DP1/Yop1p or the coiled-coil membrane proteins Climp63,

p180, and kinectin are also nearly absent from the nuclear enve-

lope and have no obvious effect on its structure. Of interest,

TRAPa was also depleted from the nuclear envelope, raising

the possibility that translocons are preferentially located in

peripheral ER sheets. Thus, distinct mechanisms may determine

the formation and function of the sheet-like domains of the

nuclear envelope and peripheral ER.

In summary, our results lead to a simple model, according to

which the basic morphological elements of the peripheral ER,

the tubules and sheets, are generated by the curvature-

stabilizing proteins. Superimposed on this mechanism, mem-

brane-bound polysomes and associated coiled-coil membrane

proteins may cooperate to form segregated rough ER sheets in

mammalian cells, domains that are functionally specialized in

protein translocation. Other factors probably contribute to the

morphology of the peripheral ER. Microtubules keep the

mammalian ER under tension and stabilize membrane tubules,

but they could also potentially form an additional scaffold that

stabilizes sheets, as suggested by the fact that both Climp63

and p180 are microtubule-binding proteins (Klopfenstein et al.,

1998; Ogawa-Goto et al., 2007). It will be interesting to elucidate

how these factors collaborate with the identified membrane-

shaping principles.


Mammalian Tissue Culture and Transfections

BSC1 cells stably expressing GFP-Sec61b and COS7 cells were grown in

DMEM containing 10% fetal bovine serum at 37�C and 5% CO2 and were

passaged every 2–3 days. GFP-Dad1 BHK cells (M3/18; Nikonov et al., 2002)

were maintained in10%CO2 at 39.5�C todegradeendogenousDad1.For trans-

lation inhibition experiments, cells were treated with 200 mM cycloheximide,

200 mM puromycin, or 100 nM pactamycin in complete media for 15 min.

To deplete Climp63, kinectin, and p180, COS7 cells were plated onto

acid-washed coverslips at 20% confluency and were transfected with

120 nM total siRNA using Oligofectamine (Invitrogen). After1.5 days, cells

were retransfected with the same amount of siRNA oligonucleotides and

then processed for immunofluorescence 1.5 days afterward. Experiments

with control siRNA oligonucleotides (QIAGEN) were done in parallel using

the same conditions. Transient DNA transfections were performed using

Lipofectamine 2000 (Invitrogen). See Supplemental Information for a list of

DNA and siRNA constructs.

Indirect Immunofluorescence and Confocal Microscopy

Indirect immunofluorescence with mammalian cells was done as described

(Shibata et al., 2008). Cells grown on acid-washed coverslips were fixed

with 4% paraformaldehyde (EMS), permeabilized with 0.1% Triton X-100

(Pierce), and immunostained with various primary antibodies and then washed

in PBS and probed with various fluorophore-conjugated secondary anti-

bodies. See Supplemental Information for a list of antibodies used.

Images were captured using a Yokogawa spinning-disk confocal on a Nikon

TE2000U inverted microscope with a 603 or 1003 Plan Apo NA 1.4 objective

lens, a Hamamatsu ORCA ER-cooled CCD camera, and MetaMorph software.

All analyses/quantifications were done on raw 16 bit images using MetaMorph.

For presentation, brightness levels were adjusted across the entire image and

were changed from 16 to 8 bits using Adobe Photoshop. Quantification was

performed as described in Supplemental Information.

Thin-Section Electron Microscopy

Thin-section EM experiments were performed as described previously

(Shibata et al., 2008) except that cells were fixed directly in culture plates.

Quantification was performed as described in Supplemental Information.

Microscopy and Image Quantification of S. cerevisiae Cells

Yeast strains and constructs used are described in Supplemental Information.

Yeast cells were imaged live in complete medium at room temperature using

an Olympus BX61 microscope, UPlanApo 100 3 /1.35 lens, Qimaging Retiga

EX camera, and IVision version 4.0.5 software. To calculate relative peripheral

sheet amounts, cortical ER images of cells expressing ssRFP-HDEL and

Rtn1-GFP were taken. Images were thresholded above background, and the

percentage of sheet area was calculated for each cell as the percentage of

area of ssRFP-HDEL that did not overlap with Rtn1-GFP using Metamorph

software. Means and standard errors were calculated using Microsoft Excel.

For presentation, brightness levels were adjusted across the entire image,

changed from 16 to 8 bits, and cropped using Adobe Photoshop.

Identification of Abundant Coiled-Coil Membrane Proteins

Mass spectrometry of dog pancreatic microsomal proteins and identification

of mRNAs coding for ER membrane proteins that are upregulated during

B cell differentiation (Luckey et al., 2006) were performed as described in

the Supplemental Information.

Modeling of Sheet versus Tubule Generation

To compute the membrane configurations (the length of the tubule as well as

the areas and shapes of the sheets) in dependence of the numbers of the

curvature-producing, Nc, and the sheet-promoting proteins, Ns, we minimize

the system energy, Ftot, for the given total membrane area, Atot. The total

energy Ftot consists of three contributions: the effective stretching energy of


the edge, Fs; the energy of the effective osmotic pressure of the sheet-

promoting proteins, Fp; and the energy of edge bending in the sheet plane, Fb.

The energy Fs is given by Fs =12kBTNc

hðLe�Ncðl0 + laÞÞ

Ncl0

i2

, wherein, Le is the total

length of the edge including the tubules; l0 is the energetically preferred

distance between the arc-like proteins measured along the edge; la is the width

of one protein arc; and kBTz4$10�21 Joule is the product of the Boltzmann

constant and the absolute temperature. According to this expression, the

length of the edge in a stress-free state is L�e =Ncðl0 + laÞ, and the effective

rigidity of the edge stretching-compression with respect to L�e is

kstr = kBT$Nc. Based on previous estimates, we take l0 = 40nm and la = 4nm

(Hu et al., 2008).

The osmotic pressure energy Fp is given by Fp = kBT$Ns$lnðNsb=AflatÞ,wherein Aflat is the flat area available to the sheet proteins and b is the area

of one sheet protein. The area Aflat is related to the total area and length of

the edge by Aflat =12ðAtot � a$LeÞ, wherein a is the membrane area absorbed

by a unit length of the edge, which can be estimated as a=p$Rez50nm

(Rez15nm is the radius of the edge cross-section), and Atot is the total

membrane area.

The energy Fb is given by Fb =12B#c

2edLe, wherein ce is the in-plane curvature

of the edge, and B is the modulus of the edge in-plane bending, which can be

estimated using the membrane bending modulus kz20kBT (Helfrich, 1973) as

BzpRekz900kBT$nm. The integration is performed over the whole edge

length, including the tubules.

Estimates supported by numerical computations show that the total length

of the edge Le and the corresponding value of the parameter G are determined

by the energies Fs and Fp and are largely independent of Fb. At the same time,

the system configuration resulting from minimization of Fb depends of the

parameter G. Therefore, we determine the system configuration in two steps.

First, we minimize the sum of Fc + Fs with respect to Le for every set of

numbers Nc and Ns and determine the corresponding function G (Nc, Ns).

Second, for every value ofG (Nc,Ns), we minimize Fb with respect to the system

shape and find the equilibrium configuration.

The Supplemental Information gives a more detailed discussion of the

model.


Supplemental Information includes Extended Experimental Procedures,

seven figures, and two tables and can be found with this article online at

doi:10.1016/j.cell.2010.11.007.

ACKNOWLEDGMENTS

We thank C. Denison, J. Minsteris, and S. Gygi for mass spectrometry analysis;

J. Baughman for microarray analysis; A. Condon and A. Boye-Doe for

technical assistance; J. Iwasa for help with illustrations; G. Kreibich,

K. Ogawa-Goto, L. Lu, and R. Yan for materials; the Nikon Imaging Center

and the Electron Microscopy facility at HMS for microscopy assistance; and

R. Klemm and A. Osborne for critical reading of the manuscript. Y.S. was sup-

ported by the American Heart Association and is a Howard Hughes Medical

Institute postdoctoral fellow. W.A.P. is supported by the Intramural Research

Program of the National Institute of Diabetes and Digestive and Kidney

Diseases. T.A.R. is a Howard Hughes Medical Institute Investigator. M.M.K.

is supported by the Israel Science Foundation (ISF) and the Marie Curie

network ‘‘Virus Entry.’’





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Abortive HIV Infection MediatesCD4 T Cell Depletion and Inflammationin Human Lymphoid TissueGilad Doitsh,1 Marielle Cavrois,1,5 Kara G. Lassen,1,5 Orlando Zepeda,1 Zhiyuan Yang,1 Mario L. Santiago,1,4

Andrew M. Hebbeler,1 and Warner C. Greene1,2,3,*1Gladstone Institute of Virology and Immunology, 1650 Owens Street, San Francisco, CA 94158, USA2Department of Medicine3Department of Microbiology and ImmunologyUniversity of California, San Francisco, CA 94143, USA4Present address: University of Colorado, Denver, Aurora, CO 80045, USA5These authors contributed equally to this work*Correspondence: [email protected]

DOI 10.1016/j.cell.2010.11.001

SUMMARY

The mechanism by which CD4 T cells are depletedin HIV-infected hosts remains poorly understood.In ex vivo cultures of human tonsil tissue, CD4T cells undergo a pronounced cytopathic responsefollowing HIV infection. Strikingly, >95% of thesedying cells are not productively infected but insteadcorrespond to bystander cells. We now show thatthe death of these ‘‘bystander’’ cells involves abor-tive HIV infection. Inhibitors blocking HIV entry orearly steps of reverse transcription prevent CD4T cell death while inhibition of later events in the virallife cycle does not. We demonstrate that the nonper-missive state exhibited by themajority of resting CD4tonsil T cells leads to accumulation of incompletereverse transcripts. These cytoplasmic nucleic acidsactivate a host defense program that elicits a coordi-nated proapoptotic and proinflammatory responseinvolving caspase-3 and caspase-1 activation. Whilethis response likely evolved to protect the host, itcentrally contributes to the immunopathogeniceffects of HIV.

INTRODUCTION

Despite extensive efforts over the past quarter century, the

precise mechanism by which HIV-1 causes progressive deple-

tion of CD4 T cells remains debated. Both direct and indirect

cytopathic effects have been proposed. When immortalized

T cell lines are infected with laboratory-adapted HIV-1 strains,

direct CD4 T cell killing predominates. Conversely, in more

physiological systems, such as infection of lymphoid tissue

with primary HIV-1 isolates, the majority of dying cells appear

as uninfected ‘‘bystander’’ CD4 T cells (Finkel et al., 1995; Jekle

et al., 2003).

Various mechanisms have been proposed to contribute to

the death of these bystander CD4 T cells including the action

of host-derived factors like tumor necrosis factor-a, Fas ligand

and TRAIL (Gandhi et al., 1998; Herbeuval et al., 2005), and viral

factors like HIV-1 Tat, Vpr, and Nef released from infected cells

(Schindler et al., 2006; Westendorp et al., 1995). Considerable

interest has also focused on the role of gp120 and gp41 Env

protein in indirect cell death, although it is not clear whether

death signaling involves gp120 binding to its chemokine

receptor or gp41-mediated fusion. It is also unclear whether

such killing is caused by HIV-1 virions or by infected cells

expressing Env.

Most studies have focused on death mechanisms acting prior

to viral entry. Less is known about the fate of HIV-1-infected CD4

T cells that do not express viral genes, in particular naive CD4

T cells in tissue that are refractory to productive HIV infection

(Glushakova et al., 1995; Kreisberg et al., 2006). In these cells,

infection is aborted after viral entry, as reverse transcription is

initiated but fails to reach completion (Kamata et al., 2009;

Swiggard et al., 2004; Zack et al., 1990; Zhou et al., 2005).

Human lymphoid aggregated cultures (HLACs) prepared from

tonsillar tissue closely replicate the conditions encountered by

HIV in vivo and thus form an attractive, biologically relevant

system for studying HIV-1 infection (Eckstein et al., 2001).

Lymphoid organs are the primary sites of HIV replication and

contain more than 98% of the body’s CD4 T cells. Moreover,

events critical to HIV disease progression occur in lymphoid

tissues, where the network of cell-cell interactions mediating

the immune response deteriorates and ultimately collapses.

Primary cultures of peripheral blood cells do not fully mimic

the cytokine milieu, the cellular composition of lymphoid tissue,

nor the functional relationships that are undoubtedly important

in HIV pathogenesis. Finally, HLACs can be infected with

a low number of viral particles in the absence of artificial mito-

gens, allowing analysis of HIV cytopathicity in a natural and

preserved environment. In this study, we used the HLAC system

to explore the molecular basis for HIV-induced killing of CD4

T cells.


RESULTS

Selective Depletion of CD4 T Cells by X4-Tropic HIV-1To explore depletion of CD4 T cells by HIV-1, HLACs made from

freshly dissected human tonsillar tissues were infected with

a GFP reporter virus (NLENG1), prepared from the X4-tropic

NL4-3 strain of HIV-1. This reporter produces fully replication-

competent viruses. An IRES inserted upstream of the Nef gene

preserves Nef expression and supports LTR-driven GFP expres-

sion (Levy et al., 2004), allowing simultaneous quantification of

the dynamics of HIV-1 infection and T cell depletion. NL4-3

was selected because tonsillar tissue contains a high per-

centage of CD4 T cells expressing CXCR4 (90%–100%).

Productively infected GFP-positive cells appeared in small

numbers 3 days after infection, peaked on days 6–9, and

decreased until day 12, when few CD4 T cells remained in the

culture (Figure 1). Fluorescence-linked antigen quantification

(FLAQ) assay of HIV-1 p24 (Hayden et al., 2003) confirmed the

accumulation of viral particles in the medium between day 3

and days 8 to 9, when a plateau was reached (data not shown).

Interestingly, when HIV-1 p24 levels plateaued no more than

1.5% of all cells (about 5% of CD4 T cells) were GFP-positive.

Figure 1. Massive Depletion of CD4 T Cells in HLACs Containing Small Number of Productively Infected Cells

Kinetics of spreading viral infection versus depletion of CD4 T cells after infection of HLACs with a replication-competent HIV reporter virus encoding GFP.

CD4 downregulation in GFP-positive cells likely represents the combined action of the HIV Nef, Vpu, and Env proteins expressed by this virus. Ratios of viable

CD4 versus CD8 T cells in HIV-infected and uninfected cultures are also shown. Flow cytometry plots represent live-gated cells, based on the forward-scatter

versus side-scatter profile of the complete culture. These data are the representative results of six independent experiments utilizing tonsil cells from six different

donors.


However, although the number of CD4 T cells was not markedly

altered in infected cultures through six days, the culture was

almost completely devoid of CD4 T cells by day 9. CD8 T cells

were not depleted in infected cultures, and CD4 T cells were

not depleted in uninfected cultures. These findings reveal

marked and selective depletion of CD4 T cells in HLAC cultures.

However, due to the nature of the assay, we could not definitively

conclude whether the principal mechanism of depletion involved

direct or indirect effects of HIV-1.

Extensive Depletion of Nonproductively Infected CD4 TCells in HLACsTo determine if indirect killing (formerly indicated as ‘‘bystander’’)

of CD4 T cells accounted for most of the observed cellular

depletion, we took advantage of a reported experimental

strategy (Jekle et al., 2003) that unambiguously distinguishes

between the death of productively and nonproductively infected

cells (Figure 2A). After 6 days of coculture, survival analysis

of CFSE-labeled cells by flow cytometry (Figure 2B) showed

Figure 2. CD4 T Cell Depletion in HIV-1-Infected HLACs Predominantly Involves Nonproductively Infected Cells

(A) Experimental strategy to assess indirect cell killing in HIV-1-infected human lymphoid cultures. Fresh human tonsil tissue from a single donor is processed into

HLAC, and then separated into two fractions. One fraction is challenged with HIV-1 and cultured for 6 days, allowing viral spread. On day 5, the uninfected fraction

is treated with AZT (5 mM) and labeled with CFSE (1 mM). On day 6, the infected and CFSE-labeled cultures are mixed and cocultured in the presence of AZT.

Because of its site of action, AZT does not block viral output from the HIV-infected cells but prevents productive infection of CFSE-labeled cells. After 6 days

of coculturing, the number of viable CFSE-positive cells is determined by flow cytometry.

(B) Flow cytometry analysis of the mixed HLACs. Indirect killing is determined by gating on live CFSE-positive cells in the mixed cultures. Effector cells are either

infected or uninfected cells.

(C) Extensive depletion of nonproductively infected CD4 T cells by HIV-1. CFSE-labeled cells mixed with uninfected or infected cells were cultured in the presence

of 5 mM AZT alone or together with 250 nM AMD3100. Data represent live CFSE-positive cells 6 days after coculture with infected or uninfected effector cells. The

absence of productive infection in the CFSE-positive cells was confirmed by internal p24 staining and monitoring GFP expression following infection with the

NLENG1 HIV-1 reporter virus (data not shown).

(D) Preferential depletion of nonproductively infected CD4 T cells by HIV-1. The absolute numbers of viable CFSE-positive CD4 and CD8 T cells and B cells were

determined. Percentages are normalized to the number of viable CFSE-positive cells cocultured with uninfected cells in the presence of AZT, as depicted by an

asterisk. Error bars represent standard deviations of three samples from the same donor. This experiment is representative of more than 10 independent

experiments with more than 10 donors of tonsillar tissues.

See also Figure S1.


extensive depletion of CD4 T cells in cultures mixed with HIV-

infected cells but not in those mixed with uninfected cells (Fig-

ure 2C). The relative proportion of CD8 T cells was not altered.

CD3+/CD8– T cells were similarly depleted, indicating that the

loss was not an artifact of downregulated surface expression

of CD4 following direct infection. Loss of CFSE-labeled CD4

T cells was prevented by AMD3100, which blocks the engage-

ment of gp120 with CXCR4, but not by the reverse transcriptase

inhibitor AZT. Thus, productive viral replication is not required for

CD4 T cell death.

To estimate the absolute numbers of all CFSE-labeled cell

subsets, we added a standard number of fluorescent beads

to the cell suspensions (Figure 2D). In contrast to the sharp

decline in CD4 T cells, the absolute numbers of CD8 T and

B cells were unaltered. Separating the HLAC into distinct

cell types revealed that cell death occurred in purified popula-

tions of CD4 T cells suggesting that other cell types did not

mediate the killing. (Figure S1 available online). In all in-

stances, CD4-specific killing was prevented by AMD3100

but not AZT. Importantly, the extent of CD4 T cell depletion

in the presence of AZT was similar to that observed when

no antiviral drugs were added (Figure 2C and Figure 1, re-

spectively). Together, these results suggest that indirect

killing is the predominant mechanism for CD4 T cell depletion

in HIV-infected HLACs.

HIV gp41-Mediated Fusion Is Necessary for Depletionof Nonproductively Infected CD4 T CellsStudies with AMD3100 and AZT indicated that indirect CD4 T cell

killing is mediated by events occurring between gp120-CXCR4

binding and reverse transcription. Engagement of the chemokine

coreceptor induces conformational changes in gp41, resulting in

insertion of viral fusion peptide on gp41 into the target T cell

membrane. To determine if the gp120-CXCR4 interaction alone

or later events involving viral fusion are required for indirect

killing, we evaluated the effects of enfuvirtide (T20), a fusion

inhibitor that blocks six-helix bundle formation by gp41, a prereq-

uisite for virion fusion and core insertion.

We first determined the optimal concentrations of T20 that

block viral infection (Figure 3A). In NL4-3-infected cells, T20 began

to inhibit infection at concentrations > 2 mg/ml; complete inhibition

required 10 mg/ml. In cells infected with a primary viral isolate,

WEAU 16-8 (Figure S2), infection was completely inhibited by

0.1 mg/ml of T20. T20 did not inhibit infection with a T20-resistant

mutant, SIM (Rimsky et al., 1998), regardless of concentration.

Next, we investigated the effect of T20 on indirect CD4 T cell

killing (Figure 3B). In the absence of T20, high levels of indirect

killing were observed. T20 concentrations that blocked infection

also greatly inhibited indirect killing. T20 did not inhibit indirect

killing in cultures containing SIM-infected cells. Thus, blocking

gp41-mediated fusion prevents indirect killing.

Figure 3. HIV-1 Fusion Is Necessary to Induce Killing of Nonproductively Infected Cells

(A and C) Concentrations of T20 that block viral infection. HLACs were infected with the indicated clones of HIV-1 in the presence of the indicated concentrations

of T20 or no drugs. One hour before incubation with the virus, cells were pretreated with T20 or left untreated. At 12 hr, cells were washed extensively and cultured

under the same conditions. On day 9, the viral concentration was determined using a p24gag FLAQ assay. The amount of p24gag accumulated in the absence of

drugs by each viral clone (A) or by SKY (C) was defined as 100%.

(B and D) Effect of T20 on indirect killing. CFSE-labeled cells were cocultured with cells infected with the indicated viral clones in the presence of 5 mM AZT and the

indicated concentrations of T20. After 6 days, indirect killing in the mixed cultures was assessed. The number of viable CFSE-positive CD4 T cells cocultured with

uninfected cells in the presence of AZT was defined as 100% (data not shown). To allow successful initial infection we pseudotyped the GIA-SKY mutants with the

VSV-G envelope. NL4-3, WT lab-adapted virus; WEAU 16-8, primary virus; SIM, T20-resistant virus; GIA-SKY, T20-dependent virus; GIA and SKY, single-domain

mutant viruses. Representative data from three independent experiments with different donors are shown.

See also Figure S2.


We then examined a T20-dependent mutant, GIA-SKY (Bald-

win et al., 2004), which fuses only when T20 is present, but

cannot initiate a spreading infection in the absence of T20 (Fig-

ure 3C). Consistent with its T20 dependency, in the presence

of 1 mg/ml T20, the GIA-SKY mutant readily replicated while

growth was inhibited at higher or lower T20 concentrations.

The single-domain mutants GIA and SKY exhibited a T20-resis-

tance phenotype similar to that of SIM.

GIA-SKY-infected cells did not induce indirect killing of CD4

Tcells in theabsenceofT20 (Figure3D). Indirectkillingwasobserved

in cultures treated with 1 mg/ml T20 but was inhibited at higher or

lower concentrations. Since T20-dependent viruses were bound

to CXCR4 before T20 was added, these findings argue that

CXCR4 signaling is not sufficient to elicit indirect CD4 T cell killing.

Indirect Killing Requires a Close Interaction betweenUninfected and HIV-Infected CellsNext we examined whether indirect killing requires close contact

with HIV-infected cells or instead can be fully supported by

virions accumulating in the supernatants of the infected histocul-

tures. We found that cell-free supernatants from HIV-infected

histocultures were much less efficient at inducing indirect killing

(Figure 4A). To exclude the possibility that the concentration of

virions in the supernatants was too low, we repeated this exper-

iment using a 20-fold concentrated virion supernatants (1 mg

p24/ml) but failed to detect indirect CD4 T cell killing (Figure 4B).

Together, these findings suggest that close cell-cell contact is

likely required for indirect killing.

To further explore the potential requirement of close cell-cell

contact for indirect killing (Sherer et al., 2007; Sourisseau et al.,

2007), we repeated these assays using cells that had been washed

daily with fresh RPMI to prevent accumulation of HIV-1 virions and

soluble factors. Such cell washing did not affect the ability of the

resultant infected cells to mediate indirect CD4 T cell killing (Fig-

ure 4B), suggesting that virions released into the medium do not

participate in indirect killing. We confirmed these findings using

a transwell culture system. CSFE-labeled cells and HIV-infected

cells were mixed or physically separated by a transwell insert with

1 mm pores, which allows free diffusion of virions but not cells. Indi-

rect killing was substantial in the mixed cultures but not in the trans-

well cultures (Figure 4C). Together, these findings indicate that

indirect killing requires close interaction between CFSE-labeled

and HIV-1-infected cells, consistent with in vitro (Garg et al.,

2007; Holm and Gabuzda, 2005) and in vivo studies showing that

apoptotic nonproductively infected cells in human lymph nodes

often cluster near productively infected cells (Finkel et al., 1995).

Indirect Killing Requires Fusion of Virionsfrom Nearby HIV-Producing CellsIndirect killing required gp41-mediated fusion and close interac-

tion with HIV-infected cells, suggesting that cell death may be

caused by the fusion of HIV-1 virions to CD4 T cells, syncytia

formation, or hemifusion (mixing of lipids in the absence of fusion

pore formation) mediated by Env present on HIV-infected cells

interacting with neighboring CD4 T cells. HIV-1 virions (Holm

et al., 2004; Jekle et al., 2003; Vlahakis et al., 2001), cell-medi-

ated fusion (LaBonte et al., 2000; Margolis et al., 1995), and

hemifusion (Garg et al., 2007) have been proposed to be involved

in indirect killing. Therefore, the requirement for cell-cell interac-

tion in indirect killing may be mediated either by effective delivery

of HIV-1 virions or by cell-associated Env.

To discriminate between virion-mediated and cell-associated

Env induction of indirect killing, we tested the effects of HIV

protease inhibitors. These inhibitors act during the budding

process, resulting in immature viral particles that cannot fuse

with target cells (Wyma et al., 2004). We first assessed the effect

of protease inhibitors on viral maturation. NL4-3 viruses carrying

a b-lactamase-Vpr (BlaM-Vpr) reporter protein were produced

in 293T cells in the presence or absence of the HIV protease

inhibitor amprenavir. We also produced a mutant virus, TR712,

encoding a form of gp41 lacking 144 of the 150 amino acids in

the C-terminal cytoplasmic tail. This deletion largely relieves

the impaired fusogenic properties of immature HIV-1 particles

(Wyma et al., 2004). Protein analysis of viral lysates showed

that the NL4-3 and TR712 virions appropriately cleaved gp160

to generate gp120 in the presence and absence of amprenavir.

However, in the presence of amprenavir, an uncleaved form of

p55 Gag polypeptide rather than the mature p24 CA protein

accumulated in both NL4-3 and TR712 virions (Figure 4D). These

results confirm that amprenavir treatment of virus producing

cells results in the accumulation of immature particles containing

normal levels of incorporated Env proteins.

To test the ability of these viruses to fuse with target cells, we

used an HIV virion-based fusion assay that measures b-lacta-

mase (BlaM) activity delivered to target cells upon the fusion

of virions containing BlaM fused to the Vpr protein (BlaM-Vpr)

(Cavrois et al., 2002). Immunoblotting for BlaM confirmed that

NL4-3 and TR712 virions incorporated Blam-Vpr in the presence

or absence of amprenavir (Figure 4D).

Next, SupT1 cells were infected with mature or amprenavir-

treated immature NL4-3 or TR712 virions containing BlaM-Vpr.

Immature NL4-3 viruses displayed a 90% decline in fusogenic

properties (Figure 4E). In contrast, immature TR712 retained

40% fusion capacity, indicating that the impaired fusion is not

a result of a defective BlaM enzyme. Thus, immature virions

generated in the presence of amprenavir display greatly reduced

ability to fuse with target cells. Importantly, protease inhibitors

did not affect the function of Env proteins expressed on infected

cells and did not block cell-cell fusion (Figure S3C).

We next investigated the effect of protease inhibitors on

indirect killing. Remarkably, three different protease inhibitors

inhibited indirect killing as efficiently as AMD3100 (Figure 4F).

These results indicated that HIV-1 virions, not HIV-infected cells,

are responsible for indirect CD4 T cell killing. Additionally, reca-

pitulating the efficient viral delivery of close cell-cell interactions

by spinoculation of free virions resulted in extensive and selec-

tive indirect killing of CD4 T cells while sparing CD8 T cells and

B cells (Figures S3A and S3B). Thus, although indirect killing in

lymphoid cultures requires a close interaction between nonpro-

ductively and productively infected cells, this killing involves

virions rather than cell-associated Env.

Nonpermissive CD4 T Cells Die from Abortive InfectionBased on these findings, we hypothesized that ‘‘indirect killing’’

involves an abortive form of infection, like that which occurs in

nonpermissive resting CD4 T cells. These naive CD4 T cells


exhibit an early post-entry block to HIV-1 infection that can be

relieved by activation with phytohemagglutinin (PHA) and inter-

leukin-2 (IL-2) (Kreisberg et al., 2006; Santoni de Sio and Trono,

2009; Unutmaz et al., 1999; Zack et al., 1990). To test this

hypothesis, we compared the killing of activated and nonacti-

vated CFSE-labeled cells in HLACs.

Figure 4. Killing of Nonproductively Infected CD4 T Cells Requires Fusion of Virions from Nearby HIV-1-Producing Cells

(A) Supernatants from HIV-infected HLACs are less efficient at inducing indirect killing than mixing of HIV-infected and uninfected HLACs.

(B) HIV-1 virions released into the medium do not participate in indirect killing. Replacing the mixed culture with fresh RPMI every 24 hr did not impair indirect killing.

Challenging HLACs with supernatants containing 20-fold more histoculture-derived virions (1 mg p24/ml) than normally accumulated in mixed cultures containing

infected cells (50 ng p24/ml) did not induce indirect killing. Percentages are normalized to the number of viable CFSE-positive cells depicted by an asterisk.

(C) CFSE-labeled cells are not killed when HIV-infected HLAC is physically separated by a 1 mm –pore transwell system that allows free diffusion of HIV-1 parti-

cles. Values represent the levels of viable CFSE-positive cells after 6 days of culture in the presence of the indicated drugs. Red, HIV-infected cells; blue, unin-

fected cells; green, CFSE-labeled cells.

(D) Mature and immature viruses carry equivalent amounts of envelope protein and Blam-Vpr, but differ in their content of capsid and Gag precursor. NL4-3 and

TR712 viruses were generated in 293T cells with or without amprenavir, lysed and subjected to SDS-PAGE immunoblotting analysis for gp120, p55 Gag, p24 CA,

Blam-Vpr, and free Blam.

(E) Immature viruses have reduced capacity to enter cells. SupT1 cells were mock infected or infected with mature or immature NL4-3 or TR712 virions con-

taining Blam-Vpr. After loading of cells with CCF2 dye, fusion was analyzed by flow cytometry. Percentages are the fraction of cells displaying increased cleaved

CCF2 fluorescence, indicating virion fusion.

(F) Protease inhibitors inhibit indirect killing. CFSE-labeled cells were cocultured with NL4-3-infected or uninfected cells in the presence of AZT (5 mM) alone or

together with AMD3100 (250 nM). To the indicated cultures were added 5 mM of amprenavir, saquinavir, or indinavir. Percentages are normalized to the number

of viable CFSE-positive cells depicted by an asterisk. Error bars represent the SD obtained with three independent samples from the same donor.

See also Figure S3.


CFSE-labeled cells were activated with PHA and IL-2 two days

before mixing with effector cells, and contained a large

percentage of dividing CD25 and CD69 positive cells. Nonacti-

vated (resting) CFSE-labeled cells did not divide and typically

contained a small percentage of cells expressing CD25 and

CD69 (Figure 5A). Either in the presence or absence of AZT,

killing of resting CFSE-labeled CD4 T cells was robust (Figure 5B,

columns 4 and 5, and 16 and 17). In sharp contrast, activated

Figure 5. Death of Abortively Infected CD4 T Cells Is Due to Impaired Reverse Transcription

(A) Status of mixed HLACs containing either resting or activated CFSE-labeled cells, 4 days after coculturing with effector cells. Activated CFSE-labeled cells

were stimulated with PHA and IL-2 48 hr before mixing, but not during coculturing with effector cells. To avoid direct killing of activated CFSE-labeled cells in

cultures with no drugs, cell killing was terminated and analyzed 4 days after coculturing.

(B) AZT renders activated CFSE-labeled CD4 T cells sensitive to indirect killing. Resting or activated CFSE-labeled cells were cocultured with effector cells in the

presence of no drugs, AZT (5 mM) alone, or AZT and AMD3100 (250 nM). Data are from two independent experiments performed with tonsil cells from two different

donors.

(C) AZT-induced killing is lost when AZT-resistant viruses are tested. Resting or activated CFSE-labeled cells were cocultured with cells infected with NL4-3 or

HIV-1 clones #629 and #964 in the presence of no drugs, AZT (0.5 mM) alone, or AZT and AMD3100 (250 nM). AZT-sensitive and AZT-resistant sub-clones are

depicted. Data are representative of three independent experiments with three different donors.

(D) NNRTIs prevent killing of abortive infected CD4 T cells. Resting or activated CFSE-labeled cells were cocultured with infected or uninfected effector cells, in

the presence of no drugs, AZT (5 mM), AMD3100 (250 nM), the NNRTIs efavirenz (100 nM), and nevirapine (1 mM), or the integration inhibitors raltegravir (30 mM)

and 118-D-24 (60 mM). Killing of resting CFSE-labeled CD4 T cells was blocked with equal efficiency by NNRTIs and AMD3100 (columns 15, 16), but not by

integration inhibitors (columns 17, 18). In combination, NNRTIs prevented cell death induced by AZT in activated CFSE-labeled cells (compare column 38 to

44 and 45). Data are representative of four independent experiments with four different donors.

The absolute numbers of CFSE-labeled CD8 T cells and B cells was unaltered in these experiments (data not shown). Percentages are normalized to the number

of viable CFSE-positive cells depicted by an asterisk.

See also Figure S4.


Figure 6. Cytoplasmic HIV-1 DNA Triggers Proapoptotic and Proinflammatory Responses in Abortively Infected CD4 T Cells

(A) Critical reactions in HIV-1 reverse transcription as detected by probes monitoring different regions within the Strong stop, Nef, and Env DNA fragments. RDDP,

RNA-dependent DNA polymerase. Adapted from S.J. Flint et al., Principles of Virology, 2000 ASM Press, Washington DC, with permission.

(B) NNRTIs prevent accumulation of DNA elongation products. The amount of viral DNA detected by a particular probe was calculated as a fold change relative

to cells treated with no drugs (i.e., calibrator). A b�actin probe was used as an internal reference. Mean cycle threshold (Ct) values of calibrator samples are

depicted. CD4 T cells were infected with WT NL4-3 produced in 293T cells, or with a Dvif NL4-3 collected from supernatants of infected HLAC, as described

in Figure S4C. Data are representative of two independent experiments performed with cells from two different donors.

(C and D) Abortive HIV-1 infection generates a coordinated proapoptotic and proinflammatory response involving caspase-3 and caspase �1 activation. HLACs

were spinoculated with no virus or with NL4-3 and AZT (5 mM), Efavirenz (100 nM), and T20 (10 mg/ml), as indicated (see Figures S3A and S3B). After 3 days, cells

were assessed by flow cytometry for intracellular levels of proinflammatory cytokines, serine 37 phosporylated p53, and activated caspases as indicated.

Ethidium monoazide was used to exclude dead and necrotic cells from the annexinV binding analysis. Data are representative of three independent experiments

with three different donors.

(E) Death of abortively infected CD4 T cells requires caspase activation. CSFE-labeled cells were cocultured with effector cells in the presence of 20 mM of Z-VAD-

FMK, a general caspase inhibitor,or Z-FA-FMK, a negativecontrol for caspase inhibitors.AZT (5mM); AMD3100 (250 nM).Percentages are normalized to the number

of viable CFSE-positive cells depicted by an asterisk. Error bars represent standard error of the mean of three experiments from three different HLAC donors.

(F) Abortive HIV infection promotes the maturation and secretion of IL-1b in tonsillar CD4 T cells. Isolated tonsillar CD4 T cells were either untreated, or stimulated

with PMA (phorbol-12-myristate-12-acetate, 0.5 mM) and the potassium ionophore nigericin (10 mM), or spinoculated with or without NL4-3 in the presence of


CFSE-labeled CD4 T cells were not depleted in the absence of

AZT, but were extensively depleted in cultures containing AZT

(Figure 5B, columns 10 and 11 and 22 and 23). Addition of

AMD3100 prevented the AZT-induced killing of activated

CFSE-labeled cells, excluding nonspecific toxic effects of AZT

in the activated cells (Figure 5B, columns 12 and 24).

The ability of AZT to promote indirect killing of activated CD4

T cells suggested that cell death is triggered by impaired reverse

transcription. To investigate this possibility, we repeated the

experiment with two pairs of AZT-resistant HIV-1 clones, 629

and 964 (Larder et al., 1989). We first determined that concentra-

tions of 0.5 mM AZT block viral replication in NL4-3-infected and

AZT-sensitive clones and achieve half maximal inhibitory effect

in AZT-resistant clones (Figures S4A and S4B).

When resting CFSE-labeled cells were used, the extent of

killing by the AZT-resistant HIV-1 viruses was similar to that

obtained with NL4-3 with or without AZT (Figure 5C resting

CFSE-positive cells), demonstrating a redundant function for

endogenous termination of reverse transcription and AZT. Alter-

natively, when activated CFSE-labeled cells were tested, AZT-

resistant HIV-1 clones did not deplete CFSE-labeled CD4

T cells in the presence of AZT (Figure 5C, columns 29 and 35).

Death of Abortively Infected CD4 T Cells Is Triggeredby Premature Termination of Viral DNA ElongationWe next asked what stage of reverse transcription triggers abor-

tive infection cell death. AZT inhibits DNA elongation but not

early DNA synthesis (Arts and Wainberg, 1994). We therefore

examined whether blocking early DNA synthesis with nonnu-

cleoside reverse transcriptase inhibitors (NNRTIs) would have

the same effect as AZT. Impaired reverse transcription may

also lead to abortive integration, causing chromosomal DNA

breaks and a genotoxic response. To exclude this possibility,

we used integrase inhibitors. To discriminate between the cyto-

pathic response induced by endogenous termination of reverse

transcription and the response induced by AZT, we separately

assessed resting and activated CFSE-labeled cells.

Remarkably, the NNRTIs, efavirenz and nevirapine, blocked

indirect killing of resting CD4 T cells as efficiently as AMD3100

(Figure 5D, columns 15 and 16). These findings suggested that

allosteric inhibition of reverse transcriptase induced by these

NNRTI’s interrupts reverse transcription sufficiently early to

abrogate the death response. In contrast, the integrase inhibitors

raltegravir and 118-D-24 did not prevent abortive infection killing

(Figure 5D, columns 17 and 18), suggesting that cell death

involves signals generated prior to viral integration. NNRTIs

also protected activated CFSE-labeled cells from death induced

by AZT (Figure 5D, column 38 versus columns 44 and 45),

demonstrating that a certain degree of DNA synthesis is required

to elicit the cytopathic response.

This notion was further strengthened in findings obtained with

vif-deficient (Dvif) HIV-1 particles where reverse transcription is

inhibited during strong-stop DNA synthesis due to incorporated

APOBEC3G (A3G) (Bishop et al., 2008; Li et al., 2007). Abortively

infected CD4 T cells were not depleted by Dvif NL4-3-infected

cells (Figures S4C and S4D), indicating that termination of

reverse transcription before the completion of strong-stop

DNA synthesis is not sufficient to generate a cytopathic

response. Other HIV-1 mutants containing substitutions in

RNase H and nucleocapsid that promote early defects in reverse

transcription failed to elicit indirect CD4 T cell killing (Figures S4E

and S4F). Together, these findings indicate that accumulation of

reverse-transcribed DNA, rather than any inherent activity of the

HIV-1 proteins, is the key factor that triggers the death response.

Abortively Infected CD4 T Cells Commencebut Do Not Complete Reverse TranscriptionWe next examined the status of HIV-1 reverse transcription in

tonsillar CD4 T cells after infection. Specifically, we investigated

the effect on reverse transcription after treatment with NNRTIs,

such as efavirenz and nevirapine, which prevent the death of

abortively infected CD4 T cells, or with AZT or integrase inhibitor

(raltegravir) that do not prevent CD4 T cell death. Taqman-based

quantitative real-time PCR (QPCR) was used to quantify the

synthesis of reverse transcription products in isolated CD4

T cells from HLAC 16 hr after infection with NL4-3. We designed

specific QPCR primers and probes (Table S1) to monitor

sequential steps in reverse transcription including generation

of strong-stop DNA, first template exchange (Nef), and DNA

strand elongation (Env) (Figure 6A). Reverse transcription

products corresponding to strong-stop DNA were similar in

untreated CD4 T cells or cells treated with AZT, NNRTIs, or

raltegravir but were greatly reduced in cells treated with

AMD3100 or in cultures infected with Dvif NL4-3 where arrest

occurs prior to the completion of strong-stop DNA synthesis

(Figure 6B columns 1–8). In contrast, the accumulation of later

reverse transcription products detected by the Nef and Env

probes were dramatically inhibited by the NNRTIs but not by

raltegravir. Levels of Nef (Figure 6B, columns 10 and 11) and

Env (columns 18 and 19) DNA products were similar in untreated

cells and cells treated with AZT, indicating that reverse transcrip-

tion in most tonsillar CD4 T cells naturally terminates during DNA

chain elongation, coinciding with the block induced by AZT. The

minor inhibition detected by AZT is likely due to a small number

of permissive CD4 T cells in the culture. These results show that

abortively infected CD4 T cells accumulate incomplete reverse

AZT (5 mM), AMD3100 (250 nM), and efavirenz (100 nM) as indicated. After 3 days, half of the cells were lysed and subjected to SDS-PAGE immunobloting anal-

ysis. On day 5, the supernatants from the rest of the cells were collected and subjected to SDS-PAGE immunobloting analysis. The IL-1b antibody detects the

pro-IL-1b (37kD) and the mature cleaved form (17kD). Data are the representative results of five independent experiments using tonsillar CD4 T cells isolated

from five different donors.

(G) DNA reverse transcription intermediates induce an IFN-stimulatory antiviral innate immune response (ISD). ISRE-GFP reporters were transfected with 1 mg of

HIV-1 reverse transcription intermediate products as indicated by numbers (detailed description in Figure S5E), empty DNA plasmid, or polyinosinic:polycytidylic

acid [poly(I:C)], and were analyzed by flow cytometry after 48 hr. Data are representative of three independent experiments; error bars show the SD for three

independent samples from the same experiment.



transcription products representative of DNA strand elongation.

Blocking earlier steps of reverse transcription by NNRTIs or by

genetic mutations like deletion of Vif or mutation of RNase H

restricts accumulation of such products, and prevents abortive

infection-induced cell death (Figure S6A).

DNA Reverse Transcription Intermediates Elicita Coordinated Proapoptotic and ProinflammatoryResponse in Abortively Infected CD4 T CellsWe next evaluated whether HIV-mediated indirect killing of CD4

T cells is associated with deregulation of cytokine production or

a DNA damage response. To facilitate a vigorous and synchro-

nized killing effect, HLACs were spinoculated with NL4-3 virions

in the presence of various antiviral drugs. Interestingly, based on

immunostaining after cytokine capture, abortively infected CD4

T cells expressed IFN-b, and high levels of the proinflammatory

interleukin 1b (IL–1b), but not tumor necrosis factor (TNFa)

(Figure 6C). Phosphorylation of S37 p53 was not observed,

suggesting that abortive HIV-1 infection does not induce a

DNA damage cascade. Abortively infected CD4 T cells also

displayed caspase-1 and caspase-3 activity along with appear-

ance of annexin V (Figure 6D). T20 and efavirenz but not AZT

prevented activation of these caspases, indicating that apo-

ptosis was induced by abortive HIV-1 infection. Cell death was

completely prevented by Z-VAD-FMK, a pan-caspase inhibitor,

suggesting that caspase activation is required for the observed

cytopathic response (Figure 6E). Such mode of cytokine produc-

tion and caspase activation was not observed in CD8 T or B cells

(Figures S5B and S5C).

We next examined whether abortive HIV-1 infection signals for

the maturation and secretion of IL-1b. In cells, IL–1b activity is

rigorously controlled. Cells can be primed to express inactive

pro-IL-1b by various proinflammatory signals. However, the

release of bioactive IL-1b requires a second signal leading to

activation of inflammasomes, cleavage of pro-IL-1b by caspase

1 and secretion of the bioactive 17 kDa form of IL-1b (Schroder

and Tschopp, 2010). Interestingly, Western blot analysis re-

vealed high amounts of intracellular pro-IL-1b in untreated CD4

T cells, suggesting that tonsillar CD4 T cells are primed to release

proinflammatory mediators (Figure 6F). Stimulating the CD4

T cells with PMA and nigericin induced further accumulation

of pro-IL-1b and promoted the maturation and release of the

bioactive 17 kDa IL-1b into the supernatant. Remarkably, infec-

tion of CD4 T cells with NL4-3 in the presence of AZT similarly

resulted in maturation and release of the bioactive 17 kDa

IL-1b into the supernatant. This response was completely pre-

vented by efavirenz and AMD3100, suggesting that abortive

HIV-1 infection signals the maturation and release of bioactive

IL-1b in these CD4 T cells.

To identify the nature of the nucleic acid species that trigger

these responses, we used a recently described H35 rat hepato-

cyte cell line containing an IFN-sensitive response element

(ISRE) linked to GFP (Patel et al., 2009). H35 cells were first

infected with pseudotyped VSV-G HIV-1 virions. These virions

induced GFP expression and cell death in the presence or

absence of AZT. Importantly, the expression of GFP and cell

death response were blocked by efavirenz but not raltegravir

(Figure S5D). Thus, the H35 system successfully reconstitutes

the cytokine and cytopathic response observed in tonsillar

CD4 T cells. We next synthesized the various HIV-1 reverse

transcription intermediates and tested their ability to activate

the ISRE-GFP reporter. Interestingly, none of the RNA-contain-

ing oligonucleotides stimulated the ISRE-GFP reporter expres-

sion above baseline. In sharp contrast, ssDNA and dsDNA

oligonucleotides longer than 500 bases in length, which corre-

sponded to reverse transcription intermediates produced during

DNA elongation, evoked a potent ISRE-GFP activation (Fig-

ure 6G). Similarly, when cells were stimulated with poly(I:C), a

synthetic double-stranded RNA known to activate IRF3 via the

RIG-I pathway elicited a comparable ISRE-GFP response. Taken

together, these findings indicate that reverse transcription

intermediates generated during DNA chain elongation induce

a coordinated proapoptotic and proinflamatory innate immune

response involving caspase-3 and caspase-1 activation in

abortively infected CD4 T cells.

DISCUSSION

The mechanism through which HIV-1 kills CD4 T cells, a hallmark

of AIDS, has been a topic of vigorous research and one of the

most pressing questions for the field over the last 28 years

(Thomas, 2009). In this study, we investigated the mechanism

of HIV-1-mediated killing in lymphoid tissues, which carry the

highest viral burdens in infected patients. We used HLACs

formed with fresh human tonsil cells and an experimental strategy

that clearly distinguishes between direct and indirect mecha-

nisms of CD4 T cell depletion. We now demonstrate that indirect

cell killing involving abortive HIV infection of CD4 T cells accounts

for the vast majority of cell death occurring in lymphoid tissues.

No more than 5% of the CD4 T cells are productively infected,

but virtually all the remaining CD4 T cells are abortively infected

ultimately leading to caspase-mediated cell death. Equivalent

findings were observed in HLACs formed with fresh human

spleen (Figures S6B and S6C), indicating this mechanism of

CD4 T cell depletion can be generalized to other lymphoid

tissues.

The massive depletion of nonproductively infected CD4 T cells

is in contrast to their survival after infection of intact blocks of

tonsillar tissue in human lymphoid histoculture (HLH) (Grivel

et al., 2003). This result probably reflects differences between

the HLH and the HLAC experimental systems. In HLH, the com-

plex three-dimensional spatial cellular organization of lymphoid

tissue is preserved, but cellular movement and interaction are

restricted, both of which are required for indirect killing. In

HLAC, the tissue is dispersed, and cells are free to interact, result-

ing in a rapid and robust viral spread. While the mechanism

triggering indirect CD4 T cell death is certainly identical in both

settings, HLH allows only a slow, nearly undetectable progres-

sion of indirect CD4 T cell death. In HLAC, this process is accel-

erated, allowing the outcome to be detected in a few days. Inter-

estingly, indirect killing was also less efficient when peripheral

blood cells were tested (data not shown). It is possible that cellular

factors specifically produced in lymphoid organs are required to

accelerate indirect killing of peripheral blood CD4 T cells.

Several mechanisms have been proposed to explain indirect

CD4 T cell killing during HIV infection. Our finding that CD4


T cell death is blocked by entry and fusion inhibitors but not by

AZT, strongly suggested that such killing involves nonproductive

infection of CD4 T cells. Therefore, we focused on events that

occur after HIV-1 entry. Our investigations demonstrate that

abortive viral DNA synthesis occurring in nonpermissive, quies-

cent CD4 tonsil T cells, plays a key role in the cell death

response. Conversely, in the small subset of permissive target

cells, reverse transcription is not interrupted, minimizing the

accumulation and subsequent detection of such reverse

transcription intermediates (Figure 7).

Interrupted or slowed reverse transcription may create persis-

tent exposure to cytoplasmic DNA products that elicit an antiviral

innate immune response coordinated by activation of type I IFNs

(Stetson and Medzhitov, 2006). Such activation, termed

IFN-stimulatory DNA (ISD) response, may be analogous to the

type I IFN response triggered by the RIG-I-like receptor (RLR)

family of RNA helicases that mediate a cell-intrinsic antiviral

defense (Rehwinkel and Reis e Sousa, 2010). Our results suggest

that abortive HIV-1 infection also stimulates activation of

caspase-3, which is linked to apoptosis, and caspase-1, which

promotes the processing and secretion of the proinflammatory

cytokines like IL–1b. It is certainly possible that pyroptosis

elicited in response to caspase-1 activation also contributes to

the observed cytopathic response (Schroder and Tschopp,

2010). The release of inflammatory cytokines during CD4 T cell

death could also contribute to the state of chronic inflammation

that characterizes HIV infection. This inflammation may fuel

further viral spread by recruiting uninfected lymphocytes to the

inflamed zone. While this innate response was likely designed

to protect the host, it is subverted in the case of HIV infection

and importantly contributes to the immunopathogenic effects

characteristic of HIV infection and AIDS.

Such antiviral pathways comprise an unrecognized cell-

intrinsic retroviral detection system (Manel et al., 2010; Stetson

Figure 7. Consequences of Inhibiting Early Steps of HIV-1 Infection on CD4 T Cell Death

(A) The nonpermissive state of most CD4 T cells in lymphoid tissue leads to endogenous termination of reverse transcription during DNA chain elongation (i.e.,

‘‘killing zone’’). As a result, DNA intermediates accumulate in the cytoplasm and elicit a multifaceted proapoptotic and proinflammatory innate immune defense

programs, coordinated by IFN-stimulatory DNA (ISD) response, caspase-3, caspase-1, and IL-1b, to restrict viral spread. Different classes of antiretroviral drugs

act at different stage of the HIV life cycle. NNRTIs like efavirenz and nevirapine inhibit early steps of DNA synthesis and therefore prevent such response and the

consequence CD4 T cell death. AZT is less efficient at blocking DNA synthesis and therefore unable to abrogate this response.

(B) In permissive CD4 T cells reverse transcription proceeds, allowing HIV-1 to bypass the ‘‘killing zone’’ and move on to productive (or latent) infection.

Interrupting reverse transcription with AZT traps the virus in the ‘‘killing zone’’ and induces cell death. EFV, efavirenz; NVP, nevirapine; and RTGR, raltegravir.

See also Figure S6.


et al., 2008). Viral RNA in infected cells is recognized by

members of the RIG-I-like family of receptors that detect specific

RNA patterns like uncapped 50 triphosphate (Rehwinkel and

Reis e Sousa, 2010). Although uncapped RNA intermediates

are generated by the HIV-1 RNase H, they contain a 50 mono-

phosphate and therefore may be not recognized by the RIG-I

system (Figure 6G). In contrast to RNA receptors, intracellular

sensing of viral DNA remains poorly understood. Consequently,

it is unclear how HIV-1 DNA intermediates are detected in the

cytoplasm of abortively infected CD4 T cells. AIM2 (absent in

melanoma 2) was recently identified as a cytoplasmic dsDNA

receptor that induces cell death in macrophages through activa-

tion of caspase-1 in imflammasomes (Hornung et al., 2009). Our

preliminary investigations have not supported a role for AIM2 in

cell death induced by abortive HIV infection (data not shown),

suggesting the potential involvement of a different DNA-sensing

mechanism. We also have not identified a role for TLR9 and

MYD88 signaling in this form of cell death. Additional candidate

sensors recognizing cytoplasmic HIV-1 DNA are now under

study.

In summary, both productive and nonproductive forms of HIV

infection contribute to the pathogenic effects of this lentivirus.

The relative importance of these different cell death pathways

might well vary with the stage of HIV infection. For example,

direct infection and death might predominate during acute

infection where CCR5-expressing memory CD4 T cells in gut-

associated lymphoid tissue are effectively depleted. Conversely,

the CXCR4-dependent indirect killing we describe in tonsil tissue

may reflect later stages of HIV-induced disease where a switch

to CXCR4 coreceptor usage occurs in approximately 50% of

infected subjects.

The current study demonstrates how a cytopathic response

involving abortive viral infection of resting nonpermissive CD4

T cells can lead not only to CD4 T cell depletion but also to the

release of proinflammatory cytokines. The ensuing recruitment

of new target cells to the site of inflammation may fuel a vicious

cycle of continuing infection and CD4 T cell death centrally

contributing to HIV pathogenesis.


Culture and Infection of HLACs

Human tonsil or splenic tissues were obtained from the National Disease

Research Interchange and the Cooperative Human Tissue Network and pro-

cessed as previously described (Jekle et al., 2003). For a detailed description

see Extended Experimental Procedures.

FACS Analysis and Gating Strategy, Preparation of HIV-1 Virions,

and Virion-Based Fusion Assay

Data were collected on a FACS Calibur (BD Biosciences) and analyzed with

Flowjo software (Treestar). HIV-1 viruses were generated by transfection of

proviral DNA into 293T cells by the calcium phosphate method. Virion-based

fusion assay was performed as previously described (Cavrois et al., 2002).

Detailed protocols are provided in the supplemental experimental procedures.

Spinoculation and Taqman-Based QPCR Analysis

of HIV-1-Infected CD4 T Cells

The spinoculation method is described in detail in Figures S3A and S3B.

Isolation of HLAC CD4 T cells and QPCR protocol are described in detail

in supplemental experimental procedures. Primers and probes sequences

used to detect reverse transcription products are provided in Table S1.

QPCR reactions were performed in an ABI Prism 7900HT (Applied

Biosystems).

ISRE-GFP H35 Reporter Cells, Microscopy, and Generation

of Synthetic HIV-1 Reverse Transcription Intermediates

H35 rat hepatic cells containing an ISRE-GFP reporter were maintained as

described (Patel et al., 2009). For microscopy imaging, ISRE-GFP reporter

H35 cells were infected with a replication competent VSV-G pseudotyped

NL4-3 and analyzed using an Axio observer Z1 microscope (Zeiss). Transfec-

tions and generation of synthetic HIV-1 reverse transcription intermediates are

described in detail in Figure S5E and Extended Experimental Procedures.



six figures, and one table and can be found with this article online at

doi:10.1016/j.cell.2010.11.001.

ACKNOWLEDGMENTS

We thank David N. Levy for the NLENG1 plasmid; David Fenard for the NL4-3

variant plasmids SIM, GIA, GIA-SKY, and SKY; George M. Shaw for the WEAU

16-8 env clone; and Suraj J. Patel, Kevin R. King, and Martin L. Yarmush for the

H35 ISRE-GFP reporter cell line. The following reagents were obtained through

the AIDS Research and Reference Reagent Program, Division of AIDS, NIAID,

NIH: AMD3100, T-20, Saquinavir, Amprenavir, Indinavir, Nevirapine, Efavirenz,

and AZT-resistant HIV-1 clones #629 and #964. Special thanks to Dr. Eva

Herker for assistance with fluorescence microscopy; to Dr. Stefanie Sowinski

for help with assessing inflammatory responses in primary immune cells; and

to Jason Neidleman for stimulating discussions and technical advice. We also

thank Marty Bigos for assistance with the flow cytometry; Stephen Ordway

and Gary Howard for editorial assistance; John C.W. Carroll and Alisha Wilson

for graphics; and Robin Givens and Sue Cammack for administrative assis-

tance. Funding for this project was provided by the Universitywide AIDS

Research Program, F04-GIVI-210 (G.D.); the UCSF-GIVI Center for AIDS

Research, NIH/NIAID P30 AI027763 (M.C.); the Francis Goelet Fellowship

(K.G.L.); and the UCSF Medical Scientist Training Program, NIH/NIGMS T32

GM007618-32 (O.Z.).

Received: November 5, 2009

Revised: May 7, 2010



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Sirt3 Mediates Reduction of OxidativeDamage and Prevention of Age-RelatedHearing Loss under Caloric RestrictionShinichi Someya,1,3,5 Wei Yu,2,5 William C. Hallows,2 Jinze Xu,4 James M. Vann,1 Christiaan Leeuwenburgh,4

Masaru Tanokura,3 John M. Denu,2,* and Tomas A. Prolla1,*1Departments of Genetics and Medical Genetics2Department of Biomolecular Chemistry

University of Wisconsin, Madison, WI 53706, USA3Department of Applied Biological Chemistry, University of Tokyo, Yayoi, Tokyo 113-8657, Japan4Department of Aging and Geriatrics and The Institute on Aging, University of Florida, Gainesville, FL 32611, USA5These authors contributed equally to this work*Correspondence: [email protected] (J.M.D.), [email protected] (T.A.P.)

DOI 10.1016/j.cell.2010.10.002

SUMMARY

Caloric restriction (CR) extends the life span andhealth span of a variety of species and slows theprogression of age-related hearing loss (AHL),a common age-related disorder associated withoxidative stress. Here, we report that CR reducesoxidative DNA damage in multiple tissues and pre-vents AHL in wild-type mice but fails to modifythese phenotypes in mice lacking the mitochondrialdeacetylase Sirt3, a member of the sirtuin family.In response to CR, Sirt3 directly deacetylates andactivates mitochondrial isocitrate dehydrogenase 2(Idh2), leading to increased NADPH levels and anincreased ratio of reduced-to-oxidized glutathionein mitochondria. In cultured cells, overexpressionof Sirt3 and/or Idh2 increases NADPH levels andprotects from oxidative stress-induced cell death.Therefore, our findings identify Sirt3 as an essentialplayer in enhancing the mitochondrial glutathioneantioxidant defense system during CR and suggestthat Sirt3-dependent mitochondrial adaptationsmay be a central mechanism of aging retardation inmammals.

INTRODUCTION

It is well established that reducing food consumption by 25%–

60% without malnutrition consistently extends both the mean

and maximum life span of rodents (Weindruch and Walford,

1988; Koubova and Guarente, 2003). Caloric restriction (CR) is

also known to extend life span in yeast, worms, fruit flies,

spiders, birds, and monkeys and delays the progression of

a variety of age-associated diseases such as cancer, diabetes,

cataract, and age-related hearing loss (AHL) in mammals (Wein-

druch and Walford, 1988; Sohal and Weindruch, 1996; Someya

et al., 2007; Colman et al., 2009). Furthermore, CR reduces neu-

rodegeneration in animal models of Parkinson’s disease (Matt-

son, 2000) as well as Alzheimer’s disease (Zhu et al., 1999).

The mitochondrial free radical theory of aging postulates that

aging results from accumulated oxidative damage caused by

reactive oxygen species (ROS), originating from the mitochon-

drial respiratory chain (Balaban et al., 2005). Consistent with

this hypothesis, mitochondria are a major source of ROS and

of ROS-induced oxidative damage, and mitochondrial function

declines during aging (Wallace, 2005). A large body of evidence

suggests that CR reduces the age-associated accumulation of

oxidatively damaged proteins, lipids, and DNA through reduction

of oxidative damage to these macromolecules and/or enhanced

antioxidant defenses to oxidative stress (Weindruch and Wal-

ford, 1988; Sohal and Weindruch, 1996; Masoro, 2000). Yet,

whether the anti-aging action of CR in mammals is a regulated

process and requires specific regulatory proteins such as sir-

tuins still remains unclear.

Sirtuins are NAD+-dependent protein deacetylases that regu-

late life span in lower organisms and have emerged as broad

regulators of cellular fate and mammalian physiology (Donmez

and Guarente, 2010; Finkel et al., 2009). A previous report has

shown that life span extension by CR in yeast requires Sir2,

a member of the sirtuin family (Lin et al., 2000), linking sirtuins

and CR-mediated retardation of aging. In mammals, there are

seven sirtuins that display diverse cellular localization (Donmez

and Guarente, 2010; Finkel et al., 2009). Previous studies have

focused on the role of Sirt1 as the major sirtuin mediating the

metabolic effects of CR in mammals (Chen et al., 2005; Bor-

done et al., 2007; Chen et al., 2008). However, recent studies

indicate that upregulation of Sirt1 in response to CR is not

observed in all tissues examined (Cohen et al., 2004; Barger

et al., 2008), and currently, no study has provided conclusive

evidence that sirtuins play an essential role in CR-mediated

aging retardation in mammals. Sirt3 is a member of the

mammalian sirtuin family that is localized to mitochondria and


regulates levels of ATP and the activity of complex I of the elec-

tron transport chain (Ahn et al., 2008) and, as such, may play

a role in the metabolic reprogramming mediated by CR. A

recent study has shown that CR increases Sirt3 levels in liver

mitochondria (Schwer et al., 2009). Fasting also increases

Sirt3 protein expression in liver mitochondria, and mice lacking

Sirt3 display the hallmarks of fatty acid oxidation disorders,

indicating that Sirt3 modulates mitochondrial fatty acid oxida-

tion in mammals (Hirschey et al., 2010). Furthermore, CR

increases expression of Sirt3 in primary mouse cardiomyo-

cytes, whereas overexpression of Sirt3 protects these cells

from oxidative stress-induced cell death (Sundaresan et al.,

2008), suggesting a potential role of Sirt3 in the aging retarda-

tion associated with CR in mammals.

AHL is a universal feature of mammalian aging and is the

most common sensory disorder in the elderly (Someya and

Prolla, 2010; Liu and Yan, 2007). AHL is characterized by an

age-associated decline of hearing function associated with

loss of spiral ganglion neurons and sensory hair cells in the

cochlea of the inner ear (Someya and Prolla, 2010; Liu and

Yan, 2007). The progressive loss of neurons and hair cells in

the inner ear leads to the onset of AHL because these postmi-

totic cells do not regenerate in mammals. The onset of AHL

begins in the high-frequency region and spreads toward the

low-frequency region during aging (Keithley et al., 2004; Hunter

and Willott, 1987). This is accompanied by the loss of neurons

and hair cells beginning in the basal region and spreading

toward the apex of the cochlea of the inner ear with age.

A previous study has shown that CR slows the progression of

AHL in CBA/J mice (Sweet et al., 1988), whereas we have

shown previously that CR prevents AHL in C57BL/6J mice,

reduces cochlear degeneration, and induces Sirt3 in the

cochlea (Someya et al., 2007). Both strains of mice have been

extensively used as a model of AHL, although the age of onset

of AHL varies from 12–15 months of age in C57BL/6J mice to

18–22 months of age in CBA/J mice (Zheng et al., 1999). Exper-

imental evidence suggests that oxidative stress plays a major

role in AHL (Jiang et al., 2007; Someya et al., 2009) and that

CR protects cochlear cells through reduction of oxidative

damage and/or by enhancing cellular antioxidant defenses to

oxidative stress (Someya et al., 2007). Yet, the molecular mech-

anisms by which CR reduces oxidative cochlear cell damage

remain unknown.

In this report, we show that the mitochondrial deacetylase

Sirt3 is required for the CR-mediated prevention of AHL in

mice. We also show that Sirt3 is required for the reduction of

oxidative damage in multiple tissues under CR conditions, as

evidenced by DNA damage levels. At the mechanistic level,

Sirt3 directly deacetylates isocitrate dehydrogenase 2 (Idh2),

an enzyme that converts NADP+ to NADPH in mitochondria.

In response to CR, Sirt3 stimulates Idh2 activity in mitochon-

dria, leading to increased levels of NADPH and an increased

ratio of reduced glutathione/oxidized glutathione, the major

redox couple in the cell. In cultured cells, overexpression of

Sirt3 and/or Idh2 increases NADPH levels and protects these

cells from oxidative stress. The data presented here provide

the first conclusive evidence that CR-mediated reduction of

oxidative damage and prevention of a common age-related

phenotype (AHL) require a member of the sirtuin family in

mammals.

RESULTS

Sirt3 Is Required for the CR-Mediated Preventionof Age-Related Cochlear Cell Death and Hearing LossFirst, to investigate whether Sirt3 plays a role in the CR preven-

tion of AHL, we conducted a 10 month CR dietary study using

WT and Sirt3�/� mice that have been backcrossed onto the

C57BL/6J background. The C57BL/6J strain is considered an

excellent model to study the anti-aging action of CR because

this mouse strain is the most widely used mouse model for the

study of aging and responds to CR with a robust extension of

life span (Weindruch and Walford, 1988) and prevention of AHL

(Someya et al., 2007). We reduced the calorie intake of WT and

Sirt3�/� mice to 75% (a 25% CR) of that fed to control diet

(CD) mice in early adulthood (2 months of age), and this dietary

regimen was maintained until 12 months of age. The auditory

brainstem response (ABR), a common electrophysiological test

of hearing function, was used to monitor the progression of

AHL in these mice (Someya et al., 2009). We first confirmed

that aging resulted in increased ABR hearing thresholds at the

high (32 kHz), middle (16 kHz), and low (8 kHz) frequencies in

12-month-old WT mice (Figure 1A), indicating that these mice

displayed hearing loss. As predicted, CR delayed the progres-

sion of AHL at all tested frequencies in WT mice (Figure 1A).

Strikingly, CR did not delay the progression of AHL in Sirt3�/�

mice (Figure 1A), although CR had the same effect on body

weight reduction in both WT and Sirt3�/� mice (Figures S2A

and S2B available online). Neural and hair cell degeneration

are hallmarks of AHL (Keithley et al., 2004). In agreement with

the hearing test results, basal regions of the cochleae from

calorie-restricted WT mice displayed only minor loss of spiral

ganglion neurons (Figures 1J and 1K; see also Figures 1B, 1C,

1F and 1G) and hair cells (Figure S1E; see also Figures S1A

and S1C), whereas CR failed to protect these cells in Sirt3�/�

mice (Figures 1L and 1M; see also Figures 1D, 1E, 1H, and 1I;

Figure S1F; see also Figures S1B and S1D). Collectively, these

results demonstrate that Sirt3 plays an essential role in the CR-

mediated prevention of age-related cochlear cell death and

hearing loss in mice.

Next, to investigate whether Sirt3 plays a role in the metabolic

effects induced by CR, we conducted a 3 month CR dietary

study using WT and Sirt3�/� mice starting at 2 months of age.

Mice lacking the Sirt3 gene appeared phenotypically normal

under basal and CR conditions: Sirt3�/� mice were viable

and fertile, and no significant changes were observed in

body weight (Figures S2A and S2B), bone mineral density (Fig-

ure S2C), body fat (Figure S2D), tissue weight (Figure S2E),

serum glucose levels (Figure S3A), glucose tolerance (Fig-

ure S3B), serum Igf-1 (Figure S3C), and cholesterol (Figure S3D)

levels between control diet WT and Sirt3�/� mice or calorie-

restricted WT and Sirt3�/� mice at 5 months of age. However,

though we found that WT mice displayed lower levels of serum

insulin (Figure S3E) and triglycerides (Figure S3F) in response

to CR, no significant changes were observed in these serum

markers between control diet-fed and calorie-restricted Sirt3�/�


mice, suggesting a possible role of Sirt3 in metabolic adapta-

tions to CR.

Sirt3 Is Required for the CR-Mediated Reductionof Oxidative Damage in Multiple TissuesHow does Sirt3 reduce cochlear cell degeneration and slow the

progression of AHL in response to CR? It is well established that

CR reduces oxidative damage to DNA, proteins, and lipids in

multiple tissues in mammals (Sohal and Weindruch, 1996;

Masoro, 2000; Hamilton et al., 2001). Hence, we hypothesized

that Sirt3 may play a role in the CR-mediated reduction of oxida-

tive damage in the cochlea and other tissues. To test this hypoth-

esis, we measured oxidative damage to DNA in the cochleae,

brain (neocortex), and liver of control diet and calorie-restricted

WT and Sirt3�/� mice at 12 months of age. We found that CR

reduced oxidative DNA damage in WT mice, as determined by

measurements of 8-hydroxyguanosine and apurinic/aprimidinic

(AP) sites, but failed to reduce oxidative DNA damage in tissues

from Sirt3�/� mice (Figures 2A and 2B). In agreement with the

oxidative damage results, CR increased spiral ganglion neuron

survival (Figure 2C), outer hair cell survival (Figure 2D), and inner

hair cell survival (Figure 2E) in the basal regions of the cochleae of

WT mice, whereas CR failed to protect these cells in Sirt3�/�

Thre

shol

d (d

B SP

L)

WT

Frequency (kHz)

C

MK

E

IG

2 m

o C

D12

mo

CD

12

mo

CR

B

F

J

D

H

L

WT Sirt3-/-WT Sirt3

-/-

A

0

25

50

75

100

8 16 32

2mo CD12mo CD12mo CR

0

25

50

75

100

8 16 32

2mo CD12mo CD12mo CR

Sirt3-/-

**

*

****

**

**

*

Figure 1. CR Prevents AHL and Protects

Cochlear Neurons in WT Mice, but Not in

Sirt3�/� Mice

(A) ABR hearing thresholds were measured at 32,

16, and 8 kHz from control diet and/or calorie-

restricted WT (left) and Sirt3�/� (right) mice at

2 and 12 months of age (n = 9–12). *Significantly

different from 2-month-old WT or Sirt3�/� mice

(p < 0.05), **significantly different from 12-month-

old WT mice (p < 0.05). CD, control diet; CR,

calorie restricted diet.

(B–M) Neurons in the basal cochlear regions from

WT mice in control diet at 2 (B and C) and 12

(F and G) months of age and calorie-restricted

diet at 12 months of age (J and K). Neurons from

control diet Sirt3�/� mice at 2 (D and E) and 12

(H and I) months of age and calorie-restricted

Sirt3�/� mice at 12 months of age (L and M)

(n = 5). Arrows in the lower-magnification photos

indicate neuron regions. Scale bars, 100 mm (B,

F, J, D, H, and L) and 20 mm (C, G, J, E, I, and M).

Data are means ± SEM. See also Figure S1, Fig-

ure S2, and Figure S3.

mice (Figures 2C–2E). Together, these

results provide evidence that Sirt3 plays

an essential role in the CR-mediated

reduction of oxidative DNA damage in

multiple tissues.

Sirt3 Enhances the MitochondrialGlutathione Antioxidant DefenseSystem in Response to CRA previous study has shown that overex-

pression of Sirt3 increased mRNA

expression of the antioxidant genes

manganese superoxide dismutase (MnSOD) and catalase (Cat)

in primary cardiomyocytes and that Sirt3�/� primary cardiomyo-

cytes displayed higher levels of ROS compared to those of WT

cells (Sundaresan et al., 2009), suggesting that Sirt3 may regu-

late the antioxidant systems. Glutathione acts as the major small

molecule antioxidant in cells (Anderson, 1998; Halliwell and Gut-

teridge, 2007; Marı et al., 2009; Rebrin et al., 2003), and NADPH-

dependent glutathione reductase regenerates reduced gluta-

thione (GSH) from oxidized glutathione (GSSG) (Anderson,

1998; Marı et al., 2009). In healthy mitochondria from young

mice, glutathione is found mostly in the reduced form, GSH

(Marı et al., 2009). During aging, oxidized glutathione accumu-

lates, and hence an altered ratio of mitochondrial GSH to

GSSG is thought to be a marker of both oxidative stress and

aging (Rebrin et al., 2003; Schafer and Buettner, 2001; Marı

et al., 2009). Thus, we hypothesized that Sirt3 may regulate the

mitochondrial glutathione antioxidant system under CR condi-

tions. To test this hypothesis, we measured the ratio of

GSH:GSSG in the mitochondria of the inner ear, brain, and liver

of control diet and calorie-restricted WT and Sirt3�/� mice at

5 months of age. Mitochondrial GSSG levels decreased during

CR in the inner ear from WT mice, but not fromSirt3�/� mice (Fig-

ure 3B; see also Figure 3C). We also found that the ratios of


GSH:GSSG in mitochondria increased during CR in all of the

tested WT tissues (Figure 3A); however, CR failed to increase

the ratios of GSH:GSSG in Sirt3�/� tissues (Figure 3A). These

results are consistent with the histological, cochlear cell count-

ing, and oxidative DNA damage results that demonstrated that

CR reduces oxidative damage in WT tissues, but not in the

Sirt3�/� tissues. Thus, during CR, Sirt3 promotes a more reduc-

tive environment in mitochondria of multiple tissues, thereby

enhancing the glutathione antioxidant defense system.

Sirt3 Stimulates Idh2 Activity and Increases NADPHLevels in Mitochondria in Response to CREnzymes of mitochondrial antioxidant pathways require NADPH

to perform their reductive functions. NADP+-dependent Idh2

from mitochondria converts NADP+ to NADPH, thereby pro-

moting regeneration of GSH by supplying NADPH to glutathione

reductase (Jo et al., 2001). A previous in vitro study suggested

that Idh2 might be a target of Sirt3, as incubation of Sirt3 with iso-

citrate dehydrogenase led to an apparent increase in dehydro-

genase activity (Schlicker et al., 2008). Thus, we hypothesized

that, in response to CR, the mitochondrial deacetylase Sirt3

might directly deacetylate and activate Idh2, thereby regulating

the levels of NADPH and, consequently, the glutathione antioxi-

dant defense system.

To provide initial support for the hypothesis that Sirt3 regulates

Idh2 activity through deacetylation, we measured the acetylation

levels of Idh2 in the liver mitochondria of WT and Sirt3�/� mice

fed control and CR diets. In WT tissues, acetylation of Idh2

was substantial in the control diet fed tissues, but CR induced

an 8-fold decrease in acetylation (Figures 4A and 4B). Robust

acetylation of Idh2 was observed in Sirt3�/� mice from both

A

AP S

ites/

105

bpD

NA

Cochlea Brain

8-ox

odG

uo/1

06dG

uo

B Liver

C

OH

Cel

ls (%

)

D

Basal Region

020406080

100

WT Sirt3-/-

CDCR

020406080

100

WT Sirt3-/-

CDCR

0

8

16

24 CDCR

*

Neu

rons

/mm

2

**

IH C

ells

(%)

E

01000200030004000

WT Sirt3-/-0

1000200030004000

WT Sirt3-/-

CDCR

01000200030004000

Middle Region Apical Region

Basal Region Middle Region Apical Region

0

25

50

75

100

WT Sirt3-/-

CDCR

0

25

50

75

100

WT Sirt3-/-0

25

50

75

100

Basal Region Middle Region Apical Region

0

25

50

75

100 CDCR

0

25

50

75

100

0

25

50

75

100

*

*

*

* *

*

*

WT Sirt3-/- WT Sirt3-/-WT Sirt3-/-

WT Sirt3-/-WT Sirt3-/-WT Sirt3-/-



Figure 2. CR Reduces Oxidative DNA Damage and Increases Cell

Survival in the Cochleae from WT Mice, but Not from Sirt3�/� Mice

(A) Oxidative damage to DNA (apurinic/apyrimidinic sites) was measured in the

cochlea and neocortex from control diet and calorie-restricted WT andSirt3�/�

mice at 12 months of age (n = 4–5). AP sites, apurinic/apyrimidinic sites.

*Significantly different from 12-month-old WT mice (p < 0.05).

(B) Oxidative damage to DNA (8-oxodGuo) was measured in the liver from

control diet and calorie-restricted WT and Sirt3�/� mice at 12 months of age

(n = 4–5).

(C) Neuron survival (neuron density) of basal, middle, and apical cochlear

regions was measured from control diet and calorie-restricted WT andSirt3�/�

mice at 12 months of age (n = 4–5).

(D) OH (outer hair) cell survival (%) of basal, middle, and apical cochlear

regions was measured from control diet and calorie-restricted WT andSirt3�/�

mice at 12 months of age (n = 4–5).

(E) IH (inner hair) cell survival (%) of basal, middle, and apical cochlear regions

was measured from control diet and calorie-restricted WT and Sirt3�/� mice at

12 months of age (n = 4–5).

Data are means ± SEM. See also Figures 1B–1M.

A Inner Ear

GSH

:GSS

G

(nm

ole/

mg

prot

ein)

0

40

80

120

WT Sirt3-/-

CDCR*

B

GSS

G

(n

mol

e/m

g pr

otei

n)

Brain Liver

Inner Ear Brain Liver

C

GSH

(n

mol

e/m

g pr

otei

n)

Inner Ear Brain Liver

0

40

80

120

WT Sirt3-/-

*

0

5

10

15

20

0.0

0.3

0.5

0.8

1.0

WT Sirt3-/-

0

5

10

15

20 CDCR

0

40

80

120

0.0

0.3

0.5

0.8

1.0

0

10

20

30

40

*

0.0

0.2

0.4

0.6

WT Sirt3-/-

CDCR

*




Figure 3. Sirt3 Increases the Ratios Of GSH:GSSG in Mitochondria

during CR(A–C) Ratios of GSH:GSSG (A), GSSG (B), and GSH (C) were measured in the

inner ear, brain (neocortex), and liver from control diet and calorie-restricted

WT and Sirt3�/� mice at 5 months of age (n = 4–5). *Significantly different

from 12- or 5-month-old WT mice (p < 0.05). Data are means ± SEM.


control and CR diet-fed conditions, indicating that Sirt3 is

required for the CR-induced deacetylation of Idh2 (Figures 4A

and 4B). As predicted, CR induced Sirt3 protein levels that

were approximately three times higher than those observed

with control diet tissues in WT mice (Figure 4C).

To establish whether Idh2 activity is stimulated by Sirt3 under

CR conditions, we measured Idh2 activity in the mitochondria

from the liver, inner ear, and brain of control diet and calorie-

restricted WT and Sirt3�/� mice. We found that Idh2 activity

significantly increased during CR in all of the WT tissues (Fig-

ure 4D); however, CR failed to increase Idh2 activity in the

Sirt3�/� tissues (Figure 4D). If CR can induce a Sirt3-dependent

increase in Idh2 activity, we anticipated increased levels of

NADPH, providing the primary source of reducing equivalents

for the glutathione antioxidant system (Jo et al., 2001; Schafer

and Buettner, 2001). To test this hypothesis, we measured

NADPH levels in mitochondria of WT and Sirt3�/� mice. We

found that levels of NADPH increased during CR in all tissues

tested from WT mice (Figure 4E); however, no significant

changes in NADPH levels were observed between control diet

and CR Sirt3�/� tissues. Collectively, these results provide

evidence that, during CR, Sirt3 induces the deacetylation and

activation of Idh2, leading to increased levels of NADPH in

mitochondria of multiple tissues. We note that we observed

a reduction in Idh2 activity in liver from Sirt3�/� mice fed the

control diet and that this correlates with a slightly increased

level of acetylated Idh2 as compared to WT mice (Figure 4B).

However, we did not observe reduced Idh2 activity or reduced

NADPH levels in the inner ear or brain of Sirt3�/� mice. We

postulate that, under basal conditions (control diet fed), addi-

tional factors regulate mitochondrial Idh2 activity and NADPH

levels.

To provide direct evidence that Sirt3 deacetylates Idh2,

a number of biochemical experiments were performed.

Although most enzyme:substrate reactions are necessarily

transient interactions to promote rapid turnover, coimmunopre-

cipitation (co-IP) experiments can sometimes trap these inter-

actions. Co-IP experiments were performed in human kidney

cells (HEK293) cotransfected with Sirt3 and Idh2. We found

that precipitated Idh2-FLAG was able to co-IP Sirt3-HA (Fig-

ure 5A), whereas precipitated Sirt3-FLAG was able to co-IP

Idh2-MYC (Figure 5B), suggesting that a physical interaction

can occur between Sirt3 and Idh2 in human cells. However,

co-IP experiments do not prove a direct functional interaction.

To provide support for a functional interaction between Sirt3

and acetylated Idh2, deacetylation assays were carried out in

HEK293 cells (Figure 5C) and in vitro using purified components

(Figure 5D). Utilizing HEK293 cells, Idh2 was cotransfected with

or without Sirt3, isolated by immunoprecipitation with anti-MYC

antibody followed by western blotting with anti-acetyl-lysine

antibody. Coexpression with Sirt3 induced the deacetylation

of Idh2 to background levels (Figure 5C). For the in vitro anal-

ysis, acetylated Idh2 was prepared (see Figure S4 and Experi-

mental Procedures) and utilized as a substrate for purified

recombinant Sirt3 or Sirt5. Acetylation status was assessed

by western blotting with anti-acetyl-lysine antibody (Figure 5D),

and the resulting change in Idh2 activity was measured sepa-

rately (Figure 5E). We found that Sirt3, but not Sirt5, deacety-

lated IDH2 in an NAD+-dependent fashion (Figure 5E). The

corresponding Idh2 activity measurements indicated that de-

acetylation by Sirt3, but not Sirt5, stimulated Idh2 activity

by �100% (Figure 5E). Together, these data provide strong

biochemical evidence that Sirt3 deacetylates and stimulates

Idh2 activity and increases NADPH levels in mitochondria in

response to CR.

Inner Ear BrainLiver

0.0

0.2

0.4

0.6

WT Sirt3-/-

CDCR

0.0

0.2

0.4

0.6

0.8 CDCR* *

0.0

0.2

0.4

0.6

WT Sirt3-/-

CDCR

*

IDH

2 Ac

tivity

(μ

M/s

/μg

prot

ein)

D

A

NAD

PH/to

tal N

ADP

E Inner Ear BrainLiver

0.0

0.3

0.5

0.8

1.0

0.0

0.3

0.5

0.8

1.0

0.0

0.3

0.5

0.8

1.0 CDCR

*

*

*

Rel

ativ

e ID

H2

Acet

ylat

ion

Leve

l (%

)

C

WT

WB: -IDH2

CD

INPUT

WB: -Sirt3

WB: -IDH2

WB: -AcKIP: -IDH2

CR CD CRSirt3-/-



*

Rel

ativ

e Si

rt3 P

rote

in

Leve

l

B

0.01.0

2.03.0

4.0

CD CR

*

WT

0

50

100

150

200

WT Sirt3-/-

CDCR

WT Sirt3-/-

*

*

CD CR

Figure 4. Sirt3 Increases Idh2 Activity and NADPH Levels in Mito-

chondria by Decreasing the Acetylation State of Idh2 during CR

(A) (Top) Western blot analysis of Sirt3 and Idh2 levels in the liver from 5-month-

old WT or Sirt3�/� fed either control or calorie-restricted diet. (Bottom) Endog-

enous acetylated Idh2 was isolated by immunoprecipitation with anti-Idh2

antibody followed by western blotting with anti-acetyl-lysine antibody (n = 3).

(B and C) Quantification of the amounts of total Idh2 acetylation (B) and Sirt3

protein (C) from (A). Western blot was normalized with Idh2 levels or Sirt3 levels

quantified and analyzed by Image software (n = 3).

(D) Idh2 activities were measured in the liver, inner ear (cochlea), and brain

(neocortex) from control diet and calorie-restricted WT and Sirt3�/� mice at

5 months of age (n = 3–5).

(E) Ratios of NADPH:total NADP (NADP+ + NADPH) were measured in the liver,

inner ear, and brain (neocortex) from control diet and caloric restricted WT and

Sirt3�/� mice at 5 months of age (n = 3–5). *Significantly different from control

diet fed WT mice (p < 0.05).

Data are means ± SEM.


Overexpression of Sirt3 and/or Idh2 Increases NADPHLevels and Protects Cells from OxidativeStress-Induced Cell DeathOur physiological, histological, and biochemical results indicate

that Sirt3 mediates reduction of oxidative damage by deacetyla-

tion and stimulating the activity of Idh2, which increases NADPH

levels for antioxidant systems in mitochondria during CR. To

provide support for this mechanism, we investigated whether

Sirt3 and Idh2 are sufficient to alter the NADPH levels in cultured

cells. HEK293 cells stably transfected with vector, Sirt3, Idh2, or

Sirt3 with Idh2 were generated, and their NADPH levels were

measured. NADPH levels were significantly increased when

either Idh2 or Sirt3 or both proteins were stably overexpressed

in HEK293 cells (Figures 6A and 6B). Importantly, overexpres-

sion of both Sirt3 and Idh2 yielded a greater increase in NADPH

levels than either Sirt3 or Idh2 overexpressed alone (Figure 6A).

Finally, to investigate whether overexpression of Sirt3, Idh2, or

Sirt3 with Idh2 can protect cells from oxidative stress, the four

HEK293 cell lines were treated with oxidants H2O2 (hydrogen

peroxide) (Figure 6C) or menadione (Figure 6D), and cell viability

was measured. Overexpression of Sirt3 or Idh2 was sufficient to

protect cells from oxidative stress induced by both oxidants

(Figures 6C and 6D). Again, overexpression of both Sirt3 and

Idh2 led to higher cell viability than either Sirt3 or Idh2 overex-

pressed alone (Figures 6C and 6D). These results provide strong

biochemical evidence that Sirt3 mediates reduction of oxidative

stress by stimulating Idh2 activity and increasing NADPH levels

under stress conditions.

DISCUSSION

Sirt3 Reduces Oxidative Damage and Enhancesthe Glutathione Antioxidant Defense System underCR ConditionsA widely accepted hypothesis of how aging leads to age-related

hearing loss is through the accumulation of oxidative damage in

the inner ear (Someya and Prolla, 2010; Liu and Yan, 2007). In

support of this hypothesis, oxidative protein damage increases

in the cochlea of CBA/J mice (Jiang et al., 2007), and oxidative

DNA damage increases in the cochlea of C57BL/6J mice during

aging (Someya et al., 2009). Age-related hair cell loss is also

enhanced in mice lacking the antioxidant enzyme superoxide

dismutase 1 (McFadden et al., 1999), whereas the same mutant

animals show enhanced susceptibility to noise-induced hearing

loss (Ohlemiller et al., 1999). We have shown recently that over-

expression of mitochondrially targeted catalase delays the onset

of AHL in C57BL/6J mice, reduces hair cell loss, and reduces

oxidative DNA damage in the inner ear (Someya et al., 2009).

Of interest, overexpression of catalase in the mitochondria leads

to extension of life span in C57BL/6J mice, but overexpression

of catalase in the peroxisome or nucleus does not (Schriner

et al., 2005). Under normal conditions, catalase decomposes

A

IDH2-FLAG -Sirt3-HA +

WB: -HAINPUT

WB: -FLAG

IP

WB: -HA

α-IgG α-FLAG

++ -

+B

Sirt3-FLAG -IDH2-MYC +

WB: -MYCINPUT

WB: -FLAG

IP

WB: -MYC

α-IgG α-FLAG

++ -

+

C

Sirt3-FLAG -IDH2-MYC +

WB: -MYC

INPUT

WB: -AcK

WB: -FLAG

++

IP: α-MYC

NAD+ -IDH2-FLAG +

WB: -FLAG

COOMASSIE BLUE

WB: -AcK

++

IP: α-FLAG

D

SIRTUIN - --+ +

+Sirt3

-+ +

+Sirt5

E

Rel

ativ

e ID

H2

Activ

ity (%

)

Sirt3 Sirt5

050

100150200250

IDH

2

IDH

2+N

AD

IDH

2+Si

rt3

IDH

2+Si

rt3+N

AD

IDH

2+Si

rt5

IDH

2+Si

rt5+N

AD

*

Figure 5. Sirt3 Directly Deacetylates Idh2

and Stimulates Activity

(A and B) Sirt3 interacts with Idh2. Idh2 or Sirt3

were immunoprecipitated from HEK293 cell

lysates with IgG antibody or FLAG beads. Precip-

itated Idh2-FLAG was detected by anti-FLAG anti-

body, and co-IP Sirt3-HA was detected by anti-HA

as indicated (A). Precipitated Sirt3-FLAG was

detected by anti-FLAG antibody, and co-IP Idh2-

MYC was detected by anti-MYC as indicated (B)

(n = 3).

(C) Sirt3 deacetylates Idh2 in HEK293 cells. Idh2

was cotransfected with or without Sirt3, isolated

by immunoprecipitation with anti-MYC antibody

followed by western blotting with anti-acetyl-

lysine antibody (n = 3).

(D) Sirt3, but not Sirt5, deacetylates Idh2 in vitro.

Acetylated Idh2 was prepared as outlined in the

Experimental Procedures and was incubated

with purified recombinant Sirt3 or Sirt5 with or

without NAD+ at 37�C for 1 hr. Acetylation status

was assessed by western blotting with anti-

acetyl-lysine antibody (n = 3). An anti-FLAG

western shows that equivalent Idh2 protein levels

were used, and Coomassie staining shows puri-

fied Sirt3 and Sirt5.

(E) In vitro deacetylation of Idh2 by Sirt3, but not

Sirt5, stimulates Idh2 activity. Acetylated Idh2 in

buffer (Tris [pH 7.5], with or without 1 mM NAD,

and 1 mM DTT) was incubated with purified

50 nM Sirt3 or Sirt5 (Hallows et al., 2006) at

37�C for 1 hr, followed by Idh2 activity assay

(n = 3). *Significantly different from Idh2 alone

(p < 0.05).

Data are means ± SEM. See also Figure S4.


hydrogen peroxide in the peroxisome, whereas in mitochondria,

hydrogen peroxide is decomposed into water by glutathione

peroxidase or peroxiredoxin (Finkel and Holbrook, 2000; Marı

et al., 2009). Hence, these results suggest that mitochondrial

ROS play a critical role in cochlear aging, AHL, and aging in

general.

We have demonstrated that Sirt3 mediates the CR reduction

of oxidative DNA damage in multiple tissues and that these

effects are likely to arise through an enhanced mitochondrial

glutathione antioxidant defense system. As discussed earlier,

the GSH:GSSG ratio is thought to be a marker of oxidative stress

(Rebrin and Sohal, 2008). Experimental evidence indicates that

aging results in a decrease in the ratio of GSH:GSSG in the mito-

chondria of brain, liver, kidney, eye, heart, and testis from aged

C57BL/6J mice due to elevated levels of GSSG, whereas CR

decreases the ratio of GSH:GSSG in the mitochondria of these

tissues by lowering GSSG levels (Rebrin et al., 2003, 2007).

Our findings demonstrate that CR increases these ratios of

GSH:GSSG in the mitochondria of brain, liver, and inner ear

from WT mice but fails to increase the ratios in the same tissues

from Sirt3�/� mice. Consistent with these results, CR reduced

oxidative DNA damage in tissues from WT mice but failed to

reduce such damage in tissues from Sirt3�/� mice. CR also

increased spiral ganglion neuron and hair cell survival in the

WT cochlea, but not in Sirt3�/� mice. Tissues that are composed

of postmitotic cells such as the brain and the inner ear are partic-

ularly vulnerable to oxidative damage because of their high

energy requirements and inability to undergo regeneration.

Therefore, we speculate that the Sirt3-mediated modulation of

C

Cel

l Via

bilit

y (%

)

B

pCDNA3-Sirt3-FLAG -pBabe-IDH2-FLAG

IDH2-FLAG

Sirt3-FLAG

INPUT

+-

+

WB: α-FLAG

-+-

+

A

[NA

DP

H] (

nmol

/mg

prot

ein)

0

60

120

180

VEC

Sirt3

IDH

2

Sirt3

+ID

H2

**

***

0

30

60

90

120

VEC

VEC

Sirt3

IDH

2

Sirt3

+ID

H2

0

30

60

90

120

VEC

VEC

Sirt3

IDH

2

Sirt3

+ID

H2

**

**

*

**

**

*

+- + + +1 mM H2O2 +- + + +

25 μM Menadione

Cel

l Via

bilit

y (%

)

D

Figure 6. Overexpression of Sirt3 and/or

Idh2 Is Sufficient to Increase NADPH Levels

and Protects HEK293 Cells from Oxidative

Stress

(A and B) (A) NADPH concentrations were sig-

nificantly increased when either Idh2 or Sirt3 or

both were stably overexpressed in HEK293 cells.

Measurements with errors are shown for the four

different stable cell populations from each type

of transfection (vector alone, Sirt3, Idh2, and

Sirt3 with Idh2) (n = 3). *Significantly different

from vector alone (p < 0.05); **Significantly dif-

ferent from Idh2 or Sirt3 (p < 0.05). (B) Western

blotting confirms Idh2 and Sirt3 stable expression.

(C and D) Sirt3 and/or Idh2 overexpression is suffi-

cient to protect HEK293 cells from the exogenous

oxidants hydrogen peroxide (H2O2) (C) and mena-

dione (D). The four different stable cells were tran-

siently exposed to either 1 mM H2O2 or 25 mM

menadione (n = 16).

Data are means ± SEM.

the glutathione antioxidant defense sys-

tem may play a central role in reduction

of oxidative stress in multiple tissues

under CR conditions, leading to aging

retardation. We also note that other mito-

chondrial effects of Sirt3, such as regula-

tion of fatty acid oxidation (Hirschey et al.,

2010) and modulation of complex I

activity (Ahn, et al., 2008), are likely to contribute to the metabolic

adaptations in response to CR.

Idh2 Regulates the Redox State of Mitochondria underCR ConditionsA large body of evidence indicates that the antioxidant defense

systems do not keep pace with the age-related increase in

ROS production, and thus the balance between antioxidant

defenses and ROS production shifts progressively toward

a more pro-oxidant state during aging (Sohal and Weindruch,

1996; Rebrin and Sohal, 2008). This balance is determined in

part by the ratios of interconvertible forms of redox couples,

such as GSH/GSSG, NADPH/NADP+, NADH/NAD+, thioredox-

inred/thioredoxinoxid, and glutaredoxinred/glutaredoxinoxid. The

GSH/GSSH couple is thought to be the primary cellular determi-

nant of the cellular redox state because its abundance is three to

four orders of magnitude higher than the other redox couples

(Rebrin and Sohal, 2008). NADPH is the reducing equivalent

required for the regeneration of GSH and the GSH-mediated

antioxidant defense system, which includes glutathione peroxi-

dases, glutathione transferases, and glutathione reductase,

playing a critical role in oxidative stress resistance (Halliwell

and Gutteridge, 2007). GSH is synthesized in the cytosol and

transported into the mitochondria through protein channels in

the outer mitochondrial membrane (Halliwell and Gutteridge,

2007; Anderson, 1998). Although GSH can cross the outer mito-

chondrial membrane through these channels, GSSG cannot be

exported into the cytosol (Olafsdottir and Reed, 1988). Thus,

GSSG is reduced to GSH by mitochondrial NADPH-dependent


glutathione reductase, preventing accumulation of GSSG in the

mitochondrial matrix (Schafer and Buettner, 2001; Marı et al.,

2009). We have demonstrated that Sirt3 directly deacetylates

and activates Idh2 under CR conditions. In response to CR,

deacetylated Idh2 displays increased catalytic activity, which is

correlated with increased NADPH levels in the mitochondria of

multiple tissues from WT mice, but not from Sirt3�/� mice.

Hence, we speculate that Idh2 may be a major player in regu-

lating the redox state of mitochondria under CR conditions given

its role in mitochondrial NADPH production. A previous study

has shown that Idh2 is induced in response to ROS in mouse

fibroblasts, whereas decreased levels of Idh2 lead to higher

ROS and accumulation of oxidative damage to DNA and lipids

(Jo et al., 2001). Our in vitro findings demonstrate that overex-

pression of Sirt3 and/or Idh2 increases NADPH levels and

protects cells from oxidative stress-induced cell death. Thus,

these observations underlie a critical role for Idh2 in the genera-

tion of NADPH in mitochondria under conditions of CR, providing

reducing capacity for the glutathione antioxidant system and

increasing oxidative stress resistance.

A Role for Sirt3 in CR-Mediated Prevention of AHLThe mouse is considered a good model for the study of human

AHL because the mouse cochlea is anatomically similar to that

of humans (Steel et al., 1996; Steel and Bock, 1983). Most in-

bred mouse strains display some degree of AHL, and the age

of onset of AHL is known to vary from 3 months in DBA/2J

mice to more than 20 months in CBA/CaJ mice (Zheng et al.,

1999). The C57BL/6J mouse strain, which is the most widely

used mouse model for the study of aging, displays the classic

pattern of AHL by 12–15 months of age (Hunter and Willott,

1987; Keithley et al., 2004). We have previously shown that

AHL in C57BL/6J mice occurs through Bak-mediated apoptosis

and that it can be prevented by the intake of small molecule anti-

oxidants (Someya et al., 2009). We note that C57BL/6J and many

other mouse strains carry a specific mutation (Cdh23753A) in the

Cdh23 gene, which encodes a component of the hair cell tip

link, and this mutation is known to promote early onset of AHL

in these animals (Noben-Trauth et al., 2003). Of interest, the

Cdh23753A allele may increase the susceptibility to oxidative

stress in hair cells because a Sod1 mutation greatly enhances

AHL in mice carrying Cdh23753A, but not in mice wild-type for

Cdh23 (Johnson, et al., 2010). However, oxidative damage

increases with age in the cochlea of both C57BL/6J mice and

the CBA/J mouse strain that does not carry the Cdh23753A allele,

indicating that oxidative stress plays a role in AHL independent

of Cdh23 (Someya et al., 2009; Jiang et al., 2007; Zheng et al.,

1999). In both strains, the loss of hair cells and spiral ganglion

neurons begins in the base of the cochlea and spreads toward

the apex with age (Keithley et al., 2004; Hunter and Willott,

1987). Importantly, CR slows the progression of AHL in both

C57BL/6J and CBA/J strains (Someya et al., 2007; Sweet

et al., 1988). Therefore, the protective effects of Sirt3 in AHL

are likely to be of general relevance to AHL.

It is thought that some of the effects of CR in aging retardation

require significant reduction of body weight through reducing

food consumption. In agreement with this hypothesis, obesity

promotes a variety of age-related diseases, such as cardiovas-

cular disease, diabetes, high blood pressure, hypertension,

and certain cancers (Paeratakul et al., 2002; Poirier et al.,

2006). Obesity is also associated with an increased risk of

mortality (Poirier et al., 2006; Lee et al., 1993). Of interest, CR

failed to reduce oxidative damage in multiple tissues and slow

the progression of AHL in CR Sirt3�/� mice, despite the fact

that these mice were lean (Figures S2A and S2B). Thus, these

results suggest that weight loss may not be sufficient for the

anti-aging action of CR. Instead, we postulate that critical meta-

bolic effectors such as Sirt3 mediate the positive effects of CR.

ConclusionsIn summary, we propose that, in response to CR, Sirt3 activates

Idh2, thereby increasing NADPH levels in mitochondria. This in

turn leads to increased ratios of GSH:GSSG in mitochondria

and decreased levels of ROS, resulting in protection of inner ear

cells and prevention of AHL in mammals (Figure 7). Because we

observed similar effects of CR in the mitochondrial GSH/GSSG

ratios in multiple tissues, we postulate that this may be a major

mechanism of aging retardation by CR. We also postulate that

pharmaceutical interventions that induce Sirt3 activity in multiple

tissues will mimic CR by increasing oxidative stress resistance

and preventing the mitochondrial decay associated with aging.


Animals

Male and female Sirt3+/� mice were purchased from the Mutant Mouse

Resource Centers (MMRRC) at the University of North Carolina-Chapel Hill

Figure 7. A Model for the CR-Mediated Prevention of AHL in

Mammals

In response to CR, SIRT3 activates IDH2, thereby increasing NADPH levels in

mitochondria. This in turn leads to an increased ratio of GSH:GSSG and

decreased levels of ROS, thereby resulting in protection from oxidative stress

and prevention of AHL in mammals.


(Chapel Hill, NC). In brief, these mice were created by generating embryonic

stem (ES) cells (Omni bank number OST341297) bearing a retroviral promoter

trap that functionally inactivates one allele of the Sirt3 gene (MGI, 2010).

Male and female C57BL/6J mice were purchased from Jackson Laboratory

(Bar Harbor, ME). Sirt3+/� mice have been backcrossed for four generations

onto the C57BL/6J background. All animal studies were conducted at the

AAALAC-approved Animal Facility in the Genetics and Biotechnology Center

of the University of Wisconsin-Madison. Experiments were performed in

accordance with protocols approved by the University of Wisconsin-Madison

Institutional Animal Care and Use Committee (Madison, WI).

Dietary Study

Details on the methods used to house and feed mice have been described

previously (Pugh et al., 1999). Mice are housed individually. Control diet (CD)

groups were fed 86.4 kcal/week of the precision pellet diet AIN-93M (BioServ,

Frenchtown, NJ), and caloric-restricted (CR) groups were fed 64.8 kcal/week

(a 25% CR) of the precision pellet diet AIN-93M 40%DR (BioServ, Frenchtown,

NJ). The schedule of feeding for control diet was 7 g on Mondays and Wednes-

days and 10 g on Fridays, whereas the schedule of feeding for calorie-

restricted diets was 5 g on Mondays and Wednesdays and 8 g on Fridays.

This dietary regimen was maintained from 2 months of age until 5 months of

age for a 3 month CR study and from 2 months of age until 12 months of

age for a 10 month CR study.

ABR Hearing Test

At 12 months of age, ABRs were measured with a tone burst stimulus at 8, 16,

and 32 kHz using an ABR recording system (Intelligent Hearing System, Miami,

FL) as previously described (Someya et al., 2009). Mice were anesthetized

with a mixture of xylazine hydrochloride (10 mg/kg, i.m.) (Phoenix Urology of

St. Joseph, St. Joseph, MO) and ketamine hydrochloride (40 mg/kg, i.m.)

(Phoenix Urology of St. Joseph).

Measurement of DNA Oxidation Levels

At 12 months of age, cochlea and neocortex were collected, and DNA was

extracted with ethanol precipitation. DNA concentrations for each sample

were adjusted to 0.1 mg/ml, and numbers of apurinic/apyrimidinic (AP) sites

were determined using the DNA Damage Quantification Kit (Dojindo, Rockville,

MD) and performed according to the manufacturer’s instructions and as previ-

ously described (Kubo et al., 1992; Meira, et al., 2009; McNeill and Wilson,

2007). Liver was also collected from the same mice, and 8-hydroxyguanosine

levels (8-oxo-7,8-20-deoxyguanosine/106 deoxyguanosine) in the DNA were

determined using a HPLC-ECD method as previously described (Hofer

et al., 2006).

Measurement of Total GSH and GSSG

Just after mitochondrial lysate preparation, 100 ml of the lysate was mixed with

100 ml of 10% metaphosphoric acid, incubated for 30 min at 4�C, and centri-

fuged at 14,000 3 g for 10 min at 4�C. The supernatant was used for the

measurements of mitochondrial glutathione contents. Total glutathione

(GSH + GSSG) and GSSG levels were determined by the method of Rahman

et al. (2006). All samples were run in duplicate. The rates of 2-nitro-5-thioben-

zoic acid formation were calculated, and the total glutathione (tGSH) and

GSSG concentrations in the samples were determined by using linear regres-

sion to calculate the values obtained from the standard curve. The GSH

concentration was determined by subtracting the GSSG concentration from

the tGSH concentration.

Idh2 Acetylation Analysis

Antibodies used for western blotting included anti-Idh2 antibody (Santa Cruz,

Santa Cruz, CA), anti-Sirt3 antibody (gift of Dr. Eric Verdin, UCSF), protein A/G

plus agarose (Santa Cruz, Santa Cruz, CA), and pan-acetylated lysine (gener-

ated following the procedure of Zhao, et al. [2010], GeneTel Laboratories LLC,

Madison, WI). For immunoprecipitation, liver mitochondria lysates were incu-

bated with anti-Idh2 antibody overnight at 4�C. Then protein A/G plus agarose

were added and incubated for 3 hr. After resins were washed, samples were

boiled with SDS loading buffer and subjected to western blotting (Smith

et al., 2009).

Idh2 Activity

Activities of Idh2 were measured by the Kornberg method (Kornberg, 1955). In

brief, 20 ml of the mitochondrial lysate sample was added in each well of a

96-well plate, and then 180 ml of a reaction mixture (33 mM KH2PO4dK2HPO4,

3.3 mM MgCl2, 167 mM NADP+, and 167 mM (+)-potassium Ds-threo-isocitrate

monobasic) was added in each well. The absorbance was immediately read at

340 nm every 10 s for 1 min in a microplate reader (Bio-Rad, Hercules, CA). All

samples were run in duplicate. The reaction rates were calculated, and the

Idh2 activity in the sample was defined as the production of one mmole of

NADPH per sec.

In Vitro Deacetylation Assay

Idh2-FLAG was transfected into HEK293 cells, which were then treated with

5 mM nicotinamide for 16 hr. Nicotinamide is a widely used sirtuin inhibitor.

Nicotinamide treatment leads to increased acetylation of Idh2, with a corre-

sponding decrease in enzymatic activity (Figure S4). Idh2 from cell lysates

was immunoprecipitated with anti-FLAG beads at 4�C for 2 hr, and then

Idh2-FLAG on beads was utilized in 200 ul deacetylation buffer (Tris

[pH 7.5], with or without 1 mM NAD, and 1 mM DTT) and incubated with puri-

fied 50 nM Sirt3 or Sirt5 (Hallows et al., 2006) at 37�C for 1 hr. Aliquots were

removed for Idh2 activity assay and western blotting with anti-FLAG antibody

or anti-acetyl-lysine antibody.

Statistical Analysis

All Statistical analyses were carried out by one-way ANOVA with post-Tukey

multiple comparison tests using the Prism 4.0 statistical analysis program

(GraphPad, San Diego, CA). All tests were two-sided with statistical signifi-

cance set at p < 0.05.


Supplemental Information includes Extended Experimental Procedures, four

figures, and one table and can be found at doi:10.1016/j.cell.2010.10.002.

ACKNOWLEDGMENTS

We thank S. Kinoshita for histological processing. This research was sup-

ported by NIH grants AG021905 (T.A.P.) and GM065386 (J.M.D.), the National

Projects on Protein Structural and Functional Analyses from the Ministry of

Education, Culture, Sports, Science, and Technologies of Japan, and Marine

Bio Foundation.

Received: July 19, 2010


Accepted: September 30, 2010

Published online: November 18, 2010

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FOXO/4E-BP Signaling in DrosophilaMuscles Regulates Organism-wideProteostasis during AgingFabio Demontis1,* and Norbert Perrimon1,2,*1Department of Genetics2Howard Hughes Medical Institute

Harvard Medical School, 77 Avenue Louis Pasteur, Boston, MA 02115, USA

*Correspondence: [email protected] (F.D.), [email protected] (N.P.)DOI 10.1016/j.cell.2010.10.007

SUMMARY

The progressive loss of muscle strength during agingisa commondegenerativeevent of unclearpathogen-esis. Although muscle functional decline precedesage-related changes in other tissues, its contributionto systemic aging is unknown. Here, we show thatmuscle aging is characterized in Drosophila by theprogressive accumulation of protein aggregatesthat associate with impaired muscle function. Thetranscription factor FOXO and its target 4E-BP re-move damaged proteins at least in part via theautophagy/lysosome system, whereas foxo mutantshave dysfunctional proteostasis. Both FOXO and4E-BP delay muscle functional decay and extend lifespan. Moreover, FOXO/4E-BP signaling in musclesdecreases feeding behavior and the release of insulinfrom producing cells, which in turn delays the age-related accumulation of protein aggregates in othertissues. These findings reveal an organism-wideregulation of proteostasis in response to muscleaging and a key role of FOXO/4E-BP signaling in thecoordination of organismal and tissue aging.

INTRODUCTION

Aging of multicellular organisms involves distinct pathogenic

events that include higher mortality, the progressive loss of

organ function, and susceptibility to degenerative diseases,

some of which arise from protein misfolding and aggregation.

Recent genetic studies in the mouse, the nematode Caenorhab-

ditis elegans, and the fruitfly Drosophila melanogaster have

expanded our understanding of the evolutionarily conserved

signaling pathways regulating aging, with the identification of

several mutants that have prolonged or shortened life spans

(Kenyon, 2005). Manipulation of longevity-regulating pathways

in certain tissues is sufficient to extend life expectancy, indi-

cating that some tissues have a predominant role in life span

extension (Libina et al., 2003; Wang et al., 2005; Wolkow et al.,

2000). For example, foxo overexpression in the Drosophila fat

body extends life span, indicating a key role of this tissue in

the regulation of longevity (Giannakou et al., 2004; Hwangbo

et al., 2004). In addition, because most tissues undergo progres-

sive deterioration during aging (Garigan et al., 2002), it is thought

that organismal life span may be linked to tissue senescence.

However, our understanding of the mechanisms regulating

tissue aging and their interconnection to life span is limited. For

example, analysis in Drosophila has revealed that the prevention

of age-dependent changes in cardiac performance does not

alter life span (Wessells et al., 2004), raising the possibility that

functional decline in distinct tissues may have different

outcomes on the systemic regulation of aging.

The Insulin/IGF-1 signaling pathway has been implicated in the

control of aging across evolution via its downstream signaling

component FOXO (DAF-16 in C. elegans), a member of the

fork-head box O transcription factor family (Salih and Brunet,

2008). FOXO regulates the expression of a series of target genes

involved in metabolism, cell growth, cell proliferation, stress

resistance, and differentiation via direct binding to target gene

promoter regions (Salih and Brunet, 2008). Mutations in foxo/

daf-16 reduce life span and stress resistance in both C. elegans

and flies, indicating a key role in organism aging (Junger et al.,

2003; Salih and Brunet, 2008). In addition to regulating life

span, FOXO has been reported to prevent the pathogenesis of

some age-related diseases. For example, FOXO reduces the

toxicity associated with aggregation-prone human mutant

Alzheimer’s and Huntington’s disease proteins (proteotoxicity)

in C. elegans and mice, suggesting that regulating protein

homeostasis (proteostasis) during aging may have a direct effect

on the pathogenesis of human neurodegenerative diseases

(Cohen et al., 2006; Hsu et al., 2003; Morley et al., 2002).

However, little is known on the protective mechanisms induced

in response to FOXO signaling and whether they vary in different

aging tissues and disease contexts.

Among the plethora of age-related pathological conditions,

the gradual decay in muscle strength is one of the first hallmarks

of aging in many organisms, including Drosophila, C. elegans,

mice and, importantly, humans (Augustin and Partridge, 2009;

Herndon et al., 2002; Nair, 2005; Zheng et al., 2005). However,

despite its medical relevance, the mechanisms underlying

muscle aging are incompletely understood. Functional changes

in skeletal muscles temporally precede the manifestation of


Figure 1. FOXO Signaling in Skeletal Muscles Preserves Proteostasis during Aging

(A–D) Electron micrographs of immunogold-labeled Drosophila skeletal muscles of wild-type flies at one (A and B) and 5 weeks of age (C and D). Protein

aggregates (PA) are detected in the cytoplasm in proximity to mitochondria (Mt) and myofibrils (Myof) in old (C and D) but not young flies (A and B). Numerous

gold particles (indicative of anti-ubiquitin immunoreactivity) localize to filamentous structures at 5 weeks of age (C and D), while only a few are present in muscles

from young flies. Scale bars are 1 mm (A and C) and 500 nm (B and D).


aging in other tissues (Herndon et al., 2002), and reduced muscle

strength is associated with an increased risk in developing

Alzheimer’s and Parkinson’s diseases (Boyle et al., 2009; Chen

et al., 2005). However, although aging-related changes in

skeletal muscles have been proposed to affect physiological

processes in distal organs (Nair, 2005), whether or not muscle

senescence modulates the pathogenesis of degenerative events

in other tissues is unknown.

The fruit fly Drosophila is an excellent model to study muscle

aging. The progressive decline in muscle strength and function

observed in humans is recapitulated in this system (Rhodenizer

et al., 2008), which is amenable to extensive genetic manipula-

tion. By using this model organism, we have searched for the

molecular mechanisms responsible for muscle aging and

found that decreased protein quality control plays a role in the

pathogenesis of age-related muscle weakness. Interestingly,

increased activity of the transcription factor FOXO and its target

Thor/4E-BP are sufficient to delay this process and preserve

muscle function at least in part by promoting the basal activity

of the autophagy/lysosome system, an intracellular protein

degradation pathway that removes damaged protein aggregates

(Rubinsztein, 2006).

Moreover, we report that FOXO/4E-BP signaling in muscles

extends life span and regulates proteostasis organism-wide by

regulating feeding behavior, release of insulin from producing

cells, and 4E-BP induction in nonmuscle tissues. Thus, we

propose a model by which FOXO/4E-BP signaling in muscles

preserves systemic proteostasis by mimicking some of the

protective effects of decreased nutrient intake.

RESULTS

Loss of Proteostasis during Muscle AgingIs Prevented by FOXOTo detect cellular processes that are responsible for decreased

muscle strength in aging flies, we monitored cellular changes in

indirect flight muscles of wild-type flies by immunogold-electron

microscopy (IEM). In older flies, we detected filamentous cyto-

plasmic structures that were instead absent in muscles from

young flies (Figures 1A–1D). Filamentous materials present in

these structures stained with an anti-ubiquitin antibody (Fig-

ure 1D), a marker for proteins that are polyubiquitinated, sug-

gesting that the cytoplasmic structures are aggregates of

damaged proteins. Aggregates were variable in size and were

detected in both resin-embedded sections (Figure 1) and cryo-

sections (data not shown) of thoracic muscles of the old but

not the young flies, in parallel with an increase in the overall

number of gold particles (Figure 1E). To test the hypothesis

that muscle function during aging may decrease due to defects

in protein homeostasis, we better characterized the age-related

deposition of protein aggregates by immunofluorescence. In

agreement with the IEM analysis (Figures 1A–1E), we observed

that aging skeletal muscles progressively accumulate aggre-

gates of polyubiquitinated proteins (ranging up to several mm)

that colocalize with p62/Ref(2)P, an inclusion body component

(Figures 1F and1I). The cumulative area of protein aggregates

increases during aging (Figure 1L), suggesting that the progres-

sive protein damage, together with a decrease in the turnover of

muscle proteins, may result in the age-related decline of muscle

strength.

To better characterize how protein quality control is linked with

aging in muscles, we analyzed the deposition of protein

aggregates in syngenic flies with foxo overexpression. Foxo

overexpression results in its activation (Giannakou et al., 2004;

Hwangbo et al., 2004) and was achieved specifically in muscles

via the UAS-Gal4 system using the Mhc-Gal4 driver (see Fig-

ure S1 available online). Increased FOXO activity in muscles

did not affect developmental growth and differentiation (as esti-

mated by body weight and sarcomere assembly) (Figure S2), and

resulted in the delayed accumulation of aggregates containing

polyubiquitinated proteins and Ref(2)P during aging (Figures 1G

and 1J, compare with control muscles in Figures 1F and 1I).

Next, we tested whether foxo null animals display accelerated

muscle aging, and found an increased accumulation of protein

aggregates (Figures 1H and1K), indicating that FOXO is both

necessary and sufficient to modulate muscle proteostasis

(Figure 1L).

To further corroborate these findings, we overexpressed

either the wild-type or the constitutive-active foxo transgenes

using the Dmef2-Gal4 muscle driver in combination with the

temperature-sensitive tubulin-Gal80ts transgene to achieve

adult-onset foxo overexpression in muscles (Figure S3). Trans-

gene overexpression significantly preserved muscle proteosta-

sis in both cases, while the controls displayed an increased

accumulation of protein aggregates (Figure S3). All together,

these results indicate that protein homeostasis depends on

FOXO activity during muscle aging.

4E-BP Controls Proteostasis in Responseto Pten/FOXO ActivityTo dissect the stimuli that encroach on FOXO to control proteo-

stasis, we tested whether Pten overexpression phenocopies

FOXO activation. Consistent with its role in activating FOXO,

we found that Pten decreased the accumulation of protein

(E) The number of gold particles, indicative of ubiquitin immunoreactivity, significantly increases in old age (standard error of the mean [SEM] is indicated with n;

**p < 0.01).

(F–L) Immunostaining of indirect flight muscles from flies with (UAS-foxo/+;Mhc-Gal4/+) or without (Mhc-Gal4/+) foxo overexpression at 1 week (F and G) and

5 weeks of age (I and J), and foxo homozygous null (MhcGal4, foxo21/25) flies (H and K). Polyubiquitin (red) and p62/Ref(2)P (green) immunoreactivities reveal

an increased deposition of aggregates containing polyubiquitinated proteins during aging in muscles of control flies (F and I), and, to a lesser extent, in muscles

overexpressing foxo (G and J). Conversely, muscles from foxo null animals display an accelerated deposition of protein aggregates (H and K) in comparison with

controls (F and I). Note the significant increase in the cumulative area of protein aggregates (indicative of both aggregate size and number) in (K) versus (I), and in (I)

versus (J), indicating that the control of protein homeostasis is linked to FOXO activity in muscles (quantification in [L]) (SEM is indicated with n; *p < 0.05, **p <

0.01). Representative polyubiquitin and Ref(2)P immunoreactivities are shown in insets. Phalloidin staining (blue) outlines F-actin, which is a component of muscle

myofibrils. Scale bar is 20 mm (F–K).

See also Figure S1, Figure S2, and Figure S3.


aggregates during aging (Figures 2B and 2E; see controls in

Figures 2A and 2D).

Next, we examined the responses induced by Pten/FOXO

signaling. First, we examined whether FOXO activity delays

protein damage by inducing chaperones that are key for protein

quality control (Tower, 2009). In response to FOXO activity in

muscles, we detected an increase in the mRNA levels of

Hsp70 and its cofactors involved in protein folding (Hip, Hop,

Hsp40, and Hsp90) but not in protein degradation (Chip and

Chap) (Figure S4 and Table S1). FOXO regulates directly the

expression of Hsp70 and its cofactors, as estimated with Lucif-

erase transcriptional reporters based on the proximal promoter

region of target genes (Figure S4 and Table S2). On this basis,

we tested whether Hsp70 overexpression preserves proteosta-

sis during aging but found little changes in the age-related accu-

mulation of protein aggregates (Figure S4). Thus, we conclude

that additional FOXO-dependent responses are involved.

Among the FOXO-target genes, Thor/4E-BP has a key role

in delaying aging by regulating protein translation (Zid et al.,

2009; Tain et al., 2009). However, the cellular mechanisms that

Figure 2. 4E-BP Preserves Proteostasis in Response to Pten/FOXO Signaling

(A–F) Immunostaining of muscles overexpressing Pten and constitutive active (CA) 4E-BP. In both cases, a decrease in the accumulation of polyubiquitin protein

aggregates is observed at 5 weeks of age in comparison with age-matched controls, suggesting that these interventions can preserve proteostasis in aging

muscles. Scale bar is 20 mm. Hsp70 overexpression has instead limited effects (Figure S4, Table S1, Table S2).

(G) A reduction in the cumulative area of protein aggregates is observed upon increased activity of either Pten or 4E-BP in comparison with controls (SEM is

indicated with n; **p < 0.01, ***p < 0.001).

(H) Relative quantification of Thor/4E-BP mRNA levels from thoraces of syngenic flies at 1 and 5 weeks of age. A significant increase in 4E-BP expression is

detected in response to fasting and Pten and FOXO activity (**p < 0.01, ***p < 0.001; SEM is indicated with n = 4).


are regulated by 4E-BP are largely unknown. To test whether

4E-BP controls proteostasis during muscle aging, we overex-

pressed a constitutive active form of 4E-BP in muscles and

observed limited accumulation of protein aggregates during

aging (Figures 2C and 2F) compared with controls (Figures 2A

and 2D). All together, increased activity of Pten or 4E-BP sig-

nificantly decreases the cumulative area of protein aggregates

(Figure 2G).

In addition, a significant increase in 4E-BP mRNA levels is

induced in muscles upon Pten, foxo overexpression, and fasting

(Figure 2H). All together, these findings suggest that 4E-BP is key

to control proteostasis in response to Pten/FOXO signaling.

FOXO/4E-BP Signaling Regulates Proteostasis viathe Autophagy/Lysosome SystemWhile FOXO/4E-BP signaling mounts a stress resistance

response that may decrease the extent of protein damage due

to various stressors (Salih and Brunet, 2008; Tain et al., 2009),

we wondered whether it regulates the removal of damaged

proteins via macroautophagy. In this process, entire regions of

the cytoplasm are sequestered in a double membrane vesicle

(autophagosome) that subsequently fuses with a lysosome,

where the autophagic cargo is degraded (Rubinsztein, 2006).

Although the primary role of autophagy is to mount an adaptive

response to nutrient deprivation, its basal activity is required

for normal protein turnover (Hara et al., 2006). In agreement

with this notion, suppression of basal autophagy leads to the

accumulation of polyubiquitin protein aggregates in a number

of contexts (Korolchuk et al., 2009; Rubinsztein, 2006).

To test whether autophagy is regulated in response to FOXO

signaling in muscles, we used a GFP-tagged version of the

autophagosome marker Atg5 (Rusten et al., 2004). While the

number of Atg5-GFP punctae decreases during aging in control

muscles (Figures 3A and 3B), it is in part maintained in response

to foxo overexpression (Figures 3C and 3D, and quantification in

Figure 3E). In addition, given the interconnection between

the lysosome system and autophagy, we monitored a GFP-

tagged version of the lysosome marker Lamp1 (lysosome-asso-

ciated membrane protein 1) and detected an overall increase in

the number of GFP punctae in response to overexpression of the

autophagy inducer kinase Atg1, foxo, and 4E-BP CA in muscles

at both 1 and 5 weeks of age (Figures 3G–3I and 3K–3M in

comparison with controls in Figures 3F and 3J and quantification

in Figure 3N).

Closer inspection revealed that the abundance of Lamp1-GFP

vesicles inversely correlates with the progressive deposition of

polyubiquitin protein aggregates, suggesting that FOXO/4E-BP

signaling regulates proteostasis at least in part via the

autophagy/lysosome system. To further test this hypothesis,

we analyzed the age-related changes in autophagy gene expres-

sion, which have been previously used as a correlative measure-

ment of autophagic activity (Gorski et al., 2003; Simonsen et al.,

2008). Interestingly, the expression of several autophagy genes

involved in autophagosome induction (Atg1), nucleation (Atg6),

and elongation (Atg5, Atg7, and Atg8) progressively declines

during aging in muscles (Figure 3O), suggesting that gene

expression changes likely contribute to the accumulation of

damaged proteins. Conversely, foxo overexpression increased

the basal expression of several Atg genes at both young and

old age, suggesting that their increased expression contributes

to the beneficial effects of FOXO on proteostasis. To test this

hypothesis, we knocked down Atg7 levels in foxo-overexpress-

ing flies and analyzed the deposition of polyuiquitinated protein

aggregates. Interestingly, RNAi treatment brought about a

�50% decrease in Atg7 mRNA levels and resulted in a partial

increase in the buildup of insoluble ubiquitinated proteins at

8 weeks, compared with age-matched, mock-treated flies

(white RNAi) and 1-week-old flies (Figure 3P).

All together, these findings suggest that FOXO/4E-BP

signaling prevents the buildup in protein damage, at least in

part by promoting the basal activity of the autophagy/lysosome

system.

Prevention of Muscle Aging by FOXO and 4E-BPExtends Life SpanTo evaluate whether preserving proteostasis can prevent

functional alterations in aging muscles, we assessed muscle

strength with negative geotaxis and flight assays (see Experi-

mental Procedures). As shown in Figures 4A and 4B, muscle

functionality gradually decreases in aging flies, resulting in

impaired climbing and flight ability. Notably, foxo (Figure 4A)

and 4E-BP activity (Figure 4B) significantly preserve muscle

strength during aging. Thus, FOXO and 4E-BP prevent both

the cellular degenerative events and the functional decay of

aging muscles.

Epidemiological studies in humans have associated muscle

senescence with increased mortality (Nair, 2005), implying that

muscle aging may have organism-wide consequences beyond

muscle function. To ask whether the prevention of muscle aging

affects the organism life span, we manipulated the activity of

components of the Akt pathway in muscles and scored for their

effects on viability. As shown in Figures 4C and 4D, either Pten,

foxo, or 4E-BP CA overexpression in muscles is sufficient to

significantly extend longevity by increasing the median and

maximum life span. 4E-BP increased life span also in foxo

heterozygous null animals (Figure 4D), while Hsp70 overexpres-

sion on the other hand showed little effects (Figure S5). All

together, these findings indicate that the extent of muscle aging

is interconnected with the life span of the organism.

FOXO/4E-BP Signaling in Muscles InfluencesFeeding Behavior and the Release of Insulinfrom Producing CellsConsidering that both fasting and FOXO induce 4E-BP expres-

sion (Figure 2H), we wondered whether the systemic effect of

FOXO signaling on life span extension can result, at least in

part, from reduced food intake.

To test this hypothesis, we examined whether feeding

behavior would be decreased in adults with FOXO and 4E-BP

activation in muscles. We first monitored the amount of

liquid food ingested using the CAFE assay (capillary feeding)

(Ja et al., 2007). Interestingly, feeding was decreased in

response to FOXO/4E-BP signaling in muscles (Figure 5A). To

substantiate this finding, we measured the ingestion of blue-

colored food (Xu et al., 2008) and detected significant differ-

ences in food intake with this assay (Figure 5B), confirming


that feeding behavior is affected. Next, to assess whether

decreased feeding behavior arises from developmental defects,

we measured the body weight of adult flies, which is a sensitive

indicator of developmental feeding (Demontis and Perrimon,

2009), but found no significant differences (Figure 5C). Thus,

the behavior of flies overexpressing foxo and 4E-BP CA in

muscles most likely is not caused by developmental defects.

To assess the metabolic status, we monitored the glucose

concentration (glycemia) in the hemolymph. Similar to wild-

type flies starved for 24 hr, we detected a significant decrease

Figure 4. FOXO/4E-BP Signaling Preserves Muscle Function and Extends Life Span

(A) Muscle function gradually decreases during aging as indicated by an increase in the percentage of flies with climbing and flight defects. However, foxo

preserves their function in comparison with controls (flight ability: n[flies] = 10 (week 1 and 5) and 30 (week 8) with n[batch] = 3 (week 1 and 5) and 2 (week 8);

standard deviation (SD) is indicated and *p < 0.05. Climbing ability: (n[Mhc-Gal4/+] = 1264, n[Mhc-Gal4/UAS-foxo] = 966, with n indicating the number of flies

at day 1; p < 0.001).

(B) Similar to FOXO, 4E-BP activity also results in decreased age-related flight and climbing deficits in comparison with controls (flight ability: n[flies] R 10 (week 1

and 5) and 25 (week 8) with n[batch]R 3 (week 1 and 5) and 2 (week 8); SD is indicated and *p < 0.05. Climbing ability: (n[Mhc-Gal4/+] = 204, n[Mhc-Gal4/UAS-4E-

BP CA] = 403, p < 0.001).

(C) Survival of flies during aging. Foxo overexpression in muscles significantly extends the median and maximum life span (median and maximum life span:

Mhc-Gal4/+ = �61 and 82 days (n = 1264); UAS-foxo tr.#1/+;Mhc-Gal4/+ = �73 and 100 days (n = 1184); Mhc-Gal4/UAS-foxo tr.#2 = �76 and 94 days

(n = 966); p < 0.001).

(D) Life span of flies with increased Pten and 4E-BP activity in muscles is extended in comparison with matched controls (median and maximum life span of

4E-BP: Mhc-Gal4/+ = �63 and 78 days (n = 204); Mhc-Gal4/UAS-4E-BP CA = �71 and 84 days (n = 403); Pten: Mhc-Gal4/+ = �55 and 76 days (n = 162);

Mhc-Gal4/UAS-Pten = �66 and 88 days (n = 130); p < 0.001). Similar increase in life span is brought about by 4E-BP CA overexpression in foxo21 heterozygous

null flies.


Figure 3. FOXO and 4E-BP Regulate Proteostasis at Least in Part via the Autophagy/Lysosome System

(A–E) Immunostaining of muscles expressing the marker of autophagosomes Atg5-GFP reveals a significant increase in their number (E) and maintenance at

1 and 5 weeks of age upon foxo overexpression (C and D) in comparison with controls (A and B). In (E), SEM is indicated with n; *p < 0.05 and **p < 0.01.

(F–N) Immunostaining of muscles expressing the lysosomal marker Lamp1-GFP and overexpressing either Atg1, foxo, or 4E-BP CA. Note an increase in the

number of lysosomes (N) at both 1 (G-I) and 5 weeks of age (K–M), which inversely correlates with polyubiquitin immunoreactivity in comparison with control

muscles (F and J). Scale bar is 10 mm (A–D and F-–M). In (N), SEM is indicated with n; *p < 0.05 and ***p < 0.001.

(O) Relative mRNA levels of autophagy genes from thoraces of 1- and 5-week-old flies decrease during normal muscle aging, while their expression increases and

persists in response to FOXO. SEM is indicated with n = 4; *p < 0.05, **p < 0.01 and ***p < 0.001.

(P) RNAi treatment against Atg7 results in a �50% knockdown of its mRNA levels in muscles and partially impairs FOXO-mediated proteostasis, as indicated by

the increased detection of ubiquitin-conjugated proteins in Triton X-100 insoluble fractions at 8 weeks (old, red) in comparison with mock-treated (white RNAi) and

young flies (1 week old, black). Normalized values based on a-tubulin levels are indicated.


of glycemia in flies with FOXO and 4E-BP activation in muscles

(Figure 5D). All together, these findings suggest that FOXO and

4E-BP act as a metabolic brake in muscles that, by influencing

Figure 5. FOXO Signaling in Muscles Partially

Mimics Systemic Metabolic Changes Associated

with Fasting by Modulating Feeding Behavior

(A–C) Flies in which FOXO/4E-BP activity has been altered

specifically in muscles consume less food than matched

controls. Food consumption was determined via capillary

feeding CAFE assay over 2 hr periods (A), and by moni-

toring the ingestion of blue colored food in 24 hr (B). Error

bars represent SEM with n[measurements] = 44, 46, 52,

37, 103, and 61 in (A) and n = 2 in (B), with *p < 0.05,

**p < 0.01, ***p < 0.001. Decreased feeding does not result

from developmental defects, as indicated by similar body

weights of flies analyzed (C) (error bars represent SD with

n R 3).

(D) Relative glucose levels (glycemia) in the hemolymph of

flies overexpressing either foxo or 4E-BP CA in muscles,

and matched controls. Manipulation of FOXO/4E-BP

signaling in muscles brings about a reduction of glycemia

similar in part to that of wild-type flies starved for 24 hr, as

estimated with the glucose hexokinase assay (SEM is

indicated with n = 5, and **p < 0.01, ***p < 0.001).

(E–H) Immunostaining of Dilp-producing median neurose-

cretory cells in the brain of starved wild-type flies, flies

overexpressing foxo in muscles, and controls. Increase

in the immunoreactivity of the insulin-like peptide Dilp2

(green) is detected in producing cells in response to either

starvation (F) or foxo overexpression in muscles (H), in

comparison respectively with fed wild-type flies (E) and

controls with no foxo overexpression in muscles (G).

Smaller changes in Dilp5 levels are observed. Phalloidin

staining (blue) detects F-actin (scale bar is 20 mm; images

in [E]–[H] have the same magnification).

(I) Quantification of the intensity of staining indicates

that differences in Dilp2 fluorescence are significant

(SD is indicated with n[measurements] = 35, 69, 37, and

96 from n[brains] = 2, 4, 3, and 4; *p < 0.05).

(J–L) Quantification and immunostaining of adipose tissue

(peripheral fat body of the abdomen) from 2 week old flies.

(J) Note a significant increase in nuclear b-galactosidase

immunoreactivity (red) in the adipose tissue from flies

with a nuclear 4E-BP-lacZ reporter and foxo overexpres-

sion in muscles (L) in comparison with controls (K). F-actin

(green) and DAPI staining (indicative of nuclei, blue) are

shown. Scale bar is 20 mm. In (J), SEM is indicated with

n = 20 and ***p < 0.001.

feeding behavior, mimic at least in part the

physiological changes that are associated with

fasting.

To gain mechanistic insights into the systemic

regulation of aging by FOXO/4E-BP signaling

in muscles, we next monitored the release of

insulin-like peptides (Dilps) from the Dilp-

producing median neurosecretory cells in the

brain, which have been previously shown to

mediate the response of life span to nutrition

in Drosophila (Broughton et al., 2010). We

detected a significant accumulation of the

insulin-like peptide Dilp2 (and to a lesser extent,

Dilp5) in starved wild-type flies in comparison with fed flies

(Figures 5E and 5F). Increased immunoreactivity indicates

decreased release of Dilps and has been previously shown to


occur in response to starvation (Geminard et al., 2009). Next, we

tested whether similar changes would occur upon FOXO

signaling in muscles and found a partial accumulation of Dilps

(Figures 5G–5I).

Assuming that decreased Dilps secretion may result in

systemic FOXO activation, we monitored its activity using a

nuclear 4E-BP-lacZ transcriptional reporter. By immunostaining

adipose tissues with anti-b-galactosidase antibodies, we

detected higher 4E-BP expression upon foxo activation in

muscles in comparison with controls (Figures 5J–5L). Thus,

FOXO signaling in muscles appears to systemically activate

4E-BP expression in other tissues by regulating food intake

and insulin release.

FOXO/4E-BP Signaling in Muscles RegulatesProteostasis in Other Aging TissuesOur demonstration that FOXO/4E-BP signaling in muscles

extends life span in Drosophila and induces a systemic fasting-

like response, along with the observation that muscles undergo

age-related structural and functional changes precociously in

comparison with other tissues (Herndon et al., 2002; Zheng

et al., 2005), raises the possibility that muscle senescence may

influence the progression of age-related degenerative events in

the entire organism.

To test this hypothesis, we examined whether, in addition to

life span extension, FOXO signaling in muscles can affect protein

homeostasis in other tissues. As in the case of muscles (Figure 1

and Figure 2), we found that Ref(2)P/polyubiquitin aggregates

progressively accumulate in aging retinas (Figures 6A and 6D),

brains (Figures 6B and 6E), and adipose tissue (Figures 6C and

6F) (peripheral fat body of the abdomen). However, foxo overex-

pression in muscle resulted in decreased accumulation of

protein aggregates in other aging tissues (Figures 6D–6F; quan-

tification in Figure 6G). Similar changes were observed in

response to 4E-BP activity in muscles in comparison with

syngenic controls (Figure 6H). Importantly, this regulation is

muscle nonautonomous, as Mhc-Gal4 drives transgene expres-

sion only in muscles (and not in the retina, brain or adipose

tissue) (Figure S1). To further test the finding that FOXO/4E-BP

signaling in muscles delays the systemic impairment of proteo-

stasis in other tissues (Figures 6A–6H), we analyzed by western

blot the ubiquitin levels of Triton X-100 insoluble fractions, which

included protein aggregates, from either thoraces (which mainly

consist of foxo-overexpressing muscles) or heads and abdo-

mens (which are enriched in nonmuscle tissues and muscles

with little foxo overexpression) (Figure S1), at 1 and 8 weeks of

age. In agreement with the increased deposition of protein

aggregates observed during aging by immunofluorescence

(Figure 1, Figure 2, and Figures 6A–6F), ubiquitin levels were

dramatically increased in the Triton X-100 insoluble fractions

from control thoraces, and head and abdominal extracts at

8 weeks of age, in comparison with 1 week of age (Figure 6I).

However, ubiquitin levels were only partially increased in old

foxo-overexpressing flies in both thoracic and head and abdom-

inal extracts. No substantial differences were instead detected in

the Triton X-100 soluble fractions (data not shown). Similar

results were obtained by 4E-BP CA but not Hsp70 overexpres-

sion in muscles (Figure 6I; Figure S5), indicating that 4E-BP

activity in muscles also confers systemic protection from the

age-related decline in proteostasis. To test whether this effect

is muscle-specific, we overexpressed foxo in the adipose tissue

(abdominal fat body) with the S106GS-Gal4 driver, and analyzed

the deposition of polyubiquitinated proteins in Triton X-100

insoluble fractions from thoraces. Under these conditions, we

seemingly detected no differences (Figure S6), suggesting that,

although other tissues may be involved, muscles may play a

key role in this regulation. Altogether, these observations sug-

gest that FOXO and 4E-BP activity in muscles mitigates the

loss of proteostasis nonautonomously by influencing feeding

behavior, insulin release from producing cells, and 4E-BP activity

in other tissues.

DISCUSSION

By using a number of behavioral, genetic, and molecular assays,

we have described a mechanism in the pathogenesis of muscle

aging that is based on the loss of protein homeostasis (proteo-

stasis) and the resulting decrease in muscle strength (Figure 7).

Increased activity of Pten and the transcription factor FOXO is

sufficient to delay this process, while foxo null animals experi-

ence accelerated loss of proteostasis during muscle aging.

Pten and FOXO induce multiple protective responses, including

the expression of folding chaperones and the regulator of protein

translation 4E-BP that has a pivotal role in preserving proteosta-

sis. FOXO and 4E-BP preserve muscle function, at least in part

by sustaining the basal activity of the autophagy/lysosome

system, which removes aggregates of damaged proteins.

However, additional mechanisms may be involved. For example,

the proteasome system may degrade damaged proteins and

thus avoid their accumulation in aggregates (Rubinsztein,

2006). Thus, perturbation in proteasome assembly and subunit

composition may contribute to muscle aging in response to

FOXO activity. In addition, whereas overexpression of a single

chaperone had limited effects, interventions to effectively limit

the extent of protein damage are likely to delay the decay in

proteostasis by decreasing the workload for the proteasome

and autophagy systems (Tower, 2009).

By comparing the accumulation of polyubiquitinated proteins

in aggregates of aging muscles, retinas, brains, and adipose

tissue, we have found that reduced protein homeostasis is a

general feature of tissue aging that is particularly prominent in

muscles (Figure 1, Figure 6, and Figure S6). The observation

that muscle aging is characterized by loss of proteostasis further

suggests some similarity between muscle aging and neurode-

generative diseases, many of which are characterized by the

accumulation of protein aggregates (Rubinsztein, 2006).

Mechanical, thermal, and oxidative stressors occur during

muscle contraction (Arndt et al., 2010), and therefore muscle

proteins may be particularly susceptible to damage in compar-

ison with other tissues. While our findings refer to the loss of

proteostasis in the context of normal aging, it is likely that a better

understanding of this process will help cure muscle pathologies

associated with aging, as some of the underlying mechanisms of

etiology may be shared. For example, most cases of inclusion

body myositis (IBM) arise over the age of 50 years, defining

aging as a major risk factor for the pathogenesis of this disease.


Figure 6. Systemic Proteostasis Is Remotely Controlled by FOXO/4E-BP Signaling in Muscles

(A–F) Aggregates of polyubiquitinated proteins accumulate during aging in the retina (A and D), brain (B and E), and the adipose tissue (C and F) of control flies

(Mhc-Gal4/+), but to a lesser extent in tissues from flies overexpressing foxo in muscles (UAS-foxo/+;Mhc-Gal4/+), as indicated by polyubiquitin (red) and p62/Ref

(2)P (green) stainings. Phalloidin staining (blue) outlines F-actin. Note that Mhc-Gal4 does not drive transgene expression in these tissues (Figure S1). Scale bar is

10 mm.

(G and H) The age-related increase in the cumulative area of protein aggregates is significantly less prominent in tissues from flies overexpressing foxo (G) or

4E-BP CA (H) in muscles in comparison with controls (SEM is indicated with n; *p < 0.05. **p < 0.01, and ***p < 0.001).

(I) Ubiquitin levels (indicative of protein aggregates) are detected in Triton X-100 insoluble fractions from thoraces, and head and abdominal tissues from flies

overexpressing foxo in muscles or control flies at 1 (young, black) and 8 (old, red) weeks of age. Ubiquitin levels are increased in old flies in comparison with young


Interestingly, muscle weakness in patients with IBM is character-

ized by the accumulation of protein aggregates (Needham and

Mastaglia, 2008), which we have now described as occurring

in the context of regular muscle aging in Drosophila. Thus,

FOXO may interfere with the pathogenesis of muscle degenera-

tive diseases in addition to muscle aging. Studies in animal

disease models of IBM will be needed to test this hypothesis.

There is an apparent contradiction between our findings and

the data describing the FOXO-dependent induction of muscle

atrophy in mice (Bodine et al., 2001; Sandri et al., 2004), a serious

form of muscle degeneration that results in decreased muscle

strength (Augustin and Partridge, 2009). The observation that

different degrees of FOXO activation can promote stress resis-

tance, or rather cell death (Salih and Brunet, 2008), could explain

why FOXO activity can be protective or rather detrimental during

muscle aging. In particular, while physiologic FOXO activation

can preserve protein homeostasis and muscle function, its

excessive activation may lead to decreased muscle function

due to hyperactivation of the protein turnover pathways. Consis-

tent with this view, the autophagy pathway has also been

involved in both muscle atrophy (Mammucari et al., 2007;

Zhao et al., 2007) and in the preservation of muscle sarcomere

organization (Arndt et al., 2010; Masiero et al., 2009), high-

lighting the importance of fine-tuning the degree of activation

of stress resistance pathways to maintain muscle homeostasis.

In addition, the output of FOXO activity may radically differ in

growing versus preexisting myofibers. In particular, our present

study indicates that FOXO protects preexisting myofibers

Figure 7. FOXO/4E-BP Signaling in Muscles

Controls Proteostasis and Systemic Aging

Muscle aging is characterized by protein damage

and accumulation of cytoplasmic aggregates.

Loss of protein homeostasis (proteostasis) associ-

ates with the progressive decrease in muscle

strength and can affect the life span of the

organism. Pten/FOXO signaling induces multiple

targets including several folding chaperones

and the regulator of protein translation 4E-BP.

FOXO/4E-BP activity regulates muscle proteosta-

sis at least in part via the autophagy/lysosome

pathway of protein degradation, preserves muscle

function, and extends life span. In addition, FOXO/

4E-BP signaling in muscles decreases feeding

behavior that, similar to fasting, results in reduced

insulin release from producing cells. This in turn

promotes FOXO and 4E-BP activity in other

tissues, preserving proteostasis organism-wide

and mitigating systemic aging.

flies in extracts from both muscles (thoraces) and nonmuscle tissues (heads and abdomens). However, flies overexpressing foxo in muscles have reduced

deposition of protein aggregates at 8 weeks of age in both muscles and nonmuscle tissues. Similar results are obtained in response to increased 4E-BP activity

in muscles (I), but not Hsp70 (Figure S5). Quantification of ubiquitin-conjugated proteins normalized to a-tubulin or histone H3 levels is indicated.

See also Figures S1, Figure S5, and Figure S6.

against age-dependent changes in pro-

teostasis while also blunting develop-

mental muscle growth in flies (Demontis

and Perrimon, 2009), as observed in

mammals (Kamei et al., 2004). Thus,

deleterious effects of FOXO activation as observed in mamma-

lian muscles may result from the inhibition of the growth of novel

myofibers in postnatal development and adulthood, a process

which is thought to be limited to development in Drosophila

(Grefte et al., 2007).

An interesting observation of our study is that interventions

that decrease muscle aging also extend the life span of the

organism. In particular, our work raises the prospect that the

extent of muscle aging may be a key determinant of systemic

aging (Figure 7). Reduced muscle proteostasis may be detri-

mental per se for life expectancy, presumably due to the involve-

ment of muscles in a number of key physiological functions.

Consistent with this view, overexpression in muscles of aggrega-

tion-prone human Huntington’s disease proteins is sufficient to

decrease life span (Figure S7). Moreover, FOXO signaling in

muscles regulates proteostasis in other tissues, via the inhibition

of feeding behavior and the decreased release of insulin from

producing cells, which in turn promote 4E-BP activity systemi-

cally. Thus, we propose that FOXO/4E-BP signaling in muscles

regulates life span and remotely controls aging events in other

tissues by bringing about some of the protection associated

with decreased food intake.

In mammals, muscles produce a number of cytokines involved

in the control of systemic metabolism (Nair, 2005; Pedersen and

Febbraio, 2008). For example, interleukin-6 (IL-6) is produced by

muscles and has been proposed to control glucose homeostasis

and feeding behavior through peripheral and brain mechanisms

(Febbraio and Pedersen, 2002; Plata-Salaman, 1998). Thus,


a muscle-based network of systemic aging as observed in flies

may occur in humans.

This study supports the common belief that preserving muscle

function is beneficial for overall aging (Boyle et al., 2009; Chen

et al., 2005), and the notion that muscles are central tissues

to coordinate organism-wide processes, including aging and

metabolic homeostasis (Nair, 2005). Moreover, the observation

that FOXO signaling in muscles influences aging events in other

tissues suggests that the systemic regulation of aging relies on

tissue-to-tissue communication (Russell and Kahn, 2007), which

may provide the basis for interventions to extend healthy life

span.


Drosophila Strains and Life Span Analysis

Details on fly strains can be found in Extended Experimental Procedures.

For longevity measurement, male flies were collected within 24 hr from

eclosion and reared at standard density (20 flies per vial) on cornmeal/soy

flour/yeast fly food at 25�C. Dead flies were counted every other day and

food changed. For each genotype, at least two independent cohorts of flies,

raised at different times from independent crosses, were analyzed. For starva-

tion treatments, flies were kept in normal vials with 1.5% agar as a water

source for the period of time indicated. For all experiments, Mhc-Gal4 females

were mated with male transgenic and syngenic control flies, and the resulting

male offspring analyzed in parallel by comparing transgene expressing

flies with matched controls flies having the same genetic background. For

transgene expression with the Gal4-UAS system, flies were reared at 25�C.

Behavioral and Metabolic Assays

Flight ability was scored according to Park et al. (2006), and negative geotaxis

assays were performed as previously described (Rhodenizer et al., 2008). In

brief, flies were gently tapped to the bottom of a plastic vial, and the number

of flies that could climb to the top of the vial after 20 s was scored. Quantifica-

tion of the glucose concentration in the hemolymph, and capillary (CAFE) and

blue-colored food feeding assays were done as previously described

(Geminard et al., 2009; Xu et al., 2008) and are described in detail in Extended


Immunostaining, Confocal and Electron Microscopy,

and Image Analysis

For whole-mount immunostaining of the fly tissues, indirect flight muscles, and

peripheral fat body of the abdomen, retinas, and brains were dissected from

male flies and fixed for 30–40 min in PBS with 4% paraformaldehyde and

0.2% Triton X-100. After washing, samples were incubated overnight with

appropriate primary and secondary antibodies. Image analysis was done

with ImageJ and Photoshop. Immuno-gold electron microscopy was done

similar to Nezis et al., (2008). See Extended Experimental Procedures for

further information and a list of the antibodies used.

Quantitative Real-Time RT-PCR

qRT-PCR was done as previously described (Demontis and Perrimon, 2009).

Total RNA was prepared from fly thoraces and qRT-PCR was performed

with the QuantiTect SYBR Green PCR kit (QIAGEN). Alpha-Tubulin 84B was

used as normalization reference. Relative quantification of mRNA levels was

calculated using the comparative CT method.

Statistical Analysis

Statistical analysis was performed with Excel (Microsoft) and p values were

calculated with Student’s t tests and log-rank tests.

Western Blot and Biochemical Analysis of Detergent-Insoluble

Fractions

Western blot and biochemical analysis of detergent-insoluble fractions were

done substantially as previously described (Nezis et al., 2008). In brief,

dissected flies were homogenized in ice-cold PBS with 1% Triton X-100 and

protease inhibitors, and the resulting unsoluble pellet resuspended in RIPA

buffer with 5% SDS and 8M urea. See Extended Experimental Procedures

for a complete protocol.



seven figures, and two tables and can be found with this article online at

doi:10.1016/j.cell.2010.10.007.

ACKNOWLEDGMENTS

We are grateful to Andreas Brech, Didier Contamine, Ernst Hafen, Pierre

Leopold, Susan Lindquist, Ioannis Nezis, Amita Sehgal, Marc Tatar, Robert

Tjian, John Tower, the DRSC/TRiP, and members of the Perrimon lab for fly

stocks, reagents, and advice. We thank Maria Ericsson for assistance with

electron microscopy, Christians Villalta for embryo injection, and Chris Bakal,

Rami Rahal, and Jonathan Zirin for critically reading the manuscript. This work

was supported by the NIH (1P01CA120964-01A1) and a Pilot Project Grant

from the Paul F. Glenn Labs for the Molecular Biology of Aging. F.D. is an

Ellison Medical Foundation/AFAR postdoctoral fellow. N.P. is an investigator

of the Howard Hughes Medical Institute.

Received: February 3, 2010

Revised: June 24, 2010



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Reelin and Stk25 Have OpposingRoles in Neuronal Polarizationand Dendritic Golgi DeploymentTohru Matsuki,1 Russell T. Matthews,1 Jonathan A. Cooper,3 Marcel P. van der Brug,2,4 Mark R. Cookson,2

John A. Hardy,2,5 Eric C. Olson,1 and Brian W. Howell1,*1Department of Neuroscience and Physiology, SUNY Upstate Medical University, Syracuse, NY 13210, USA2Laboratory of Neurogenetics, National Institute on Aging, National Institutes of Health, Bethesda, MD 20892, USA3Division of Basic Sciences, Fred Hutchinson Cancer Research Center, Seattle, WA 98109, USA4Present address: Department of Neuroscience, The Scripps Research Institute, Jupiter, FL 33458, USA5Present address: Department of Molecular Neuroscience and Reta Lila Weston Laboratories, University College, Queens Square House,

London WC1 3BG, UK*Correspondence: [email protected]

DOI 10.1016/j.cell.2010.10.029

SUMMARY

The Reelin ligand regulates a Dab1-dependent sig-naling pathway required for brain lamination andnormal dendritogenesis, but the specific mecha-nisms underlying these actions remain unclear. Wefind that Stk25, a modifier of Reelin-Dab1 signaling,regulates Golgi morphology and neuronal polariza-tion as part of an LKB1-Stk25-Golgi matrix protein130 (GM130) signaling pathway. Overexpression ofStk25 induces Golgi condensation and multipleaxons, both of which are rescued by Reelin treat-ment. Reelin stimulation of cultured neurons inducesthe extension of the Golgi into dendrites, which issuppressed by Stk25 overexpression. In vivo, Reelinand Dab1 are required for the normal extension of theGolgi apparatus into the apical dendrites of hippo-campal and neocortical pyramidal neurons. Thisdemonstrates that the balance between Reelin-Dab1 signaling and LKB1-Stk25-GM130 regulatesGolgi dispersion, axon specification, and dendritegrowth and provides insights into the importance ofthe Golgi apparatus for cell polarization.

INTRODUCTION

The development of the exquisite morphology of neurons is

a carefully orchestrated process that optimizes the ability of indi-

vidual neurons to receive signals, integrate them, and transmit

the output to target cells. Neuronal polarization, first observed

as the rapid growth of a process that will ultimately become an

axon, followed by the asymmetrical development of dendrites

are key steps in morphological and functional maturation

(Arimura and Kaibuchi, 2005). Interestingly, the Golgi apparatus

has been implicated in these different aspects of neuronal

polarity. In the nascent neuron, the position of the Golgi and

the adjoined centrosome correlates with the site of axon emer-

gence, which becomes the future basal side of a mature pyra-

midal neuron (de Anda et al., 2005, 2010; Zmuda and Rivas,

1998). Later, the Golgi apparatus is positioned on the apical

side of pyramidal neurons, proximal to the major apical dendritic

tree and opposite to the axon and minor basal dendrites (Horton

et al., 2005). Dispersion of the Golgi apparatus away from

the apical pole leads to a loss of dendrite asymmetry in these

cells, with equal-sized apical and basal dendrites (Horton et al.,

2005). Furthermore, specialized Golgi outposts, which populate

dendrites, promote the elaboration of dendritic branches (Ye

et al., 2007). However, it remains to be determined how Golgi

positioning within neurons is regulated.

Mutations in the genes encoding the Reelin-Dab1 signaling

pathway lead to profound defects in neuronal positioning and

dendritogenesis during brain development (Niu et al., 2004;

Rice et al., 2001). The lamination of the cerebral cortex, hippo-

campus, and cerebellum is disorganized and appears approxi-

mately inverted compared to normal. Reelin is a secreted ligand

that is produced in discreet layers in the developing brain

(D’Arcangelo et al., 1995; Ogawa et al., 1995). Genetic and

biochemical studies have shown that it regulates a signal trans-

duction pathway requiring the ApoE receptors ApoER2 and

VLDLR (D’Arcangelo et al., 1999; Hiesberger et al., 1999;

Trommsdorff et al., 1999), the cytoplasmic adaptor protein

Dab1 (Howell et al., 2000), and Src family kinases (Arnaud

et al., 2003; Bock and Herz, 2003). Disparate functions have

been proposed for Reelin-Dab1 signaling, though a clear biolog-

ical response to clarify its role in brain development is lacking

(Chai et al., 2009; Cooper, 2008; Forster et al., 2010; Sanada

et al., 2004).

The severity of dab1-dependent phenotypes depends on the

genetic background (Brich et al., 2003). We have recently identi-

fied stk25 as a modifier of dab1 mutant phenotypes (unpublished

data). Here we characterize the role of Stk25 (also YSK1, Sok1) in

nervous system development. Previous work has implicated

Stk25 in regulating Golgi morphology through the Golgi matrix

protein GM130 (Preisinger et al., 2004), which we confirm here.


GM130 regulates the fusion of ER-to-Golgi vesicles with the

Golgi cisternae and the fusion of Golgi cisternae into elongated

ribbons (Barr and Short, 2003; Puthenveedu et al., 2006). Deple-

tion or mitotic phosphorylation of GM130 leads to Golgi frag-

mentation and reduced efficiency of biosynthetic processing

(Lowe et al., 1998; Marra et al., 2007; Puthenveedu et al., 2006).

The protein kinase LKB1 and its associated factors STRAD

and MO25 are known to be important for neuronal polarization,

axon specification, and dendrite growth (Asada et al., 2007;

Barnes et al., 2007; Shelly et al., 2007). In this study, we find

that Stk25 is part of an LKB1 cell polarization pathway. Stk25,

LKB1, and GM130 are shown to regulate Golgi morphology

and axon initiation. In addition, we show that Stk25 and Reelin-

Dab1 signaling have antagonistic effects on neuronal polariza-

tion and the morphology and subcellular distribution of the

Golgi. As the position of the Golgi plays roles in cell polarization,

process extension, and cell migration (Fidalgo et al., 2010;

Horton et al., 2005; Yadav et al., 2009; Ye et al., 2007), this

evidence is fundamental for understanding the molecular control

of neuronal morphogenesis and provides new insights into the

biological role of Reelin-Dab1 signaling.

RESULTS

Stk25 Regulates Neuronal PolarityStk25 has previously been shown to regulate the polarized

migration of epithelial cells. As other Ste20-like kinases have

roles in neuronal polarization (Jacobs et al., 2007; Preisinger

et al., 2004), we sought to assess a role for Stk25 in neuronal

polarization by using hippocampal neuronal cultures (Dotti and

Banker, 1987). These neurons have a stereotypic morphology

and program of differentiation and respond to Reelin-Dab1

signaling (Matsuki et al., 2008). Soon after plating, they extend

short uniform processes that have the potential to develop

into either axons or dendrites (Arimura and Kaibuchi, 2007). By

stage III, 48 to 72 hr later, one of the processes can be identified

as an axon whereas the other processes differentiate into

dendrites.

We reduced Stk25 levels by infection with a lentivirus carrying

GFP and Stk25 shRNA and identified axons 6 days later using

SMI-312, a pan-axonal neurofilament marker. Depletion of

Stk25 inhibited axon specification. At least 30% of the Stk25

shRNA lentivirus-infected, GFP-positive neurons lacked an

axon (Figures 1B and 1F, lane 2), whereas axons were detected

in all neurons infected with either empty vector (EV) or control

shRNA vectors (Figures 1A and 1F, lanes 1 and 3 and insets).

The longest process in Stk25 shRNA-expressing cells was also

much shorter than the long axons of control cells (Figures 1A,

1B, and 1F, lane 2), consistent with a failure to induce an axon.

To assess whether axon absence was specifically caused by

reduced Stk25 expression, we tested for rescue by Stk25 over-

expression. Both kinase-active and kinase-inactive versions of

an shRNA-resistant Stk25 (Stk25*) were expressed as red fluo-

rescent protein (RFP) fusion proteins in cultures that were also

infected with the GFP-expressing, Stk25 shRNA virus (Figures

S1A–S1D available online). Both kinase-active and kinase-inac-

tive Stk25*-RFP rescued the axon-less phenotype caused by

Stk25 knockdown (Figure 1F, lanes 7–9). This suggests that

the axon-less phenotype in Stk25 shRNA-expressing cells was

the specific result of reducing Stk25 expression and that Stk25

kinase activity is not required for axon production.

To investigate whether Stk25 affected axon initiation or main-

tenance, we examined stage III hippocampal neurons (Figures

1D and 1E). We found that 56% ± 5% of Stk25 knockdown

neurons lacked an axon compared to only 7% ± 8% of control

samples (Figure 1G). The longest neurite in Stk25 knockdown

neurons was also significantly shorter than the incipient axon in

control cultures. Moreover, overexpression of Stk25 induced

multiple axons. Expression of either the wild-type or kinase-inac-

tive Stk25*-RFP fusion proteins, or an Stk25-green fluorescent

protein (GFP) fusion that has previously been shown to be bio-

logically active (Preisinger et al., 2004), induced multiple SMI-

positive axons in approximately 45%–50% of neurons as

compared to 15% ± 3% in GFP-alone expressing controls

(Figures 1C and 1F, lanes 5, 6, 8, and 9). Stk25 overexpression

did not increase axon length (Figure 1F). Taken together, the

results show that Stk25 regulates axon initiation but not axon

growth in cultured neurons.

Reelin-Dab1 Signaling Suppresses Multiple AxonProductionStk25 is expressed at relatively high levels in Reelin-Dab1

responsive cells in the developing cortical plate (Figure S1E)

and in the adult hippocampus and cerebellar Purkinje cells (Fig-

ure S1F). Because we identified stk25 in a screen for modifiers of

dab1 mutant phenotypes (unpublished data), we examined

whether Reelin-Dab1 signaling might have an undiscovered

role in axon initiation. Hippocampal neurons were cultured

from dab1�/� mutant embryos and infected with GFP-express-

ing lentiviruses to survey their morphology. Surprisingly, approx-

imately 30% of the dab1�/� mutant neurons produced multiple

axons as compared to approximately 15% of the wild-type

neurons (Figure 1H). To determine whether the multiple axon

phenotype in dab1�/� mutant neurons was sensitive to Stk25

expression level, we examined the effect of knocking down

Stk25. Significantly fewer dab1�/� mutant neurons infected

with the Stk25 shRNA-expressing lentivirus produced multiple

axons than the GFP-expressing control sample (Figure 1H). In

addition, a significant number of the Stk25 shRNA-expressing

neurons completely lacked axons. This shows that Reelin-

Dab1 signaling regulates axon initiation and that the multiple

axon phenotype in dab1�/� mutant mice is dependent upon

Stk25 expression.

Congruent with this result, growth of neurons in the presence

of Reelin suppressed the multiple axon phenotype caused

by Stk25 overexpression (Figure 1I). This treatment did not,

however, lead to the loss of axon production, which would be ex-

pected if Stk25 function was abolished. None of these treat-

ments affected axon length. Therefore, Reelin-Dab1 signaling

appears to counteract the effects of high Stk25 expression

without completely blocking its function in axon induction.

Stk25 Regulates Axon Formation and DendriteAsymmetry In VivoTo investigate whether Stk25 regulates neuronal differentiation

in vivo, we electroporated the Stk25 shRNA-expressing vector


into the hippocampi of fetal mice. The brains of these mice were

analyzed for GFP expression and neuronal polarization of Ctip2-

positive, pyramidal neurons in the CA1 region of the hippo-

campus at postnatal day 7 (P7). Stk25 shRNA did not interfere

with the positioning of neurons, but their apical dendrites were

Figure 1. Stk25 Expression Regulates Axon

Differentiation in Culture

(A) Primary hippocampal neurons (E17.5) infected

with the GFP-expressing EV-control virus had

typical pyramidal neuron morphologies, including

a long SMI-positive axon (inset a) and shorter

dendrites.

(B) Neurons infected with the Stk25 shRNA virus

had shorter processes and frequently lacked

long (>250 mm) SMI-positive processes that met

the criteria for axons (inset b). An SMI-positive

process (arrowhead) from a noninfected neuron

runs parallel to the GFP-positive process (arrow).

(C) Cells overexpressing Stk25 wild-type (WT)-

GFP had multiple SMI-positive axons (insets c, c0 ).(D) At stage III (2DIV), EV-control infected neurons

had one dominant SMI-positive axon.

(E) In contrast, Stk25 shRNA-expressing neurons

often lacked SMI-positive, axon-like processes.

(F) The number of neurons with 0, 1, 2, or more

axons and the length of the longest processes

were determined for neurons infected with

the indicated viruses. For rescue experiments,

neurons were coinfected with the Stk25 shRNA

(GFP-positive) and either RFP, Stk25* WT-RFP,

or Stk25* K49R-RFP expressing viruses (lanes

7–9, Figure S1).

(G) At stage III (2DIV), many Stk25 shRNA-

expressing neurons lacked axons as compared

to a small percentage of EV-control infected

neurons.

(H) The number of neurons with multiple axons

was increased in dab1�/� (lane 2) compared to

wild-type neurons (lane 1, duplicated from F),

and this was reduced by Stk25 shRNA expression

(lanes 3).

(I) Primary hippocampal neurons that were in-

fected with either GFP- or Stk25 WT-GFP-ex-

pressing viruses were split into three groups and

grown in either neurobasal (NB), control-condi-

tioned (CCM), or Reelin-conditioned (RCM) media

for 6 days.

Statistical significance (*,**,***p < 0.0001,

Student’s t test, compared between the sample

pairs: (F) 1:2; 4:5,6,7; 7:8,9; (G) 1:2; (H) 1:2, 2:3;

n > 60; (I) 5:6; n indicated in bars). Bars: (C)

50 mm; (a) 10 mm; (c0) 5 mm; and (E) 20 mm. See

also Figure S1.

significantly longer (Figures 2A, 2B, and

2E). In addition, approximately 40% of

the strongly GFP-positive, Stk25 shRNA-

expressing neurons lacked identifiable

axon initial segments, detected using

anti-phospho-IkBa antibodies, suggest-

ing that axons were either absent or failed

to mature normally (Figures 2D and 2F;

Movie S1). By comparison, all of the GFP-positive, EV-control

electroporated neurons examined had axon initial segments

(Figures 2C and 2F; Movie S1). This suggests that Stk25 regulates

axon specification and dendrite growth in hippocampal pyra-

midal neurons in vivo.


In addition to having longer apical dendrites, the basal

dendrites of Stk25 shRNA-expressing neurons were also atyp-

ical. Normal pyramidal neurons have long, thick apical dendrites

and much thinner and shorter basal dendrites (Horton et al.,

2005; Figures 2G and 2K; Movie S2). The apical dendrites of

Stk25 shRNA-expressing neurons had normal thickness, but

the basal dendrites were thicker than normal (Figures 2H and

2K; Movie S2). We were not able to measure the length of the

basal dendrites. Therefore, there is evidence for growth of both

apical and basal dendrites, and this reduced the distinction

between apical and basal dendrites in terms of thickness. This

suggests that Stk25 is needed for normal axon production and

dendrite asymmetry in vivo.

Stk25 Interacts with STRADa and Acts on the LKB1Signaling PathwayThe functions of Stk25 resemble those reported for LKB1-

STRAD signaling (Barnes et al., 2007; Kishi et al., 2005; Shelly

et al., 2007). This pathway has a prominent role in cell polarity

control across numerous cell types from Caenorhabditis elegans

to man. LKB1 is partially regulated by binding STRAD, which

both shuttles it from the nucleus to the cytoplasm and stabilizes

it. We therefore investigated whether Stk25 associates with the

LKB1-STRAD signaling complex. By immunoprecipitating

tagged fusion proteins coexpressed in HEK293T cells, we found

that both wild-type and kinase-inactive HA-Stk25 coimmuno-

precipitated with myc-STRADa (Figure S2A). Identifying Stk25

Figure 2. Stk25 Regulates Neuronal Polarity during Brain Development

(A) EV-control vector (GFP-positive, green) electroporated at E16.5 in utero was expressed in Ctip2-positive (red), hippocampal-pyramidal neurons at P7.

(B) Stk25 shRNA-expressing neurons (GFP-positive) were appropriately positioned in the CA1 layer, and their apical dendrites extended further than EV-control.

(C) GFP-expressing, EV-control transfected CA1 neurons had the typical pyramidal shape and phospho-IkBa- (red), GFP-positive (green) axon initial segments

(Sanchez-Ponce et al., 2008) (Movie S1).

(D) In contrast, a high percentage of strongly GFP-positive, Stk25 shRNA-expressing neurons were often misshapen and lacked axon initial segments (Movie S1).

(E) Quantification of apical dendrite length in EV-control and Stk25 shRNA hippocampi.

(F) Quantification of the number of GFP-, Ctip2-positive pyramidal neurons that had axon initial segments (n indicated in bar.)

(G) In EV-control neurons, the Golgi apparatus (trace of GRASP65 signal) is concentrated on the apical side of the neuron (Movie S2).

(H) In Stk25 shRNA-expressing neurons, the Golgi apparatus is broadly distributed throughout the neuron (Movie S2).

(I) Scheme used to determine Golgi distribution in (J).

(J) The Golgi distribution in apical, lateral (combined), or basal quadrants was quantified.

(K) The diameters of the largest apical and basal processes were determined (*p < 0.0005, Student’s t test, n R 12, neurons from three animals).

Bars: (B) 200 mm; (D and H) 10 mm. Error bars indicate standard error of the mean (SEM) in all figures.


as a direct or indirect STRAD-binding protein suggests a poten-

tial role for Stk25 on the LKB1 pathway.

To investigate whether Stk25 is important for LKB1 function,

we took two approaches. We examined whether (1) Stk25 is

required for LKB1-STRAD-regulated epithelial cell polarization

and (2) Stk25 overexpression rescues the LKB1 knockdown

phenotype in neurons.

We first tested whether reduced Stk25 expression would

inhibit the LKB1-STRAD-dependent polarization of W4 intestinal

epithelial cells. These cells have been engineered to constitu-

tively express LKB1 and express STRAD in response to doxycy-

line, which leads to their polarization (Baas et al., 2004). Most W4

cells infected with EV and control shRNA lentiviruses became

polarized within 24 hr of doxycycline treatment (Figures S2C

and S2E). In contrast, only 20% of cells infected by the human-

ized (h) Stk25 shRNA lentivirus were polarized by doxycycline

treatment (Figures S2C and S2E).

Furthermore, expression of either wild-type or kinase-inactive

Stk25*-RFP rescued STRAD-induced polarization in Stk25

shRNA-expressing W4 epithelial cells (Figure S2F). Collectively,

these experiments show that the Stk25 protein, not its kinase

activity, is required for LKB1-STRAD-regulated epithelial cell

polarization.

We then confirmed that LKB1 knockdown leads to a loss of

axon initiation in cultured hippocampal neurons (Figure 3A;

Barnes et al., 2007; Shelly et al., 2007). We tested whether

Stk25 can rescue or bypass the LKB1 requirement by overex-

pressing Stk25* wild-type (WT)-RFP in LKB1 shRNA-expressing

neurons (Figure 3B). Ninety-two percent of LKB1 knockdown

neurons that expressed Stk25* WT-RFP produced at least one

axon compared to only 48% of RFP-, LKB1 shRNA-coexpress-

ing neurons (Figure 3E). These results are consistent with a role

of Stk25 on the LKB1 pathway to regulate axon induction.

GM130 Interacts with Stk25 and Regulates AxonInductionThe Golgi matrix protein GM130, which has critical roles in regu-

lating Golgi dynamics, was identified in a yeast two-hybrid

screen as an Stk25 binding partner (Preisinger et al., 2004). We

confirmed this interaction by coimmunoprecipitating tagged

Figure 3. Stk25-RFP Overexpression Rescues the Neuronal Polarization Defect Caused by LKB1 but Not by GM130 Knockdown

(A) Expression of LKB1 shRNA (GFP-positive, green) in hippocampal neurons led to an increase in the number of neurons that lack an axon at 6DIV in cells also

expressing RFP (red). (a) Longest process lacks SMI immunoreactivity.

(B) In contrast, overexpressing Stk25* WT-RFP in LKB1 knockdown neurons rescued axon production. (b) Long, axon-like process is SMI positive.

(C) GM130 knockdown (GFP-positive) also caused a reduction in axon production in RFP-positive cells. (c) No SMI imunoreactivity was detected in processes of

the GFP-, RFP-positive neuron.

(D) Stk25* WT-RFP expression did not rescue axonogenesis in GM130 knockdown neurons. (d) Longest process is SMI negative.

(E) Axon number and the length of the longest processes were quantified for the indicated treatment groups. (Lane 1 was duplicated from Figure 1F lane 1.)

(*p < 0.005 compared to lane 1, **p = 0.01 compared to lane 2, Student’s t test.)

Bars: (D) 50 mm; (d) 5 mm. See also Figure S2.


fusions of GM130 and Stk25 (Figure S2B). Interestingly, kinase-

inactive Stk25 consistently immunoprecipitated with GM130

more efficiently than wild-type, suggesting that Stk25-depen-

dent phosphorylation may destabilize the complex.

Stk25 colocalizes with GM130 at the Golgi apparatus of HeLa

cells (Preisinger et al., 2004). To determine whether Stk25 local-

izes to the Golgi complex in neurons, we raised an antibody to

a region of Stk25 that is divergent from the close relatives

Mst3 and Mst4 (Extended Experimental Procedures). Endoge-

nous Stk25 expression overlapped with the GM130-positive

cis-Golgi in neurons at stage III, coincident with axon specifica-

tion (Figure S2D).

To asses whether GM130 plays a role in neuronal differentia-

tion, we examined GM130 shRNA-expressing neurons for

defects in polarity. Similar to Stk25 and LKB1 knockdown

neurons, knockdown of GM130 reduced axon number at 6DIV

(Figure 3C). GM130 knockdown also caused a significant reduc-

tion in axon initiation in stage III (2DIV) neurons (data not shown).

Stk25*-RFP overexpression in GM130-deficient cells did not

rescue axon number at 6DIV (Figure 3D), which suggests that

GM130 is required for neuronal polarization downstream of

Stk25.

Figure 4. Golgi Apparatus Morphology Is

Regulated by Stk25, LKB1, and GM130

Expression and Reelin Signaling

(A) Stage III neurons that were infected with the

EV-control virus had typical cis-Golgi ribbons

(GRASP65, Movie S3). In contrast, the cis-Golgi

in Stk25 shRNA-, LKB1 shRNA-, or GM130

shRNA-expressing neurons was fragmented

(Movie S3). GFP signal was omitted for clarity.

(B) Significantly more Stk25 knockdown neurons

had fragmented Golgi complexes compared to

the EV-control and the control shRNA (n, as indi-

cated). LKB1 and GM130 knockdown also

caused significant Golgi fragmentation as com-

pared to EV-control infected neurons. Stk25*-

RFP expression rescued Golgi fragmentation in

LKB1 shRNA but not GM130 shRNA-expressing

neurons.

(C) Neurons overexpressing either Stk25 WT-GFP

or Stk25 K49R-GFP had condensed cis-Golgi

(GRASP65 signal) compared to EV-controls when

grown in either neurobasal or control-CM. Growth

in Reelin-CM partially rescued the Golgi appear-

ance in Stk25-overexpressing cells. GM130 and

GRASP65 colocalized under all conditions (not

shown).

(D) Golgi volume (upper panel) and the length of

the longest Golgi ribbon (lower panel) were deter-

mined (*p < 0.0001, Student’s t test, n indicated in

bars).

Bars: 5 mm. See also Figure S3.

Stk25, GM130, and LKB1 RegulateGolgi DistributionPreviously it was shown that GM130

regulates Golgi morphology in HeLa cells

(Puthenveedu et al., 2006). Given that

Stk25, LKB1, and GM130 regulate axon

initiation, and the position of the Golgi apparatus early in differen-

tiation normally coincides with axonal localization (de Anda et al.,

2005, 2010), we examined whether Stk25, LKB1, and GM130

regulate Golgi morphology (Figure 4). Individually knocking

down Stk25, LKB1, and GM130 in stage III primary hippocampal

neurons resulted in dispersion of Golgi elements in a high

percentage of cells, in contrast to the typical elongated

morphology observed in the EV-control neurons (Figures 4A

and 4B; Movie S3).

Interestingly, the Golgi fragmentation caused by LKB1 knock-

down was rescued by Stk25*-RFP overexpression (Figure 4B),

suggesting that Stk25 overexpression can compensate for

reductions in LKB1 signaling. In contrast, Golgi fragmentation

in GM130 shRNA-expressing cells was not rescued by Stk25

overexpression (Figure 4B). Overexpression of either Stk25

WT-GFP or Stk25 K49R-GFP led to the condensation of the

Golgi into a smaller volume (Figure 4C, neurobasal). Therefore,

increasing or decreasing Stk25 expression from endogenous

levels has different consequences for Golgi morphology, in addi-

tion to having the opposite effects on axon production. These

results suggest an LKB1-Stk25-GM130 pathway for Golgi regu-

lation in cultured neurons.


Importantly, Stk25 knockdown in hippocampal pyramidal

neurons also caused Golgi fragmentation in vivo, as determined

by use of in utero electroporation. Normally, the Golgi is strictly

localized to the apical side of the soma and forms outposts in

the apical dendrite (Horton et al., 2005; Figures 2G and 2J; Movie

S2). However, in Stk25 shRNA-expressing, Ctip2-positive

neurons, the Golgi apparatus was often broadly distributed

throughout the soma (Figures 2H and 2J; Movie S2).

In summary, these results indicate that Stk25, LKB1, and

GM130 are required for normal Golgi morphology in neurons at

a time when axons are first appearing. Furthermore, the frag-

mented Golgi phenotype correlated with the loss of axon

production in neurons, and both phenotypes were rescued by

Stk25 overexpression in LKB1 knockdown cells.

Reelin Signaling Regulates Golgi MorphologyAs Stk25 and Reelin have opposing effects on axon initiation

(Figure 1H) and Stk25 affects Golgi morphology (Figures 4A

and 4B), we investigated the role of Reelin in regulating Golgi

morphology.

First we examined the appearance of the Golgi apparatus in

hippocampal and neocortical pyramidal neurons of reelin�/�

and dab1�/� mutant mice. In the pyramidal layer of the wild-

type CA1 zone and in developing neocortical layers, the Golgi

apparati were linearly organized and extended tens of microns

into the apical processes (Figure 5D; Figures S4D and S4G,

insets). The Golgi of the reelin�/� and dab1�/� mutants often

appear convoluted near the nucleus rather than extended into

a dendrite (Figures 5E and 5F; Figures S4E and S4F, insets).

The distance from the Ctip2-positive nucleus to the tip of

the Golgi ribbon was significantly decreased in reelin�/� and

dab1�/� mutants as compared to wild-type (Figure 5G and Fig-

ure S4G), indicating that the reelin and dab1 genes either directly

or indirectly regulate Golgi extension into the apical process of

pyramidal neurons.

As reelin and dab1 also regulate the proper layering of hippo-

campal pyramidal neurons (Caviness and Sidman, 1973; Goffi-

net, 1984; Rice et al., 2001) (Figures 5B and 5C), the effects of

reelin and dab1 on Golgi deployment may be indirect. Therefore,

we tested whether Reelin-Dab1 signaling acutely induces

changes in Golgi morphology or localization by treating hippo-

campal neuron cultures with Reelin for 30 min. Hippocampal

pyramidal neurons were infected with a low titer GFP-expressing

lentivirus to help visualize individual neurons. The Golgi was

largely localized close to the nucleus in control-conditioned

media (CM) and neurobasal-treated Ctip2-positive pyramidal

neurons (Figures 6A and 6C). However, in approximately

80% ± 5% of Reelin-CM-treated neurons, the Golgi apparati

extended into the largest dendritic process (Figures 6A and

6C). The distance between the nucleus and the most distal

portion of the Golgi ribbon from randomly selected Ctip2-posi-

tive neurons was significantly larger in the Reelin-CM-treated

samples compared to the control-CM- and neurobasal-treated

samples (Figure 6B). The Golgi apparatus is therefore rapidly

deployed into dendrites in response to Reelin stimulation.

We next evaluated whether the Golgi response to Reelin was

sensitive to elevated Stk25 expression levels. Hippocampal

neurons were infected with Stk25 WT-GFP or Stk25 K49R-GFP

expressing viruses after 72 hr in culture and treated analogously

to experiments described above. Expression of either Stk25

WT-GFP and Stk25 K49R-GFP reduced but did not eliminate

the Golgi extension in response to Reelin (Figures 6B and 6C).

Under these conditions, linear Golgi ribbons were observed

extending into the dendrites, but on average this was approxi-

mately 50% the distance observed in the Reelin-treated, GFP-

expressing cells (Figure 6B). Furthermore, Reelin signaling

suppressed Golgi compaction induced by Stk25 overexpression

(Figures 4C and 4D). In cultures that were grown in Reelin-CM for

2 days (Figure 4), we did not observe Golgi deployment into

dendrites. This is not surprising as components of the Reelin-

Dab1 pathway begin to be degraded within a few hours. In

60-day-old animals, Golgi extension into dendrites was also

reduced (data not shown). Therefore, Golgi deployment appears

Figure 5. The Golgi Apparatus Extends into an Apical Process in

Neonatal Hippocampus in a reelin- and dab1-Dependent Manner

(A) Ctip2-positive CA1 neurons are organized into a tight lamella in wild-type

brain.

(B) Homozygous disruption of reelin or (C) dab1 causes dispersion of these

neurons.

(D) Confocal imaging through the CA1 region of the wild-type hippocampus

revealed that the Golgi apparatus (white or green, inset) extends radially into

the presumptive apical dendrite of Ctip2-positive neurons (red, inset).

(E) In equivalent reelin�/� or (F) dab1�/� mutant sections, the Golgi is more

often convoluted proximal to the nucleus (inset). Insets were selected from

regions where isolated cells could be distinguished.

(G) The Golgi phenotype was quantified by measuring the distance from the

nucleus to the furthest tip of the Golgi ribbon. (*p < 0.0001, Student’s t test,

n indicated in bar from three animals per group.)

Bar: 200 mm in (C), 20 mm in (F), and 2 mm in inset. See also Figure S4.


to be a transient, developmental phenomenon. Thus, similar to

the manifestation of the multiple axon phenotype caused by

Stk25 overexpression or loss of dab1 gene function, the degree

of Golgi extension seems to be determined by a competition

between Reelin-Dab1 signaling and Stk25 levels.

DISCUSSION

In this study, we find that Reelin-Dab1 signaling acts in an

opposing manner to LKB1, GM130, and Stk25 to regulate the

polarization of axons, dendrites, and Golgi apparati of hippo-

campal neurons, as shown in Figure 7. Knocking down these

three proteins led to Golgi fragmentation and inhibited axon

initiation (Figure 1, Figure 3, and Figure 4). In contrast, Stk25

overexpression caused Golgi condensation and the formation

of multiple axons (Figure 1 and Figure 4). It also rescued axon

production and Golgi fragmentation caused by LKB1 knockdown

but did not rescue either phenotype caused by reduced GM130

expression (Figure 3 and Figure 4), suggesting that Stk25 func-

tions as an intermediary between LKB1 and GM130. Stk25

directly or indirectly binds to the LKB1-STRAD complex and

GM130 and may play a scaffolding role to link LKB1 signaling

to GM130 and Golgi regulation (Figure S2). Reelin-Dab1 signaling

antagonizes the effects of Stk25 overexpression on Golgi

morphology and neuronal polarization as well as inducing polar-

ized deployment of the Golgi into the apical dendrite (Figure 1,

Figure 4, and Figure 6). Together this implicates the LKB1

pathway, GM130, Stk25, and Reelin-Dab1 signaling in Golgi

regulation during neuronal polarization.

Involvement of the Golgi Apparatus in NeuronalPolarizationThe Golgi apparatus and centrosomes reorient as neurons

migrate into the cortical plate (de Anda et al., 2010; Nichols and

Olson, 2010). At the time of axon initiation, the centrosome is

near the basal pole (rear) of the cell. It then moves to the opposite

pole (front) and is important for extending an apical process that

is used for radial migration (de Anda et al., 2010). The apical

process subsequently transforms into the apical dendritic tree,

with the Golgi and centrosomes at its base (Barnes et al., 2008;

Horton et al., 2005). The same events presumably occur during

migration of hippocampal pyramidal neurons in vivo. When

hippocampal neurons are cultured, the centrosome position

determines which neurite becomes an axon (de Anda et al.,

2005). Later, the apical localization of the Golgi apparatus pro-

motes the asymmetric growth of the apical compared to the basal

dendrites (Horton et al., 2005). Consistent with this, Stk25 knock-

down led to Golgi disorganization, inhibited axon induction, and

lessened the asymmetry between the long, thick apical dendrite

and short, slender basal dendrites (Figures 2F, 2H, 2J, and 2K).

The Golgi may influence axon initiation through nucleating

microtubules, regulating secretory trafficking, or interacting

Figure 6. Reelin Stimulation Leads to Rapid Golgi Extension into Dendrites

Primary hippocampal neurons were infected with GFP-expressing viruses after 3DIV and stimulated 3 days later.

(A) The Golgi apparati in Reelin-CM-treated neurons extended tens of microns into dendrites, compared to little or no extension into dendrites of control-CM or

neurobasal-treated neurons.

(B) The distance between the nucleus and the tip of the Golgi was measured for GFP-, Ctip2-positive neurons. Expression of Stk25 WT-GFP and Stk25 K49R-GFP

caused a significant reduction in Reelin-induced Golgi extension.

(C) The Golgi of most GFP-, Ctip2-positive Reelin-CM-treated neurons extended at least 10 mm from the nucleus into or toward a dendrite. Significantly fewer

Golgi were observed in the processes of control-treated samples or Reelin-CM-treated samples that also overexpressed Stk25.

Yellow arrows indicate furthest tip of Golgi ribbon from nucleus. (*p < 0.0001, **p = 0.0002, ***p < 0.05, Student’s t test, between Reelin-CM- and control-treated

samples and between GFP- and Stk25-expressing samples treated with Reelin-CM.) Bars: 10 mm.


with the centrosome (Efimov et al., 2007; Pfenninger, 2009;

Rosso et al., 2004; Sutterlin and Colanzi, 2010). It seems less

likely that the Golgi is required to supply materials to sustain

axon growth, as none of our manipulations affected axon length,

only axon number. Therefore, the Golgi probably has a signaling

or microtubule nucleation role in axon specification. Indeed,

microtubule stabilization has been shown to enhance axon

formation (Witte et al., 2008), and inhibiting post-Golgi trafficking

disrupts axo-dendritic polarization (Bisbal et al., 2008; Yin et al.,

2008). In dendrites, however, the Golgi may have a role in

supplying materials for dendrite growth, as we detected effects

on dendrite thickness and length (Figures 2E and 2K). Deploy-

ment of the Golgi into the apical dendrite may initiate the forma-

tion of dendritic Golgi outposts, which have been shown to

promote dendrite growth and branching (Horton et al., 2005;

Ye et al., 2007).

We found that Stk25 functions in Golgi morphology and axon

specification as part of an LKB1 pathway (Figure 3 and Figure 4).

LKB1, the mammalian Par-4 homolog, is an evolutionarily

conserved cell polarity protein that is known to regulate axo-

dendritic polarity in neurons (Barnes et al., 2008). LKB1 is acti-

vated upon binding STRAD and MO25 (Alessi et al., 2006).

STRAD stabilized LKB1 in processes prior to axon production

and in the nascent axon, suggesting a role in axon specification

(Shelly et al., 2007). As a master kinase, LKB1 activates several

downstream kinases that regulate various aspects of cell

polarity. These include the Sad A and Sad B kinases, which

are required for neuronal polarization (Barnes et al., 2007; Kishi

et al., 2005). Mst4, another downstream kinase, is closely related

to Stk25. Like Stk25, it binds to GM130 and is enriched in the

Golgi apparatus (Preisinger et al., 2004). Both Mst4 and Stk25

are required downstream of LKB1-STRAD induction for polar-

ized brush border formation in epithelial cells (ten Klooster

et al., 2009; Figure S2). However, although Mst4 kinase activity

is required during this process, the kinase activity of Stk25 is

not needed to induce polarized brush border formation, regulate

Golgi morphogenesis, or polarize hippocampal neurons (Fig-

ure 1F and Figures 4C and 4D). This suggests a kinase-indepen-

dent scaffolding function for Stk25 (Figure 7), which is reminis-

cent of the pseudokinase STRAD (Lizcano et al., 2004). GM130

appears to be necessary for Stk25 effects on Golgi and neuronal

polarization; however, it may not be sufficient. By linking LKB1

signaling to GM130, Stk25 may directly regulate GM130 or indi-

rectly modulate the activity of other Golgi proteins.

Reelin-Dab1 Signaling Regulates Neuronal Polarizationand Golgi DeploymentOur work also shows that Reelin-Dab1 signaling, acting in oppo-

sition to LKB1-Stk25-GM130, affects Golgi morphology and

axon formation. The absence of Reelin or Dab1 inhibited Golgi

deployment into the apical dendrite in vivo (Figure 5 and

Figure S4), and long-term growth in Reelin opposed Golgi

condensation induced by Stk25 overexpression in vitro (Fig-

ure 4). Similarly, Dab1 absence induced supernumerary axons

in vitro (Figure 1H), the opposite effect to depleting Stk25.

However, Reelin-Dab1 and LKB1-Stk25-GM130 do not fit into

a simple epistatic relationship. For example, Stk25 depletion

reduces axon number even when Dab1 is absent, suggesting

that Stk25 does not require Dab1 to regulate axon number (Fig-

ure 1). This indicates that LKB1-Stk25-GM130 and Reelin-Dab1

act on the Golgi and axon initiation through different pathways,

and the balance between the two pathways determines the

outcome. In this respect, Golgi distribution is a quantitative trait,

not all or none, and may be influenced by other factors. Indeed,

extended Golgi were observed in a subset of neurons in reelin�/�

and dab1�/� mutant brains (Figure 5 and Figure S4). One possi-

bility is that Reelin-Dab1 and LKB1-Stk25-GM130 regulate

different aspects of Golgi morphology through different mecha-

nisms. For example, Reelin-Dab1 may regulate ER-Golgi vesicle

movement, and LKB1-Stk25-GM130 may affect vesicle fusion.

In sum, we have characterized Stk25, a modifier of the Reelin-

Dab1 pathway, and shown that it acts on the LKB1-STRAD

pathway to regulate Golgi morphology and neuronal polariza-

tion. Stk25 may play a scaffolding role to link LKB1-STRAD to

Golgi regulation through binding GM130, as the kinase activity

was shown to be dispensable for neuronal polarization and Golgi

morphogenesis. We find that Reelin-Dab1 signaling regulates

Golgi morphology and deployment into dendrites in a competi-

tive manner with Stk25. Golgi position has been shown to

enhance local secretory trafficking (Horton et al., 2005; Ye

et al., 2007); thus, this competition may regulate membrane

and protein cargo flow into proximal dendrites. Our findings

provide new insights into the regulation of morphogenic changes

in neurons that drive neuronal polarization and brain lamination.


Expression Vectors

The lentiviral vectors used in this study were based on pLentiLox 3.7 (pLL3.7)

vectors (Rubinson et al., 2003) with the following substitutions: (1) for shRNA

experiments, instead of the CMV promoter, the CMV enhancer/chicken b-actin

Figure 7. Model of Stk25 as a Scaffolding Protein Acting Competi-

tively with Reelin-Dab1 Signaling

LKB1 is known to act in complex with STRAD to regulate cellular polarity

(Alessi et al., 2006). Reelin, the receptors ApoER2 and VLDLR, and Dab1

also form a signaling complex (Hiesberger et al., 1999; Trommsdorff et al.,

1998). STK25 coimmunoprecipitates with STRAD and GM130 (Figure 2S).

Overexpression of LKB1 and STRAD is known to induce the formation of

multiple axons (Barnes et al., 2007; Shelly et al., 2007). Independent of its

kinase activity, STK25 does so also and induces Golgi condensation (Figure 1F

and Figure 4A). Knocking down LKB1, Stk25, or GM130 causes Golgi frag-

mentation/dispersion and lost axon production, the opposite to Golgi conden-

sation and multiple axon formation (Figure 1, Figure 3, and Figure 4) (Barnes

et al., 2007; Shelly et al., 2007). The overexpression phenotypes are sup-

pressed by Reelin stimulation. Dab1�/� neurons (Reelin signaling deficient)

have multiple axons and shorter dendrites (Figure 1F) (Niu et al., 2004). Reelin

stimulation induces Golgi deployment and dendrite growth, phenotypes sup-

pressed by Stk25 expression/overexpression (Figure 2 and Figure 6).


promoter (Niwa et al., 1991) directs GFP expression; (2) for fusion protein

experiments, instead of the U6 promoter the CMV enhancer/chicken b-actin

promoter directs expression. The shRNA constructs include Stk25 shRNA AG

GAGCTCCTGAAGCACAAAT and control shRNA AGTAGCTCCTAAAGCACA

CAT. The lentivirus production was as previously described (Matsuki et al.,

2008). The knockdown viruses were confirmed to reduce expression of either

Stk25, LKB1, or GM130 (Figure S1 and Figure S3). The Stk25 K49R mutant has

previously been reported to be kinase inactive, which we confirmed (Preisinger

et al., 2004 and data not shown).

Animals

All animals were used in accordance with protocols approved by the Animal

Care and Use Committees of SUNY Upstate Medical University, National Insti-

tutes of Neurological Disorders and Stroke, and the Fred Hutchinson Cancer

Research Center, following NIH guidelines. Time pregnant mice (C57BL/6

for in vitro experiments and Swiss Webster for in utero electroporations) and

rats (Sprague Dawley) were purchased from Charles River Laboratories and

Taconic. The dab1�/� (Howell et al., 1997) and reelin�/� (Jackson Labs)

mice were on the C57BL/6 strain.

Immunocytochemistry

Immunocytochemistry was done according to published methods (Matsuki

et al., 2008) and is detailed in the Extended Experimental Procedures along

with a list of the antibodies used. To measure Golgi volumes and length of

the longest Golgi ribbon, we immunostained the neurons with anti-GRASP65,

anti-GFP, and anti-Ctip2, which recognizes a CA1 and layer V pyramidal

neuron-specific transcription factor. The area of the Golgi apparatus was

calculated for each Z-plain (Image Examiner, Zeiss), multiplied by the thick-

ness of the section, and summed to determine the volume.

Cell Culture

Hippocampal neuronal cultures were isolated from embryonic day (E) 17.5

mice or E18.5 rats and grown in neurobasal samples supplemented with 2%

B27 (Invitrogen, Matsuki et al., 2008). For polarity studies, neurons (1 3 104

cells per cm2) were infected with the respective viruses on the day of culturing

and replated 2 days later on poly-L-lysine coated coverslips placed over

a monolayer of astrocytes. Axons were quantified at 2 days in vitro (DIV) or

6DIV as indicated, following standard criteria (Shelly et al., 2007). For Golgi

deployment assays, rat cultured neurons (3 3 105 cells per cm2) were infected

with low titer virus on day 3 and treated and fixed on day 6 in culture. Similar

results were obtained with mouse neurons (data not shown). The control-

and Reelin-conditioned media were collected and concentrated as previously

described (Matsuki et al., 2008).

Analysis of In Utero Electroporated Brains

To knock down Stk25 expression, DNA was injected into the lateral ventricle of

E17.5 embryos of Swiss Webster mice in utero and electroporated (70 mV) as

previously described (Olson et al., 2006) with the electrode paddles oriented to

direct the DNA into the hippocampus. Perfused brains were processed for

analysis on P7. Floating sections (70–100 mm) were immunostained with anti-

bodies described in the figure legends. Confocal images were collected with

overlapping optical sections through 30 mm, which were flattened for display.

We assessed whether axon initial segments or Golgi elements belonged to

a particular GFP-positive neuron (Figure 2), by examining movies of either

3D-rendered images or Z sections (Movie S1 and Movie S2). Golgi areas

(Figures 2G and 2H) were produced by thresholding (Adobe Photoshop) flat-

tened, 2D-negative images to match the GRASP65 signal channel in the orig-

inal and discarding the signal extraneous to the GFP-positive cells (Movie S2).

Process diameters were measured 12 mm from the nucleus (Figure 2K). These

measurements were done using Image Examiner (Zeiss). Measurement of

dendrite lengths was done using the softWoRx (AppliedPrecision).


Supplemental Information includes Extended Experimental Procedures, four

figures, and three movies and can be found with this article online at doi:

10.1016/j.cell.2010.10.029.

ACKNOWLEDGMENTS

We would like to thank Zainab Mansaray and Kristin Giamanco for experi-

mental assistance, Michael Zuber for comments on the manuscript, Hans

Clevers for cell lines, Louis Cantley and Jun-ichi Miyazaki for DNA vectors,

Arvydas Matiukas and Melissa Pepling for assistance with confocal micros-

copy, and Bonnie Lee Howell for editing. This work was supported by funds

from the NINDS intramural program and SUNY Upstate Medical University

to B.W.H.; NIH grants NS066071 to E.C.O., NS069660 to R.T.M., and

CA41072 to J.A.C.; and NIA intramural funds for M.R.C.





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Resource

A Human Genome Structural VariationSequencing Resource Reveals Insightsinto Mutational MechanismsJeffrey M. Kidd,1,4 Tina Graves,2 Tera L. Newman,1,5 Robert Fulton,2 Hillary S. Hayden,1 Maika Malig,1 Joelle Kallicki,2

Rajinder Kaul,1 Richard K. Wilson,2 and Evan E. Eichler1,3,*1Department of Genome Sciences, University of Washington School of Medicine, Seattle, WA 98195, USA2Washington University Genome Sequencing Center, School of Medicine, St Louis, MO 63108, USA3Howard Hughes Medical Institute, Seattle, WA 98195, USA4Present address: Department of Genetics, Stanford University, Stanford, CA 94305, USA5Present address: iGenix, Seattle, WA 98110, USA


SUMMARY

Understanding the prevailing mutational mecha-nisms responsible for human genome structural vari-ation requires uniformity in the discovery of allelicvariants and precision in terms of breakpoint delinea-tion. We develop a resource based on capillary endsequencing of 13.8 million fosmid clones from 17human genomes and characterize the completesequence of 1054 large structural variants corre-sponding to 589 deletions, 384 insertions, and 81inversions.We analyze the 2081 breakpoint junctionsand infer potential mechanism of origin. Threemechanisms account for the bulk of germline struc-tural variation: microhomology-mediated processesinvolving short (2–20 bp) stretches of sequence(28%), nonallelic homologous recombination (22%),and L1 retrotransposition (19%). The high qualityand long-range continuity of the sequence revealsmore complex mutational mechanisms, includingrepeat-mediated inversions and gene conversion,that are most often missed by other methods, suchas comparative genomic hybridization, single nucle-otide polymorphism microarrays, and next-genera-tion sequencing.

INTRODUCTION

Despite significant advances in the discovery and genotyping of

human genome structural variation, only a small fraction of

common structural variation has been resolved at the sequence

level (Conrad et al., 2010b; Freeman et al., 2006; Itsara et al.,

2009; Kidd et al., 2008; Lam et al., 2010; McCarroll et al.,

2008b; Redon et al., 2006). The majority of human genome struc-

tural variation has been discovered with single nucleotide poly-

morphism (SNP) microarrays and array comparative genomic

hybridization (arrayCGH), approaches that provide limited infor-

mation about the precise structure and location of identified vari-

ants. Because of their dependence on the reference genome,

array-based approaches preferentially detect deletions over

insertions and are unable to directly detect copy-number-neutral

events such as inversions. Higher-density array platforms give

a better estimation of variant sizes, but most breakpoints cannot

be resolved at a scale finer than 50 bp regions (Conrad et al.,

2010b), while targeted next-generation sequencing approaches

have difficulty resolving breakpoints within homologous

segments (Conrad et al., 2010a).

These methodological biases threaten to skew our under-

standing of the underlying mechanisms responsible for the

formation of structural variation and limit our ability to compre-

hensively discover and genotype this form of genetic variation.

We resolve the breakpoints of 1054 structural variants based

on capillary sequencing of clone inserts. The high-quality

sequence of contiguous variant haplotypes allows alternative

structures to be included in future human genome assemblies

and provides the breakpoint resolution necessary to accurately

genotype these variants in sequence data generated from

next-generation sequencing platforms. The sequences and the

associated clones also provide a resource for assessing future

methods for structural variation discovery.

RESULTS

The Human Genome Structural Variation CloneResourceThe high quality of the reference human genome is due, in large

part, to the fact that it was assembled based on capillary

sequencing of individual large insert clones whose complete

sequence was resolved prior to final genome assembly. This

strategy allowed complex duplicated and repetitive regions to

be incorporated that were missed by other approaches (Istrail

et al., 2004; She et al., 2004). Since genome structural variation

is similarly biased to these regions, we proposed that developing

clone libraries for a modest number of additional genomes would

serve as a valuable resource for characterizing complex and

difficult-to-assay regions of genome structural variation (Eichler


et al., 2007). The overall strategy involved the construction of

individual genome libraries using a fosmid cloning vector

(40 kb inserts) and capillary sequencing of the ends of the inserts

to generate a high-quality end-sequence pair (ESP). Discrep-

ancies in the length and orientation of these mapped ESPs

with respect to the reference genome serve as signatures of

copy-number variation and inversion, respectively. Since the

underlying clones can be retrieved, the complete sequence

context of the discovered structural variant can also be obtained.

Previously, we discovered and cloned 1695 structural variants

with fosmid libraries derived from nine individuals and presented

sequence of 261 structural variants (Kidd et al., 2008; Tuzun

et al., 2005). We expand this resource to include capillary end

sequencing of 4.1 million additional fosmid clones from eight

additional human genomes (Table S1, available online).

The combined set includes 13.8 million clones derived from the

genomes of six Yoruba Nigerians, five CEPH Europeans, three

Japanese, two Han Chinese, and one individual of unknown

ancestry.

Structural Variant AllelesUsing this resource, we searched for clusters of clones that

suggest a structural difference when compared to the reference.

We discovered a total of 2051 discordant regions (Table S1)

having support from multiple clones for a structure different

from the reference genome. The size distribution of the fosmid

clone inserts limited us to the detection of structural variants

greater than 5 kb in length. Inversions also tend to be biased to

larger events because of the probability of capturing a breakpoint

by a pair of end sequences. While there is no upper bound in the

detection of deletions and inversions, the direct capturing of

insertions larger than the insert size of the clone (40 kb) requires

specialized approaches. For example, new tandem duplications

may be identified with an everted clone mapping signature (Fig-

ure S1) (Cooper et al., 2008) and insertions of novel human

sequence may be identified by read pairs for which only one

end maps (Kidd et al., 2010).

We targeted 1054 structural variants (Table S1) from nine

human genomes and completely sequenced the inserts of

1167 fosmid clones (46.4 Mb of sequence). We identified 81

loci for which breakpoints could not be resolved because of diffi-

culty in clone assembly and the limits of 40 kb fosmid inserts (see

Supplemental Experimental Procedures). We defined break-

points relative to the reference genome assembly following

a two-stage procedure (Kidd et al., 2010) (Figure 1 and Table

S2). We initially distinguished copy-number changes (n = 973

insertion and/or deletions) from balanced genome structural

variants (81 inversions) (Figure 2). The analyzed variants altered

95 gene structures. We estimate that 1.04% (11/1054) of the

sequenced alleles are already known risk factors for common

and rare human diseases (Figure 3 and Table S3).

Breakpoint FeaturesUsing the 40 kb of clone-based sequence, we examined the

sequence features and inferred potential mechanism of origin

for these variants (Table 1). We identified 30 variants associated

with the expansion or contraction of a variable number of

tandem repeats (VNTRs) (Buard et al., 2000; Jeffreys et al.,

1994; Richard et al., 2008). VNTR repeat units ranged from

17 bp to 6.5 kb with copy numbers ranging from 1 to 319

copies. We identified 198 events (20% of the total insertions

and deletions) that we classified as being the result of L1

retrotransposition. Each of the 198 L1 elements associated with

the retrotransposition events has a sequence identity of at least

97.5% when compared to the L1.3 reference sequence, and

152 are at least 6 kb in size, consistent with full-length elements

that may be capable of subsequent retrotransposition (Beck

et al., 2010). We find evidence for transduction of flanking

sequence for 20% (40/198) of the sites, with the transduced

segment size ranging from 45 to 968 nucleotides (median of

81.5) (Goodier et al., 2000; Moran et al., 1999; Pickeral et al.,

2000). Using the transduced sequence as a marker, we identi-

fied the potential donor location for 30 of these retrotransposi-

tions (20 insertions in the fosmid source sample and 10

insertions in the reference genome). We identified three posi-

tions that have each given rise to multiple LINE insertions (Fig-

ure 2B), suggesting the presence of L1 donor hotspots. We

note that 11 of the 20 L1 insertions in the fosmid source

(including the three recurrent L1 donors) correspond to

elements that have been functionally determined to represent

hot L1s, according to assays performed by Beck et al. (2010).

We found two events consistent with the insertion of an intact

HERV-K element: one insertion in the reference sequence (as

indicated by clone AC209281) and an insertion contained in

clone AC226770. Both events showed less than 1% divergence

from the HERV-K sequence (Dewannieux et al., 2006) and were

flanked by long terminal repeats (Tristem, 2000). Our discovery

size thresholds (>5 kb) preclude the identification of smaller ret-

rotransposition events arising from SVA or Alu repeats that are

common when smaller structural variants are considered (Ben-

nett et al., 2008; Korbel et al., 2007; Lam et al., 2010; Mills et al.,

2006).

We divided the remaining 824 structural variants into two

broad categories. Class I consists of variants with no additional

sequence at the breakpoint junction (Figures 4A–4D and

Figure S2). Class II variants contain an additional sequence,

found across the variant junction, that is not present at either

of the other variant breakpoints (Figures 4E–4G). We also

assessed the presence of extended sequence homology

and the extent of matching sequence at the breakpoints. We

note that microhomology is a qualitative term without clear delin-

eation as 1 or 2 bp matches are expected to occur often by

chance (Figure 4) and a range of homologous match lengths is

observed (Conrad et al., 2010a; Lam et al., 2010). Similarly, there

is ambiguity in assigning events to potential mechanisms based

solely on the length of homologous segments. Consequently, we

categorize events based on observed ranges of homology and

consider assignment to specific mechanisms as speculative.

Among the class I events, 49% (289/590) of copy-number

variants contain 2–20 bp of matching sequence, indicating that

microhomology-mediated mechanisms, such as microhomol-

ogy-mediated end joining (MMEJ), contribute to a substantial

fraction (30%) of human structural variation (Table 1) (Hastings

et al., 2009; McVey and Lee, 2008; Payen et al., 2008; Roth

and Wilson, 1986). Although there is large overlap in the variant

size when broken down by extent of homologous sequence


(Figure 4C), we find that, as a class, the mean size of events

associated with microhomology (2–20 bp of matching sequence,

n = 289, mean size is 9.7 kb) is significantly smaller (p = 0.02926,

two sample t test) than those showing a hallmark of nonallelic

homologous recombination (NAHR) (R200 bp of matching

sequence, n = 177, mean size is 21.0 kb). The analyzed inver-

sions are overwhelming driven by large homologous segments

with 69% (56/81) of all analyzed inversions containing stretches

of matching sequence at least 200 bp in length. In contrast, only

30% (177/590) of the class I copy-number variants contain

matching breakpoint sequences of at least that length. It is

important to note, however, that our clone end-sequence

mapping strategy is biased toward the detection of larger inver-

sions when compared to copy-number variants. This is a direct

consequence of the probability of capturing a breakpoint that

diminishes when inversions become smaller than the clone insert

size. Overall, we find that younger Alu events and segmental

duplications contribute most significantly to the process of

NAHR (Table S4), as expected because of their higher levels of

sequence identity. The strongest enrichment is found for paired

Alu repeats at each breakpoint (5.2-fold enrichment). If each

breakpoint is treated separately, rather than requiring that an

element of the same subfamily be present at both breakpoints

of a variant, then AluY also shows a substantial degree of enrich-

ment (2.6-fold, Table S4). Since AluY is the most recently active

Alu family, dispersed AluY elements are expected to have

a higher degree of sequence identity than other Alu families (Bat-

zer and Deininger, 2002; Cordaux and Batzer, 2009). Closer

examination of the distribution of breakpoints within individual

Alus reveals a nonuniform pattern of breakpoint density (Fig-

ure 3D). The highest density of breakpoints occurs near the posi-

tion of a sequence motif (CCNCCNTNNCCNC) that has been

Align against common junction sequence

| |----| || ||||||||

|||||||| |||--||--||

| |----| || |||| ||||

|||||||| |||--||--||

| |----| || |||| |||| |||||||| |||--||--||

*1*1111* **122** **22

|||||||| |||||||||||| ||||

|| ||

| |----| || |||| |||| |||||||| |||--||--||

Combine pairwise alignments Assess identity

10 kb deletion

break 1 break 2

deletion junction

A

B

C

Figure 1. Sequence and Breakpoint Analyses

Variant breakpoints were defined based on alignments of sequences from the sequenced insertion and deletion alleles. For example, (A) the sequence of fosmid

clone AC207429 is compared with sequence from the corresponding region on chr2. A 10 kb deletion, relative to the reference sequence, is readily apparent

(indicated by the red bracket). The position of segmental duplications, common repeats (LINEs are green, SINEs are purple, and LTR elements are orange),

and RefSeq exons are shown. Sequence segments corresponding to three different breakpoint regions (red, green, and purple bars) are extracted for further

analysis.

(B) The sequence across the variant junction is aligned against each of the other two sequences and the resulting pairwise alignments are merged. The pattern of

sequence identity is assessed to identify the positions where the junction sequence switches from being a better match to the first breakpoint to being a better

match to sequence from the second breakpoint. The breakpoint coordinates correspond to the innermost positions that can be confidently assigned to be before

and after the variant boundary.

(C) The result of aligning the three segments depicted in (A). Alignment columns where the junction sequence matches the sequence from the first (leftmost)

breakpoint are indicated by a 1 while alignment columns where the junction sequence matches the second (rightmost) breakpoint are indicated by a 2. Positions

where all three sequences are the same are indicated by an asterisk (*). The red square highlights the position of the breakpoint coordinates (highlighted in red and

green text). The two breakpoints are separated by seven nucleotides found at both breakpoints with perfect identity (blue text). Highlighted in gray is a 293 bp

segment present at both breakpoints with a sequence identity of 91%. See also Tables S2 and S7.


associated with meiotic recombination hotspots, is found in

some Alu elements (Myers et al., 2008), and has also been

observed for rearrangements between human and chimpanzee

(Han et al., 2007; Sen et al., 2006).

We find that 16% (153/973) of the insertion and deletion vari-

ants and 9% (7/81) of the inversions contain additional

sequence at the variant breakpoints (class II events; Figure 4).

Many of the additional insertion sequences are relatively short

in length, consistent with nontemplate-directed repair associ-

ated with nonhomologous end joining (Figure 4B). For these

shorter sequences, no inference could be made as to the source

of the additions. However, 41% of all class II variants (66/160)

contain additional sequence at the junction at least 20 bp in

length. Of these longer fragments, 88% (58/66) map to another

location within the human genome. Since we are limited in this

study to directly capturing the breakpoints of insertions smaller

than 40 kb, we repeated this comparison with only deletions

relative to the assembly where we expect to have less of

a bias in terms of variant size. We find that the additional junction

sequences for 30 of 39 class II deletion events at least 20 bp

long map elsewhere in the genome. Seventy-three percent

(22/30) are found on the same chromosome as the variant.

In fact, eight of the insertions map less than 1 kb away from

the variant breakpoint (Figure 4G and Table S5) and all 22 are

less than 250 kb from the breakpoint. This pattern suggests

the action of a replication-associated process that involves

template switching or strand invasion (Hastings et al., 2009;

Lee et al., 2007; Smith et al., 2007). In contrast to the class I

events, only 2% of the class II events (3/160) contained

stretches of homologous sequence flanking the breakpoint

insertion confirming they arose by mechanisms other than

NAHR. Interestingly, if we examine the sequence context of

these regions, we find that 20% (30/153) of class II events

map within 5 kb of a segmental duplication. This represents

a significant enrichment for proximity to duplicated sequence

(p < 0.002 based on comparisons with randomly sampled

sequences) indicating that regions flanking segmental duplica-

tions may be generally more unstable and susceptible to

multiple mutational processes such as template switching

during replication (Itsara et al., 2009; Lee et al., 2007; Payen

et al., 2008).

Gene Conversion and Structural VariationDuring our analysis of putative NAHR events, we identified

10 structural variants having a complex pattern of exchange

inconsistent with a simple model of unequal crossover. The

breakpoint region contains an interleaved pattern of alternating

patches of sequences from flanking homologous segments

(Figure 5). These patterns are reminiscent of multiple rounds of

gene conversion, although each of these events was also asso-

ciated with a copy-number variant event. Using paralogous

sequence variants that distinguish the 50 and 30 homologous

segments, we investigated the overall extent of this nonallelic

exchange (referred to as the conversion tract length), and the

number of switches before unambiguous homology to the 50 or

30 end was re-established. We determined that most (6/10) of

the conversion tracts were relatively short (200–600 bp in length)

with a relatively consistent number (4–6) and length (30–40 bp) of

A B

Figure 2. Sequenced Structural Variant Alleles

(A) Size distribution for 1054 sequenced structural variants. Insertions, deletions, and inversions relative to the genome reference assembly are depicted sepa-

rately. Note that the bins are not of equal sizes. The mean size of the sequenced variants is 14.9 kb for deletions, 6.1 kb for insertions, and 196 kb for inversions.

Our variant selection methodology largely identifies deletions greater than �5 kb and insertions from �5 kb to �40 kb in size and is biased against inversions

smaller than �40 kb.

(B) The relationship between the donor site of transduced sequences and LINE insertion position are given for 30 events with a match to hg18 using BLAT. Rela-

tionships are shown for 20 LINE insertions in library source individuals relative to the reference (blue lines) and for 10 insertions in the genome reference (red lines).

The blue circles represent three different loci associated with multiple distinct LINE insertions. See also Figure S1 and Table S1.


switches before clear boundaries at the 50 and 30 could be re-es-

tablished (Figure S3). Seven of these events have breakpoints

that map within segmental duplications, and the remaining three

have breakpoints that map within LINEs. Three of the variants

contained at least ten switches. One variant (AC212911) showed

the largest associated conversion tract with a remarkable 182

switches extending over 7.9 kb (Figure 5D). We sequenced the

deletion allele with fosmids derived from three different individ-

uals for one event (AC226182). Each of the three deletion haplo-

types contained identical patterns of interleaved sequence,

a finding that is consistent with the creation of the pattern at

the time of variant formation, or shortly thereafter, rather than

as a result of a continual conversion process between deletion

and insertion alleles leading to a diverse set of related molecules

over time (Figure S3). It is also possible that the conversion

pattern arose before the formation of the structural variant and

that the pattern we observe in sequenced variants is merely

incidental or the result of a series of mismatch repair processes

prior to variant formation. Nevertheless, the observed switch

pattern is reminiscent of patterns of toggling previously

observed at some LINE insertions (Gilbert et al., 2005, 2002;

Symer et al., 2002) and suggests a mechanism of serial strand

invasion/repair during the rearrangement process.

Comparison with Other Genome-wide Studiesand Ascertainment BiasesIn this study we focused on systematically characterizing large

structural variants at the single base-pair level. In order to identify

events that may have been missed by the fosmid ESP approach,

we compared our set of structural variants to other studies that

have discovered and genotyped copy-number variants in the

same DNA samples. We focused on five individuals analyzed

by fosmid end sequencing (Kidd et al., 2008), Affymetrix 6.0 mi-

croarray (McCarroll et al., 2008b), and high-density oligonucleo-

tide arrayCGH (Conrad et al., 2010b). A comparison of the three

studies shows that 11%–65% of discovered variants are unique

to a single study and corresponding experimental platform (Fig-

ure 6). The limited overlap should not be surprising since each

approach preferentially identifies a subset of the total collection

of genomic variation. For example, the fosmid ESP mapping

approach can detect insertions of sequence not represented in

the genome assembly (Kidd et al., 2008, 2010), as well as

balanced events such as inversions (not depicted in Figure 6),

whereas array approaches can more readily detect copy-

number variation caused by large duplications.

Differences in ascertainment extend to the resolution of break-

point sequences. The sequenced variants described in this

chr1

TMEM50A

C1orf63 RHD

AC196511

chr1

LCE3DLCE3ELCE3ALCE3C LCE3B

Repeats

SegDupMasker

0.0

AC196522

AC207974

chr5

MST150 IRGMNEGR1

Repeats

SegDupMasker

AC210916

chr1

10 20 30 40 kb

10 20 30 40 kb10 20 30 40 kb

10 20 30 40 kb

A

B

C

D

Figure 3. Examples of Sequenced VariantsExamples of the complete sequence of structural variant alleles that have been associated with disease risk, including (A) a 45.5 kb deletion upstream of NEGR1,

(B) a 72 kb deletion of RHD, (C) a 3.9 kb and a 20.1 kb deletion upstream of IRGM, and (D) a 32 kb deletion of LCE3C. See also Table S3.


manuscript include 237 of the regions targeted for array capture

and 454 sequencing (Conrad et al., 2010a). Seventy of these

targeted events were successfully resolved by breakpoint

array-capture experiments (Table S6), with none of the events

containing extended breakpoint homology successfully resolved

by next-generation sequencing.

We also reassessed regions discovered by other studies that

were missed by the fosmid ESP approach. With the standard

fosmid analysis criteria (two or more discordant clones with suffi-

cient quality) (Tuzun et al., 2005), an overlapping deletion site is

only identified for 53% (631/1193) of the corresponding deletion

genotypes reported by Conrad et al. (2010b). The intersection

rate increases to 75% (900/1193 sample-level genotypes) if indi-

vidual deletion clones are considered with reduced quality

thresholds. This suggests that much of the variation missed by

the fosmid ESP approach is a result of random fluctuations in

the level of clone coverage and the quality of individual

sequencing reads (Cooper et al., 2008).

Experimental approaches to discover structural variation can

have reduced sensitivity in regions of segmental duplication

because of difficulty in uniquely mapping reads or designing

array probes (Cooper et al., 2008; Kidd et al., 2008; Tuzun

et al., 2005). We compared the validated structural variants

from Kidd et al. (2008) with those found by read-depth

approaches (Alkan et al., 2009). Alkan et al. (2009) identified

113 genes that differ in copy number among three individuals.

Only 38% of the genes greater than 5 kb (26/69) and identified

as copy-number variable by read-depth intersect with a struc-

tural variant (reported in Kidd et al.[2008]). This result indicates

that even the fosmid ESP approach has underascertained

copy-number variation associated with the most variable dupli-

cated sequences.

We identified 81 loci during our sequence analysis with

evidence for a nonreference structure for which we could not

unambiguously define the variant breakpoint (see Supplemental

Experimental Procedures). Of these 81 loci, 63 are associated

with segmental duplications, including ten examples of tandem

duplications. We note that 23 of these duplication-containing

loci map near gaps in the National Center for Biotechnology

Information (NCBI) build36 genome assembly or to sequences

that have been assigned to a chromosome but not fully inte-

grated into the genome reference sequence. Duplication-medi-

ated copy-number variation remains underascertained in terms

of sequence-level resolution of variant haplotypes and muta-

tional mechanism analysis. If we adjust for these biases, we esti-

mate that the fosmid ESP approach has minimally missed at

least 106 structural variants associated with segmental

duplications.

DISCUSSION

We describe a clone resource from 17 human DNA samples that

provides 135-fold physical coverage of the human genome. The

corresponding catalog and clones can be used to further charac-

terize almost any segment of human euchromatin. We used this

resource to assess breakpoint characteristics of 1054 events.

The nature of our experimental design permitted us to discover

more events mediated by larger segments of homology, providing

a more complete assessment of human genetic variation. Of

particular interest are complex events whose sequence features

have been difficult to previously assess at a genome-wide level.

The high quality and length of the sequenced fosmids combined

with defined paralogous sequence events allowed us to quantify

alternating sequence matches suggestive of interlocus gene

conversion (Bayes et al., 2003; Lagerstedt et al., 1997; Reiter

et al., 1997; Visser et al., 2005).

Using this resource, we obtained the complete structure of

several alleles that have been associated with disease, including

a deletion variant upstream of the NEGR1 gene associated with

increased body mass index (Willer et al., 2009) (clone

AC210916), two deletion polymorphisms upstream of the

IRGM gene associated with Crohn’s disease (Barrett et al.,

Table 1. Summary of Events and Inferred Mechanisms

Event Classification Insertions and Deletions Inversions Potential Mechanisms

Retroelements

L1 198 (20.3%) NA Retrotransposition

HERV-K 2 (0.2%) NA Retrotransposition

VNTR 30 (3.1%) Minisatellite, NAHR

Class I (no additional sequence at breakpoint) 590 (60.6%) 74 (91.3%)

0 or 1 matching nucleotides 82 (8.4%) 10 (12.3%) NHEJ

2–20 matching nucleotides 289 (29.7%) 8 (9.9%) NHEJ, MMEJ

21–100 matching nucleotides 28 (2.9%) 0 NAHR, other

101–199 matching nucleotides 14 (1.4%) 0 NAHR, other

R200 (NAHR) 177 (18.2%) 56 (69.1%) NAHR

Class 2 (additional sequence at breakpoint) 153 (15.7%) 7 (8.6%)

1–10 additional nucleotides 76 (7.8%) 2 (2.5%) NHEJ

>10 additional nucleotides 77 (7.9%) 5 (6.2%) NHEJ, FoSTeS,template switching

Total 973 81

The number of events that fall into each breakpoint class is given. The following abbreviations are used: NHEJ, nonhomologous end joining; FoSTeS,

fork stalling and template switching. See also Table S6.


50 100 150 200 250 300

0.02

0.04

0.06

0.08

0.10

0.12

Position in Alu

Bre

akpo

int D

ensi

ty

A B

C D

5’bkpnt CAAATGCAATGTTTATTAAGCAGGTACTTTGTGCTCAAGAGTATGATACAGAGCACTATAC209239 CAAATGCAATGTTTATTAAGCAGGTACTTTGTGCTCAAGAGTATGATACAGAGCACTAT

5’bkpnt GCTGGGAC209239 GCTGGGATTTGGCAGAGGGGGATTTGGCAGGGTCATAGGACAACAGCGGAGGGAAGGTC

AC209239 AGCTCAGGAGGCTTAGGCATGAGAATCACTTGAACCTGGTAGGCA3’bkpnt CTCAGGAGGCTTAGGCATGAGAATCACTTGAACCTGGTAGGCA

E

F

G

Figure 4. Variant Breakpoint Analyses

(A–D) Class I variants are defined as those without additional nucleotides at the breakpoint. (A) A histogram of the extent of matching breakpoint sequence (black)

and extended breakpoint homology (gray) is shown for 590 class I copy-number events. The red line corresponds to the expected distribution of breakpoint

match lengths found from 100 random permutations. Note that bin sizes are not equal. The increase in extended homology segments 250–299 bp in length corre-

sponds to variants having Alus at their breakpoints. (B) As in (A) zoomed in to show variants having a matching sequence of 20 bp or less. (C) Box plot of variant

size partitioned by length of extended breakpoint homology for 590 class I copy-number variants (red line: median; blue box: interquartile range; whiskers: within

1.53 interquartile range). (D) Breakpoint density map within a consensus Alu repeat sequence based on 269 copy-number variant events (blue box: RNA pol III

promoter; black boxes: AT-rich segment between the two monomers that make up the Alu element and the poly A tail; purple box: position of motif

(CCNCCNTNNCCNC) found in some Alus and associated with recombination hotspots [Myers et al., 2008]).

(E–G) Class II variants contain additional sequence across the breakpoint junction. (E) A class II variant containing a 55 nucleotide-long stretch of additional

sequence (in blue) that is not found at either breakpoint. (F) Histogram of the length of additional sequence found at variant breakpoints (black) and the length

of detected extended homology between breakpoint sequences (gray) for 153 class II copy-number variants. (G) Genomic location for class II unmatched

sequences (>20 bp) associated with deletions. The black lines connect the positions of a class II deletion variant (relative to the genome assembly) and the cor-

responding location where the additional sequence across the variant breakpoint can be found. The relationship for 31 deletion variants is depicted. One event

involves a match to unlocalized sequence on chromosome 1 (chr1_rand). See also Figure S2 and Tables S4 and S5.


2008; Bekpen et al., 2009; McCarroll et al., 2008a) (clone

AC207974), and the deletion of the LCE3B and LCE3C genes.

In total, we conservatively estimate that 1.04% (11/1,054) of

the discovered variants are associated with disease. This yield

of disease-causing alleles rivals that found by genome-wide

association studies using SNPs, which have identified 779

genome-wide associations based on genotyping of at least

100,000 SNPs (http://www.genome.gov/multimedia/illustrations/

GWAS2010-3.pdf).

Although the functional significance of many of the other struc-

tural variants remains to be determined, the clone resource and

availability of the complete sequence of variant haplotypes will

facilitate future disease association through the rapid design of

assays to test for association with disease (Abe et al., 2009; An

et al., 2009; Kidd et al., 2007) or direct comparison with short

sequencing reads from next-generation sequence platforms

(Kidd et al., 2010; Lam et al., 2010).

We investigated this approach for 1024 non-VNTR sequenced

structural variants (Table S7) and found that 71% (726/1024) of

InsertionAllele

DeletionAllele

AC216822

AC216064

1 200 400 600 800 1000

A

B

AC225624

1 200 400 600 800 1000 1200 1400 1600 1800

AC225305

AC203608

C

AC206476

AC212994

1 1000 2000 3000 4000 5000 6000 7000 8000

AC212911D

AccessionNumber of

switches

Conversion

tract (bp)

Variant

Size (kb)

AC225832 4 2,632 27.6AC225305 4 632 6.7AC216797 4 250 8.7AC215992 4 116 16.2AC211399 6 211 10.1AC212994 6 205 3.9AC226182 6 122 108.7AC203608 10 1,249 20.6AC225624 14 454 5.9AC212911 182 7,899 30.7

E

Figure 5. Breakpoint Assessment Using Paralogous

Sequence Variants

(A) Schematic comparison of the structures of the insertion

and deletion haplotypes of a putative NAHR variant. The blue

and red boxes represent homologous sequences present at

the breakpoints, which mediate the rearrangement. The blue

and red vertical lines identify paralogous sequence variants

that distinguish the 50 and 30 copy of the matching sequence.

Scanning along the deletion allele, which is missing the inter-

vening sequence, one observes single nucleotides specific

with the 50 breakpoint, followed by a stretch of sequence

that matches both, then sequences that match the 30 break-

point.

(B) Representation for three variants showing a classic NAHR

pattern. Each line represents the deletion allele corresponding

to the indicated variant. We note a single unexpected paralo-

gous sequence variant mismatch located 145 bp past the 30

breakpoint, which could correspond to a SNP, short gene

conversion, or alignment artifact because of the placement

of indels between 50 and 30 segments.

(C) Representation of four variants having breakpoints that

show a pattern of alternating sequences that match the 30

then 50 breakpoints.

(D) An extreme pattern of alternating matches that contains

182 switches spanning over a 7.9 kb interval.

(E) Rearrangements associated with gene conversion. See

also Figure S3.

the variants are uniquely identifiable with a read

length of 36 bp and uniqueness threshold permit-

ting up to one substitution. This includes 32 inver-

sions—balanced events that are invisible to array-

based genotyping approaches. As read lengths

increase to 100 bp, we estimate that 88% (902/

1024) of these variants could be genotyped. The

construction of complete alternative haplotypes

then facilitates the use of read-pair information to

distinguish among distinct structural configurations

(Antonacci et al., 2010).

Although, short read technologies may miss

some of the breakpoint sequences, there are

many advantages to the application of short read technology

to genome structural variation. This includes the detection of

thousands more events per individual genome, especially vari-

ants below the detection threshold of the fosmid ESP approach.

The dynamic range response and the sequence specificity of

next-generation sequencing allow absolute copy number and

the identity of duplicated genes to be accurately predicted.

One of the strengths of this clone resource, however, is that it

permits the iterative assessment of predicted variants. Clones

may be retrieved corresponding to structural variants discovered

by other methods applied to these 17 individuals, including

newly developed approaches such as methods for identifying

transposon insertions (Huang et al., 2010; Witherspoon et al.,

2010). Sequencing would provide complete information

regarding the structure of additional events, thereby providing

a resource set of sequenced variant haplotypes. The availability

of the underlying clones and potential location of the variant

within a specific DNA sample provides an approach for more

fully exploring the genetic architecture and mutational properties


of these regions. Thus, we predict that such a resource will be

a valuable complement for understanding the true complexity

of human genetic variation as human genomes become routinely

sequenced using short read sequencing technology.


Identifying and Sequencing Variant Clones

Sites of structural variation, relative to the reference genome assembly, were

identified through fosmid ESP mapping. Briefly, genomic DNA was obtained

from transformed lymphoblastoid cell lines (available from the Coriell Cell

Repository) and approximately 1 million 40 kb fragments from each individual

were cloned into fosmid vectors. Paired end sequences were obtained from

both ends of each fragment with standard capillary sequencing. The resulting

ESPs were mapped onto the reference assembly to identify clusters of multiple

clones from a single individual showing the same type of discordancy (Tuzun

et al., 2005). We previously identified 1695 structural variants that have been

experimentally validated (Kidd et al., 2008). In this manuscript, we focus on

1054 events for which complete, finished clone sequence is available. High-

quality finished sequence was obtained for all fosmid inserts with capillary-

based shotgun sequencing and assembly with the procedures established

for sequencing clones as part of the Human Genome Project. Some sequenced

clones contain gaps in simple sequence repeats that are not related to the

detected structural variants. For one individual, NA18956, additional clones

were selected with a relaxed threshold of two standard deviations larger or

smaller than the observed mean insert. In some cases, multiple clones were

sequenced for a single event, whereas in other loci a single clone sequence

appeared to contain multiple distinct variants relative to the genome reference.

Identifying Variant Breakpoints

Sequences of individual fosmid inserts were initially compared to the NCBI

build36 (UCSC hg18) genome reference assembly with the program miropeats

(Parsons, 1995) with a match threshold of �s 400. Images summarizing these

comparisons that included annotations of the repeat content, predicted and

observed segmental duplications (with DupMasker [Jiang et al., 2008]), and

RefSeq exons were prepared and examined to identify clones harboring

a structural difference relative to the build36. Clones that mapped to unas-

signed or random parts of the reference genome or that do not contain an

entire event (such as clones that contain one edge of a tandem duplication)

were omitted from analysis. Approximate variant breakpoints were determined

utilizing the context provided by long stretches of contiguous matching

sequence. In many cases, the pattern of common repeats or segmental dupli-

cations was a useful aid in this assessment.

For each variant, three sequences were extracted and aligned. In the case of

a deletion, two sequences at the variant boundaries are extracted from the

genome assembly and one sequence (termed the deletion junction sequence)

is extracted from the clone. For insertions, the junction sequence is extracted

from the genome assembly and two sequences corresponding to the variant

boundaries in the fosmid clone are extracted. For inversions, a single break-

point is directly captured in the sequenced clone. However, the position of

the other breakpoint can be inferred based on a comparison with the genome

assembly. Thus, for inversions, two sequences are extracted from the

assembly at the edges of the inferred inversion and the third sequence is

extracted from the clone. For inversion analysis, one of the chromosome-

derived segments is reverse-complimented prior to alignment.

An alignment is then constructed from the extracted breakpoint segments

(Kidd et al., 2010). First, an optimal global alignment is computed between

the junction fragment and each of the other two fragments with the program

needle with default parameters (Rice et al., 2000). These alignments are then

merged to yield a single, three-sequence alignment. From this alignment,

the innermost positions that can be confidently assigned to be before and after

the structural variant are identified. The resulting positions are used to define

membership as a class I or class II variant and correspond to the breakpoint

match length depicted in Figure 4. Extended breakpoint homology was deter-

mined with both cross_match (http://www.phrap.org/, -minmatch 4 -max-

match 4 -minscore 20 -masklevel 100 -raw -word_raw) without complexity-

adjusted scoring (Chiaromonte et al., 2002) and bl2seq (-W 7 -g F -F F -S 1

-e 20) to identify the longest extent and identity of additional matching

sequence (termed extended breakpoint homology) that included the two

breakpoints. For putative NAHR events, we additionally determined the

longest stretch of 100% perfect identity as well as a parsimonious matching

metric to account for mutations after the time of variant formation (Figure S2).

VNTR and Retroelement Analysis

Events associated with tandem repeats were characterized with the output

from miropeats (Parsons, 1995), tandem repeats finder (Benson, 1999),

DupMasker (Jiang et al., 2008), and RepeatMasker (Smit et al., 1996–2004).

Potential L1 insertions were characterized with both the TSDfinder program

(Szak et al., 2002) and the results of the breakpoint identification and charac-

terization process.

Genotyping Structural Variants with Diagnostic K-mers

Diagnostic k-mers were identified for each variant (Table S7) by extracting

overlapping k-mers of the indicated size across each sequenced breakpoint.

K-mers were then searched against the build36 genome sequence and a set

of sequenced fosmids with mrsFAST (http://mrfast.sourceforge.net/). To be

considered diagnostic, a k-mer must be unique (within the given edit distance

threshold) to the allele variant from which it was derived (Kidd et al., 2010).

ACCESSION NUMBERS

All sequence data have been deposited in GenBank under project ID 29893.


Supplemental Information includes Supplemental Experimental Procedures,

three figures, and eight tables and can be found with this article online at

doi:10.1016/j.cell.2010.10.027.

790 283

128

5

634

278

84 132

25

76

130

5

Kidd et al. N=1,206

Conrad et al. N=1,128

McCarroll et al. N=236

Figure 6. Comparison of Events Detected from Three Studies

Only variants estimated to be >5 kb are included. The Kidd et al. (2008) set

includes sites of insertion or deletion in one of the five samples relative to

the genome assembly; the Conrad et al. (2010b) set includes gains and losses

in at least one of the five samples relative to a reference arrayCGH sample; and

the McCarroll et al. (2008b) set includes CNVs that were successfully geno-

typed on the Affymetrix 6.0 platform and are variable among the five included

samples. Prior to comparison, the variant sets within each study were merged

into a single, nonredundant interval set, and any overlap among regions

between studies was sufficient regardless of which sample a variant was

detected in.


ACKNOWLEDGMENTS

We thank D. Smith and the staff at Agencourt Biosciences for library produc-

tion, E. Kirkness and staff of the J. Craig Venter Institute for end-sequence data

from the JVCI library, and L. Chen for computational assistance in the mapping

of end-sequence data. We thank S. Girirajan, J. Moran, and C. Payen for

thoughtful discussion; T. Brown for manuscript preparation assistance; and

members of the University of Washington and Washington University Genome

Centers for assistance with data generation. J.M.K. is supported by a National

Science Foundation Graduate Research Fellowship. This work was supported

by the National Institutes of Health Grant HG004120 to E.E.E., who is an inves-

tigator of the Howard Hughes Medical Institute. E.E.E is on the scientific

advisory board for Pacific Biosciences. T.L.N. is an employee and founder of

iGenix Inc.

Received: July 6, 2010




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editorad.indd 1 5/7/2010 12:25:11 PM

Editor: Trends in Molecular MedicineWe are seeking to appoint a new Editor for Trends in Molecular Medicine, to be based in the Cell Press offices in Cambridge, Massachusetts.

As Editor of Trends in Molecular Medicine, you will be responsible for the strategic development and content management of the journal. You will be acquiring and devel-oping the very best editorial content, making use of a network of contacts in academia plus information gathered at international conferences, to ensure that Trends in Molecular Medicine maintains its market-leading position.

This is an exciting and challenging role that provides an opportunity to stay close to the cutting edge of scientific advances while developing a new career away from the bench. You will work in a highly dynamic and collaborative publishing environment that includes 14 Trends titles and 12 Cell Press titles. You will also collaborate with your Cell Press colleagues to maximize quality and efficiency of content commissioning and participate in exciting new non-journal-based initiatives.

The minimum qualification is a doctoral degree in a relevant discipline, and post-doctoral training is an advantage. Previous publishing experience is not necessary—we will make sure you get the training and development you need. Good interpersonal skills are essential because the role involves networking in the wider scientific commu-nity and collaboration with other parts of the business.

To apply Please submit a CV and cover letter describing your qualifications, research interests, current salary, and reasons for pursuing a career in publishing at http://reedelsevier.taleo.net/careersection/51/jobdetail.ftl?lang=en&job=SCI0006D. No phone inquiries, please. Cell Press is an equal opportunity employer.

Applications will be considered on an ongoing basis until the closing date of November 26th, 2010.

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TMM_editorAD.pdf 1 11/2/10 12:19 PM

Cell Press is seeking a Business Project Editor to plan, develop, and implement projects that have commercial or sponsorship potential. By drawing on existing content or developing new material, the Editor will work with Cell Press’s commercial sales group to create collections of content in print or online that will be attractive to readers and sponsors. The Editor will also be responsible for leverag-ing new online opportunities for engaging the readers of Cell Press journals.

The successful candidate will have a PhD in the biological sciences, broad scientific interests, a

fascination with technology, good commercial instincts, and a true passion for both science and science communication. They should be highly organized and dedicated, with excellent written and oral communication skills, and should be willing to work to tight deadlines.

The position is full time and based in Cambridge, MA. Cell Press offers an attractive salary and

benefits package and a stimulating work environment. Applications will be considered on a rolling basis. For consideration, please apply online and include a cover letter and resume. To apply, visit the career page at http://www.elsevier.com and search on keywords “Business Project Editor.”

Cell Press Business Project Editor Position Available

businessprojecteditor.indd 1 8/4/2010 3:00:21 PM

23Brain Research take another look

www.elsevier.com/locate/brainres

One re-unified journal, nine specialist sections, 23 receiving Editors ←Authors receive first editorial decision within 30 days of submission ←

“Young Investigator Awards” for innovative work by a new generation of researchers ←

1

EDITOR-IN-CHIEFF.E. Bloom

La Jolla, CA, USA

SENIOR EDITORSJ.F. Baker

Chicago, IL, USAP.R. Hof

New York, NY, USAG.R. Mangun

Davis, CA, USAJ.I. Morgan

Memphis, TN, USAF.R. Sharp

Sacramento, CA, USAR.J.Smeyne

Memphis, TN, USAA.F. Sved

Pittsburgh, PA, USA

ASSOCIATE EDITORSG. Aston-Jones

Charleston, SC, USAJ.S. Baizer

Buffalo, NY, USAJ.D. Cohen

Princeton, NJ, USAB.M. Davis

Pittsburgh, PA, USAJ. De Felipe

Madrid, SpainM.A. Dyer

Memphis, TN, USAM.S. Gold

Pittsburgh, PA, USAG.F. Koob

La Jolla, CA, USA

T.A. Milner New York, NY, USA

S.D. Moore Durham, NC, USA

T.H. Moran Baltimore, MD, USA

T.F. Münte Magdeburg, Germany

K-C. Sonntag Belmont, MA, USA

R.J. Valentino Philadelphia, PA, USA

C.L. Williams Durham,NC, USA

Twenty-three tothe Power of One.

BresAd23_212X276:Ad 6/3/08 9:15 AM Page 1

cell1435cla.indd 1cell1435cla.indd 1 11/18/2010 10:06:05 PM11/18/2010 10:06:05 PM

Announcements/Positions Available

Columbia University’s CCTI is seeking a qualified Associate Research Scientist who will have significant research responsibilities which include directing the large animal operations/facility. Incumbent will be responsible for the research infrastructure of the CCTI. Will monitor and develop standard operating procedures for the research operation. The candidate is required to have a MD. or Ph.D. in biology with significant research experience in transplantation immunology. Salary offered is $84,000 but will commensurate with experience. Interested applicants should send a CV, letter of interest and names of three references to:

Mayra Marte-MirazColumbia University Medical Center

630W. 168th Street New York, NY 10032

or via email at [email protected].

CUMC is an EOE.


Positions Available

Bowes Research Fellows

University of California Berkeley

The Bowes Research Fellows Program at the University of California, Berkeley, is seeking nominations of outstanding recent or imminent Ph.D. and M.D. graduates to be given the freedom to establish an independent research program as an alternative to the traditional postdoctoral experience. Bowes Fellows must have demonstrated exceptional promise and maturity in their graduate careers and be eager to engage the frontiers of biomedical and life sciences. Fellows will receive funding and space sufficient to maintain a laboratory of two to three members for a term of up to five years, free from the need to obtain grant support or the distractions of classroom teaching. Fellows will have principal investigator status, making them eligible to obtain outside funding from grants or other sources as their research programs expand.

Bowes Fellows benefit from the mentorship of our faculty, as well as from the exceptional breadth of our scientific resources and the highly interactive nature of the Berkeley community. In turn, our community benefits from the creative approaches Bowes Fellows take to solving important problems. Because interdisciplinary interactions are key to innovation, we seek to attract individuals who have broad interests in the life sciences and who have diverse expertise in experimental, theoretical and/or computational approaches.

Candidates must be nominated by their current mentor or by another senior investigator who can provide an in-depth analysis of their accomplishments and future potential. Refer potential reviewers to the UC Berkeley Statement of Confidentiality found at: http://apo.chance.berkeley.edu/evalltr.html.

Selected candidates will be asked to submit a brief research plan and to arrange for additional letters of recommendation. Finalists will be invited to interview on the UC Berkeley campus. Nominations must be received by December 15, 2010 and should be sent to (email submissions are preferred):

Michael EisenChair, Bowes Research Fellows Selection Committee

Department of Molecular & Cell BiologyUniversity of California, Berkeley

Stanley Hall 304BBerkeley CA [email protected]

The University of California is an affirmative action, equal opportunity employer.


EuPA now has its own journal!

To receive more information register at:http://www.elsevier.com/locate/jprot

http://www.eupa.org/

Covered byPubMed

Editor in Chief:Juan J. Calvete, Valencia, Spain

Executive Editors:Proteomics in Cell BiologyJean-Jacques Diaz, Lyon, France

Proteomics in MicrobiologyConcha Gil, Madrid, Spain

Proteomics in Plant SystemsJesus V. Jorrín, Córdoba, Spain

Proteomics in Animal ModelsDario Neri, Zürich, Switzerland

Proteomics in Protein ScienceJasna Peter-Katalinic, Münster,Germany

Biomedical Applications ofProteomics and CongressProceedingsJean-Charles Sanchez, Geneva,Switzerland

Proteomics of Body Fluids andProteomic TechnologiesPier Giorgio Righetti, Milan, Italy

Bioinformatics in ProteomicsPeter Højrup, Odense, Denmark

For a complete listing of theeditorial board, visit thejournal’s homepage

Submitting AuthorsManuscripts can be submitted to the

Journal of Proteomics athttp://ees.elsevier.com/jprot

See online version for legend and references.848 Cell 143, November 24, 2010 ©2010 Elsevier Inc. DOI 10.1016/j.cell.2010.11.026

SnapShot: The SUMO SystemSandrine Creton and Stefan JentschMax Planck Institute of Biochemistry, Martinsried 82152, Germany

Jenstch.indd 1 11/18/10 1:43 PM

careers.cell.com

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