smith_cosb_2015

NMR studies of protein folding and binding in cellsand cell-like environmentsAustin E Smith1, Zeting Zhang3, Gary J Pielak1,2 andConggang Li3

Available online at www.sciencedirect.com

ScienceDirect

Proteins function in cells where the concentration of

macromolecules can exceed 300 g/L. The ways in which this

crowded environment affects the physical properties of

proteins remain poorly understood. We summarize recent

NMR-based studies of protein folding and binding conducted

in cells and in vitro under crowded conditions. Many of the

observations can be understood in terms of interactions

between proteins and the rest of the intracellular environment

(i.e. quinary interactions). Nevertheless, NMR studies of folding

and binding in cells and cell-like environments remain in their

infancy. The frontier involves investigations of larger proteins

and further efforts in higher eukaryotic cells.

Addresses1 Department of Chemistry, University of North Carolina-Chapel Hill,

Chapel Hill, NC 27599-3290, USA2 Department of Biochemistry and Biophysics, University of North

Carolina-Chapel Hill, Chapel Hill, NC 27599-3290, USA3 Key Laboratory of Magnetic Resonance in Biological Systems, State

Key Laboratory of Magnetic Resonance and Atomic and Molecular

Physics, National Center for Magnetic Resonance in Wuhan, Wuhan

Institute of Physics and Mathematics, Chinese Academy of Sciences,

Wuhan 430071, PR China

Corresponding authors: Pielak, Gary J ([email protected]) and Li,

Conggang ([email protected])

Current Opinion in Structural Biology 2015, 30:7–16

This review comes from a themed issue on Folding and binding

Edited by Annalisa Pastore and Guanghong Wei

http://dx.doi.org/10.1016/j.sbi.2014.10.004

0959-440X/# 2014 Elsevier Ltd. All rights reserved.

IntroductionThe cytoplasm is a complex environment where macro-

molecules can reach concentrations greater than 300 g/L

and occupy 30% of the cellular volume [1,2]. Organelles

can be even more crowded [3]. These high concentrations

of macromolecules define the environment where

proteins function. Furthermore, the surfaces of biological

macromolecules (e.g. proteins, nucleic acids) are covered

with groups capable of forming hydrogen bonds and

attractive or repulsive charge–charge interactions [4].

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The crowded nature of the cellular interior puts these

groups in close contact.

Multi-dimensional NMR spectroscopy provides infor-

mation about protein structure, dynamics, stability and

interactions at the level of individual atoms. The main

advantage of NMR for studying crowding effects is that

the introduction of NMR-active isotopes (e.g. 15N, 13C,19F) allows the protein of interest (the test protein) to be

examined in the presence of unenriched crowding agents,

even when the agents are present at biologically relevant

concentrations. The main downside of NMR is its insen-

sitivity; in general, test protein concentrations must be

greater than 10�5 M. This insensitivity means that NMR

detection of proteins in cells relies mostly on promoter-

driven overexpression of test proteins or their delivery

into cells. We use the term in-cell NMR rather than invivo NMR for several reasons: expression levels of test

proteins are hundreds of times larger than native levels,

many of test proteins are not native to the cells in which

they are studied and dense cell slurries can alter intra-

cellular pH [5�].

We focus on high-resolution solution NMR studies pub-

lished since 2011 conducted in cells and under crowded invitro conditions. We have not reviewed NMR-based

studies of membrane proteins under physiological con-

ditions or enzyme activity in cells, but direct the reader to

several recent contributions [6–10].

Quinary interactions and NMRIn 1973, Anfinsen stated that weak surface contacts play a

role in protein chemistry [11]. In 1983, McConkey rea-

lized that such interactions are above primary, secondary,

tertiary and quaternary structure [12]. He coined the term

quinary structure to define the interactions between a

protein and the rest of the intracellular environment.

These contacts organize the cytoplasm and play key roles

in metabolism [13] and signal transduction. Interest in

quinary interactions languished, however, until recent in-

cell studies returned them to the fore [14,15].

Much of the data reviewed here can be understood in

terms of quinary structure. We consider attractive quinary

interactions in terms of binding. This binding is neither as

specific nor as strong as, for instance, the interaction

between a protein-based protease inhibitor and its pro-

tease, but it is binding nonetheless.


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8 Folding and binding

Attractive quinary interactions can adversely affect NMR

spectra. Fast tumbling (i.e. rotational motion) is key to

acquiring high-resolution protein NMR spectra in

solution. Increasing the molecular weight of a globular

protein and increasing the solvent viscosity with small

viscogens degrade tumbling rates, increasing the width of

protein resonances. It was recognized several years ago

that resonances from globular proteins tend to be broader

in cells than they are when the protein is studied in buffer

alone [16] and that broadening did not arise from

increased viscosity alone. For globular test proteins,

especially in Escherichia coli cells, broadening is often

so severe that the test protein spectrum is undetectable.

This situation can lead to dangerous misinterpretations

when spectra attributed to the test protein inside cells are

actually from test protein that has leaked from the cells

into the surrounding media [17]. Along these lines, the

Christodoulou group has recently applied NMR-diffusion

methods to separate signals from inside cells from those in

the surrounding media [18].

The Spicer [19] and Ito [20] groups have identified

important exceptions to the observation that heteronuc-

lear multidimensional spectra from globular proteins in E.coli are too broad to be useful. One of these proteins, the

B1 domain of protein G (GB1, 6 kDa), has come to serve

as a test bed for in-cell NMR [19,21��,23–26].

Lila Gierasch’s lab examined the effects of increased

viscosity on several properties, including the width of

crosspeaks in 1H–15N HSQC spectra of small globular

proteins in simple buffered solutions [25]. The widths

were compared to those obtained for the same proteins in

E. coli cells. The data indicate that the large line widths in

cells cannot be completely explained by a higher

viscosity. The authors concluded that the extra broad-

ening comes from weak transient interactions (i.e. quinary

interactions) with components of the cytosol, consistent

with other observations [27].

Peter Crowley’s group obtained direct evidence for these

interactions by examining HSQC spectra of overex-

pressed cytochrome c (12 kDa) in E. coli lysates (severe

broadening prevented acquisition of in-cell spectra)

[21��]. Size exclusion chromatography as a function of

salt concentration in combination with SDS-PAGE was

used to show that the broadening is due to weak inter-

actions. Mutating positively charged residues to nega-

tively charged residues resulted in crosspeak narrowing

and smaller SDS-PAGE-measured molecular weights,

indicating that the quinary contacts involve interactions

of positively charged residues on the cytochrome with

negatively charged moieties on macromolecules in the

lysate. This important study has placed the focus on

charge–charge interactions. Other types of polar inter-

action as well as hydrophobic contacts may also play key

roles, but systematic studies have yet to be performed.


Volker Dotsch’s group moved work on quinary inter-

actions into eukaryotic cells [28�]. They injected 15N-

enriched protein into Xenopus laevis oocytes and again

found that quinary interactions resulted in broad HSQC

spectra. Specific phosphorylation of one of the proteins

(the 19 kDa WW domain of peptidyl-prolyl isomerase

Pin1) abrogated the interactions, resulting in observation

of the test protein spectrum in oocytes.

Lewis Kay’s group tackled quinary interactions with the

test protein calmodulin (17 kDa) in E. coli lysates. The

system provides information about nonspecific inter-

actions because neither calmodulin nor its specific bind-

ing partners are found in bacteria. The interactions

between components of the cytosol and calmodulin affect

protein dynamics on the ps to ns and ms timescales [29].

They measured a lower bound of �0.2 mM for the

binding constant of calmodulin with cytosolic com-

ponents (Figure 1a,b) [30�]. These results show that

quinary interactions are strong even in the absence of

natural selection. The team also examined what happens

when calmodulin is immersed in yeast lysate. This system

has both biological and physiological relevance because

yeast lysate harbors proteins that specifically bind calmo-

dulin. The specific interactions caused further broaden-

ing resulting in complete loss of the spectrum (Figure 1c).

As discussed above, some proteins have such strong

quinary interactions that tumbling is too slow for solution

NMR. However, lack of rotational motion need not be a

barrier; solid state NMR with magic angle spinning is the

tool of choice under these conditions. Reckel et al. used

solid-state NMR to examine two rotationally challenged

proteins in E. coli cells [31]. Selective 13C-enrichment in

combination with uniform 15N enrichment provided

interpretable spectra. Although solid-state spectra are

not yet as informative as solution spectra, solid state

NMR of biological macromolecules is advancing quickly

[32].

Folding intermediates and crowdingLatham and Kay studied the effects of biologically

relevant crowders on protein folding intermediates

[33��]. They chose an 8 kDa four-helix bundle (the FF

domain) as the test protein and assessed its folding

dynamics in E. coli and yeast lysates (Figure 2a–c).

The same intermediate is found in buffer and lysates,

but it is more highly populated in lysates. Thus, crowding

can affect folding paths, which suggests we may not learn

all there is to know about physiologically relevant folding

by studying proteins in buffer alone.

We close with a warning. A classic approach to under-

standing the solvent-protein friction of folding dynamics is

to quantify the conversion between states as a function of

viscosity. Using the same four-helix bundle, Sekhar et al.[34] showed that a small molecule viscogen (glycerol) and a


Protein NMR in cells and cell-like environments Smith et al. 9

Figure 1

(a) (c)

E. coli S. cerevisiae

(b)

21.5 20

25

1 0 1 0

Val136γ2

142Valγ1

121Valγ1

69Leuδ1

116Leuδ1

4Leuδ1

125Ileδ1

Val108γ2

Val108γ2

22.0

22.513C

(pp

m)

13C

(pp

m)

1H (ppm)

1H (ppm)

R2ef

f (s–1

)

VCPMG (Hz)

23.0

60

50

40

30

20

100 500 1000 1500 2000

0.95 0.90 0.85 0.80 0.75 0.70

Current Opinion in Structural Biology

Equilibrium thermodynamic effects of quinary interactions [30�]. (a) Changes in calmodulin methyl chemical shifts as a function of E. coli lysate

concentration indicate nonspecific interactions with lysate constituents on the fast NMR timescale. (b) Methyl relaxation data showing the higher

viscosity of lysate (100 g/L, orange and green) compared to buffer (red and blue) and the competition between nonspecific binding and specific

peptide binding (green and orange). (c) HSQC spectra showing that specific binding in Saccharomyces cerevisiae (yeast) lysate replaces

nonspecific binding in E. coli lysate resulting in disappearance of the calmodulin spectrum.

macromolecular viscogen (bovine serum albumin) give

different results. The discrepancy arises from the sizes

of the viscogens compared to the folding species; only

small viscogens yield correct values because, for molecules

much smaller than the test protein, the macroscopic

viscosity equals the microscopic viscosity [35].

Disordered proteins are differentThe interior of a globular protein is similar to that of a

billiard ball in that both are rigid. That is, the parts that

comprise the interior of these objects move as a group. On

the other hand, disordered proteins [2] are similar to

spaghetti in that they have local, internal motions that

are independent of global motion. In cells, the internal

motions are not affected to the same extent as the global

motion of globular proteins. Thus, disordered proteins

tend to give higher quality HSQC spectra in cells. Barnes

et al. [2] showed this differential effect in a simple

experiment by fusing the disordered protein, a-synuclein

(14 kDa), to the small globular protein, ubiquitin (9 kDa).

An in-cell HSQC spectrum of the fusion showed only

crosspeaks from the disordered protein. When the cells


were lysed and the lysate diluted, loosening ubiquitin’s

quinary interactions, the spectra of both proteins were

visible. By examining secondary shifts, Waudby et al.showed that a-synuclein appears to remain unfolded even

in E. coli [5�]. By monitoring the shift of the protein’s

histidine resonance, these authors also showed that the

internal pH of E. coli in dense slurries acidified to pH

6.2 instead of the expected physiological pH of 7.6.

Majumber et al. exploited the differential dynamic effect

on globular and disordered proteins to probe binding [36].

First, they induced expression of a disordered protein in15N-containing media and obtained the expected in-cell

spectrum. Next, they induced expression of its unen-

riched globular partner. Binding of the two proteins

hindered the local motion of the previously disordered

protein, broadening its resonances into the background.

Application of principle component analysis even allowed

the authors to define the binding site on the disordered

partner. Amata et al. also exploited the higher resolution

of in-cell spectra of disordered proteins to examine phos-

phorylation of c-Src in frog oocytes in real time [37].



Figure 2

N

(a)

(b)

Buffer

I d

5

400

300

E. c

oli L

ysat

e IΔ

v CP

MG

I (H

z)

200

100

0

400

300

Yea

st L

ysat

e IΔ

v CP

MG

I (H

z)

Buffer I ΔvCPMGI (Hz)

200

100

00 100 200 300 400

4

3

2

1

500 1000 1500

kex ( S–1 )

kex

p I (

%)

2000

1H13Cr.m.s.d. = 18.90 Hz

2500

E. coli LysateS. cerevisiae Lysate

B 1H CPMG RD

(c)

E. coli

1H13Cr.m.s.d. = 35.50 Hz

S. cerevisiae

N

H1

H1

H4

H4

H2

kNI ∼ 30 s–1

N

Fre

e E

nerg

y

Reaction Coordinate

IkIN ∼ 1770 s–1

C

H3

H2


Kinetic effects of quinary interactions [33��]. (a) Free energy diagram of the equilibrium between the native state and the intermediate of the FF

domain. (b) Percent population versus rate plot showing that the intermediate is more highly populated in E. coli lysate and yeast lysate than it is

in buffer. (c) Plots of shift changes showing that lysates do not affect the structure of the intermediate.

Minimizing effects of attractive quinaryinteractions on spectraMethods for obtaining high-resolution spectra of globular

proteins in cells are desirable because attractive quinary

interactions broaden resonances. We discuss two

approaches: 19F incorporation and optimization of con-

ventional multidimensional experiments.

19F is a wonderful in-cell NMR nucleus for eight reasons.

First, fluorine is rare in biological systems [38], which

eliminates background. Second, it is 100% abundant.

Third, the NMR sensitivity of 19F is 83% that of 1H,

making it one of the most sensitive nuclei. Fourth, its

chemical shift range is large. Fifth, as a spin-1/2 nucleus,

its relaxation properties are similar to those of the proton.

Sixth, cells can be coaxed into incorporating aromatic

fluorine-labeled amino acids [22�]. Seventh, replacing a


few hydrogen atoms with fluorine atoms has a minimal

effect on structure and stability [39].

The eighth advantage is subtle. It might be considered a

disadvantage that only a few fluorine-labeled amino acids

are incorporated. However, this situation can be advan-

tageous for studying globular proteins in cells. Simply put,

it is easier to observe a few broad resonances in a one-

dimensional spectrum of a 19F-labeled protein than it is to

observe tens to hundreds of closely packed, broad cross-

peaks in a typical multidimensional spectrum of a protein

uniformly enriched with 15N or 13C [27]. Not only can

‘less mean more’, but also the Crowley lab has shown how

to make 19F labeling of tryptophan residues in E. coli an

inexpensive endeavor by using 5-fluorindole [22�]. The

method represents at 15-fold savings compared to the

usual scheme for labeling aromatic residues, which



involves, for tryptophan labeling, growth of cells in mini-

mal media containing labeled tryptophan, an inhibitor of

aromatic amino acid (glyphosate, i.e. RoundupTM) and

unlabeled phenylalanine and tyrosine [40].

In one of the first studies to demonstrate the effect of

attractive interactions in cells, Schlesinger et al. [41]

exploited 19F labeling to show that crowding is not always

stabilizing, because, as explained in the next section,

attractive quinary interactions can overcome crowding-

induced short-range repulsions. Ye et al. went on to use19F to show that the viscosity in cells is only two-to-three

times that in buffer alone, but quinary interactions make

the protein appear much larger than it is. Their work also

emphasizes the fact that care is required in interpreting

NMR-relaxation derived in-cell viscosity measurements,

because both quinary interactions and viscosity affect

relaxation.

Despite the favorable aspects of 19F labeling, enrichment

with 15N and 13C can yield more detailed information

about structure and dynamics. One way to overcome the

broadening-induced overlap is to use selective enrich-

ment [29,30�]. Latham and Kay showed that incorporat-

ing 13C and 2H methyl-enriched Ile, Leu, Val, or Met

gave interpretable 13C–1H correlation spectra in E. coliand yeast lysates [29,30�]. There are, however, many

combinations of enrichment schemes and NMR exper-

iments. To define a strategy for optimizing in-cell NMR

in E. coli, Xu et al. explored the in-cell NMR of four highly

expressed small (<20 kDa) globular proteins in combi-

nation with several enrichment methods, including

uniform 15N enrichment with and without deuteration,

selective 15N-Leu enrichment, 13C methyl enrichment of

isoleucine, leucine, valine, and alanine (ILVA), fractional13C enrichment, and 19F labeling [25]. 19F labeling gave

acceptable results in all instances, as did 13C ILVA

labeling with one-dimensional 13C direct detection in a

cryogenic probe. ILVA enrichment, however, is expens-

ive and such simple spectra might yield little insight

relative to their cost. Xu et al. suggest that uniform 15N

enrichment and 19F labeling, both of which are low cost,

be tried first. If enrichment yields high quality HSQC

spectra, other approaches will probably work. If an HSQC

spectrum of the protein in cells is not observed, but

reasonable 19F spectra are, ILVA enrichment is worth

trying. If both experiments fail, one might try weakening

protein-cytoplasm interactions by exploring charge-

change variants [21��].

Quinary interactions and globular proteinstabilityThe effects of crowding on protein stability arise from two

phenomena: short range (steric) repulsions and longer-

range interactions. Repulsions stabilize globular proteins

for the simple reason that the crowders take up space,

favoring the compact native state over the expanded


denatured state. Longer-range interactions (also called

soft-interactions or chemical-interactions) include hydro-

gen bonding and charge-charge interactions. Charge-

charge interactions can be stabilizing or destabilizing.

Charge–charge repulsions are stabilizing because they

require the crowders to stand off from the test protein

making them occupy even more space. Attractive inter-

actions between the test protein surface and the crowders,

on the other hand, are destabilizing because unfolding of

the test protein exposes more interacting surface.

NMR-detected amide proton exchange is a tool for quanti-

fying the free energy required to expose backbone amide

protons to solvent [42]. The largest opening free energies

reflect global protein stability, that is, the free energy of

denaturation. Although the method provides opening free

energies at the residue level, we focus on the global free

energy of opening because interpretation of smaller opening

free energies remains controversial [43,44]. Importantly,

exchange experiments provide this information without

the addition of heat or denaturing co-solutes, which is

important for systems containing protein crowders because

it means crowder stability is unaffected by the experiment.

Several controls are required to ensure that the rate data

yield valid equilibrium thermodynamic parameters [45].

We apologize for the number of self-citations in this

section, but a great deal of recent work on NMR-detected

amide proton exchange and crowding has come from the

Pielak lab. That work has been conducted with the small

globular proteins, chymotrypsin inhibitor 2 (CI2, 6 kDa)

and GB1. One of the control experiments mentioned

above is to demonstrate that the crowding agent does

not affect the intrinsic rate of exchange in a disordered

peptide. This assumption is true for a number of crow-

ders, including E. coli lysate [46,47].

Synthetic crowders such as Ficoll and polyvinylpyrroli-

done have minimal attractive interactions with CI2. Thus,

these crowders stabilize CI2 and other test proteins

[35,47,48,49]. CI2, however, is destabilized by protein

crowders and the constituents of E. coli lysate [50]. The

destabilization suggests the presence of attractive crow-

der-test protein interactions, which is consistent with

other NMR experiments on CI2 [35]. Inomata et al.[51] have observed an increase in amide-proton exchange

rates for the test protein ubiquitin in human cells, a result

that is also consistent with the presence of attractive

quinary interactions.

Sarkar et al. hypothesized that the interactions involve

charge–charge contacts. To test this idea, they refined the

lysate by removing the anionic nucleic acids and nega-

tively charged proteins; CI2 was still destabilized [52].

The result indicates that there remains much to be

learned about longer-range interactions and crowding.

Sarkar and Pielak have also shown that the osmolyte



glycine betaine can mitigate the lysate’s destabilizing

effect on CI2 [53].

Monteith et al. [54�] used amide proton exchange to

measure the stability of GB1 in living E. coli cells

(Figure 3). There are two reasons one might think this

would be an easy experiment. First, GB1 is one of the few

globular proteins whose 1H–15N HSQC spectrum can be

Figure 3

(a)

(b)

harvest

repeat n times ove

cell pellet

re

HSQCspectrum

8.0

BSA

buffer

cells

7.0

6.0

K4 L5 I6 L7 A26 E27

ΔGo

’ op (

kcal

/mo

l)

quenchedlysate

pH < ice co

15N enriched GB1

Quantifying residue-level protein stability in living cells [54�]. (a) Procedure u

graphs of the free energy of opening versus residue number of GB1 showin

destabilized in a 100 g/L solution of bovine serum albumin (BSA).


directly observed in E. coli [19]. Second, D2O diffuses

quickly into cells [55]. Unfortunately, the low signal-to-

noise ratio of the in-cell HSQC spectra prevents quanti-

fication of exchange rates. Instead, the authors employed

a quenched-lysate method [55]. The authors showed that

GB1 is stabilized in cells compared to buffer by between

0.5 and 1.1 kcal/mol, which is significant given the free

energy of unfolding in buffer is 7 kcal/mol.

r 24 h

suspend inD2O

resuspend inquench buffer

aliquot/pellet

V29

Residue

Y33 T44 D46 T51 F52 T53

4,ld lyse cells

H2O

D2O

incubate at37 °C


sed to acquire in-cell NMR-detected amide 1H exchange data. (b) Bar

g that the protein is stabilized with respect to buffer in cells, but



The increased stability of GB1 in cells appears to be

related to protein charge, but the story is not yet clear. As

mentioned above, the majority of proteins in E. coli are

anionic, and so are CI2 and GB1. The resulting net

repulsive interactions should allow both proteins to retain

their rotational motion resulting in acceptably narrow

crosspeaks [21��]. The increased stability of GB1 in cells

compared to buffer suggests net repulsion between the

test protein and the cytoplasm. This increased stability of

GB1 in cells is consistent with the fact we can observe its

HSQC spectrum in cells [19,21�,24–26]. CI2 and GB1,

however, show different responses to the cytosol. That is,

even though CI2 is also negatively charged, its 15H–1H

HSQC spectrum cannot be observed in E. coli [17] and

the protein is destabilized by reconstituted cytosol

[50,52]. This differential response of similarly charged

proteins to the cytosol shows that we still have a great deal

to learn about assessing quinary interactions.

NMR in eukaryotic cellsEukaryotic cells are at the cutting edge of in-cell NMR.

Although E. coli is a good test bed because it is easy to

grow and can be coaxed into expressing the large amounts

of test protein required for NMR, the more complex

features of eukaryotic cells make them an important

subject.

Insect cells and the yeast Pichia pastoris are commonly

used to express large quantities of eukaryotic proteins.

Bertrand et al. tested this yeast as a subject for in-cell

NMR using ubiquitin as the test protein [56]. Their

results are especially informative about what can happen

when proteins are highly expressed. The authors com-

pared 1H–15N HSQC spectra after expression from a

methanol inducible promoter under two conditions.

Methanol as the carbon source gave high expression

and usable spectra. A combination of methanol and

glucose gave even higher expression, but the ubiquitin

was trapped in expression vesicles, making it essentially a

solid. The resulting diminution of rotational motion

broadened its resonances into the background. Hamatsu

et al. have shown that insect cells are also amenable to in-

cell protein NMR with a baculovirus expression system,

reporting both 15N–1H HSQC spectra and three dimen-

sional spectra of four small globular proteins, including

GB1 and calmodulin [23]. Thus, these eukaryotes should

prove useful for in-cell NMR, especially for studies of

eukaryotic-specific post-translational modifications.

Higher eukaryotic cells are fussier than E. coli. For

instance, E. coli can be examined under anaerobic con-

ditions, but most higher-eukaryotic cells are sensitive to

nutrient depletion and require oxygenation. Kubo et al.developed a clever bioreactor that solves these problems

[57��]. Cells are fixed in a gel in the sensitive part of the

probe where they can be easily perfused with fresh

oxygenated media (Figure 4). The authors tested the


bioreactor by using 31P NMR to examine ATP levels.

Without the bioreactor, ATP depletion was complete in

30 min. With the bioreactor, cells maintained their ATP

levels for the duration of the experiment (five hours).

They also investigated the in-cell 1H–13C HMQC spectra

of a 9-kDa methyl ILV 13C-enriched microtubule binding

protein. The protein was introduced into the cells via a

pore-forming toxin. Transferred cross saturation exper-

iments and competition experiments utilizing unen-

riched protein helped define the binding site and

showed specific microtubule binding. Importantly, the

binding site in cells is similar to that discovered invitro. These experiments also illuminate an important

point about protein leakage. In the absence of the bio-

reactor only 10–20% of the protein leaked, yet almost all

the signal arose from the leaked protein because the

quinary interactions in cells led to severe broadening.

The Banci group has characterized the maturation of the

mitochondrial intermembrane space (MIMS) protein,

Mia40 (16 kDa) in human cells [58]. To reach their

destination, MIMS proteins must be transported through

the outer membrane in a disulfide-reduced, unfolded

state. Overexpression resulted in accumulation of the

fully folded form in the cytoplasm because the transporter

and reductive apparatus were overwhelmed. This lack of

a physiological outcome with a physiologically relevant

protein is one reason why such studies should be called

in-cell, not in vivo, NMR. As expected, simultaneous

overexpression of the reductive machinery resulted in

more unfolded, ready-to-be-transported protein.

Banci et al. also investigated the maturation of superoxide

dismutase (SOD) in human cells [59]. The wealth of

detail available from their spectra is remarkable. The

authors could detect the monomeric partially unfolded

apo form, the Zn-bound monomer and dimer as well as

the Cu-Zn form. Furthermore, both oxidized and reduced

cysteines could be detected with [15N] cysteine enrich-

ment. This group also attached a sequence to target

Mia40 and superoxide dismutase for import into the

MIMS [60�] and showed it is possible to acquire high-

resolution data from proteins in the inter-membrane

space of intact mitochondria.

Continuing with SOD, Danielsson et al. took a different

approach [61]. Instead of expressing the protein in human

cells, they expressed and purified the isotopically

enriched protein from E. coli and covalently attached a

cell penetrating peptide to an engineered cysteine on

SOD. Initial attempts to introduce sufficient amounts

into the cell were unsuccessful. Success came after

increasing the positive charge on SOD. The charge

changes make sense considering our knowledge of cell

penetrating proteins: they must overcome the negative

charge on the test protein to which they are attached.

Another key point involving a pre-enriched protein [57��]



Figure 4

(a) (b)

(c)

10

1.0

0.5

0

1 2 3 4

Time / h

Nor

mal

ized

inte

nsity

of A

TP

βP

1 mm × 2 mm

0.5 mm × 1 mmAspirator

Trap

Sealed bySealed byParafilmParafilm

4.2 mm x 5.0 mm4.2 mm x 5.0 mmNMR tubeNMR tube

2.6 mm x 4.0 mm2.6 mm x 4.0 mmglass tubeglass tube

Glass capillaryGlass capillary

NMR magnetNMR magnet

HeLa cells immobilizedHeLa cells immobilizedby Mebiol gelby Mebiol gel

4 mm × 6 mm

DMEM injection(using syringe injector) Halting DMEM

Injection

5 6 7

15

*

*

** *

**

***

** **

*

*

20δ (13

C)

/ ppm

δ (1H) / ppm

251.0 0.6

Supernatantw/o bioreactor

Supernatantwith bioreactor

0.2

δ (1H) / ppm

1.0 0.6 0.2


(a) Bioreactor for protein NMR in eukaryotic cells [57��]. (b) Plots of relative ATP concentration versus time showing that the bioreactor maintains

ATP levels. (c) TROSY spectra showing that the bioreactor prevents the test protein from leaking into the media.

is that the NMR-active nuclei are less likely to find their

way into metabolites and other proteins that obscure the

test-protein spectrum.

We close this section by comparing the quality of in-cell

spectra from bacterial and eukaryotic cells. With rare

exceptions [19,20], multidimensional spectra from glob-

ular proteins are difficult to observe in E. coli. As dis-

cussed above, the situation is better in eukaryotic cells. It

is not entirely clear why this should be the case, but one

clue may be the lower concentration of protein plus

nucleic acid in eukaryotic cells (100–300 g/L) compared

to E. coli cells (300–500 g/L) [2]. It is important to bear in

mind, however, that eukaryotic cells are far from a pana-

cea because not all proteins that should be observable by

in-cell NMR in eukaryotes are observed [58]. The differ-

ence between what should happen and what does happen

extends to methods for getting isotopically enriched

proteins into higher eukaryotic cells. Besides injection,

which is only reasonable for oocytes [28�,37], the most


common methods are attachment of a cell-penetrating

peptide, use of a toxin to open pores in the membrane and

electroporation [62]. It is unclear, however, which method

will work best in any particular combination of test

protein and cell.

Summary and closing thoughtsNMR has taken the lead in adding to our understanding

of how quinary structure affects protein stability, folding

pathways, ligand binding and side chain dynamics. Se-

lective labeling strategies have been shown to be useful

when these interactions broaden protein resonances into

the baseline. In addition, sophisticated NMR strategies

are being applied to not only E. coli proteins, but also

eukaryotic cells. Nevertheless, the field remains in its

infancy. We are picking the low hanging fruit: typical test

proteins have molecular weights of approximately

10 kDa, but the average molecular weight an E. coliprotein is 30 kDa. The average eukaryotic protein is even

larger [63]. In addition, we need models to explain how



proteins of similar size, shape and surface properties

interact differently with the cytosol. The most important

and challenging frontier is the study of larger test

proteins, because crowding effects are predicted to scale

with molecular weight [64].

Conflict of interest statementNothing declared.

Acknowledgements

The research in our laboratories is supported by the Ministry of Science andTechnology of China (2013CB910200 to C.L.), the 1000 Young TalentsProgram (C.L.), the National Natural Science Foundation of China(21075134 to C.L.) and the U.S. National Science Foundation (MCB-1051819 and MCB-1410854 to G.J.P.). We thank Elizabeth Pielak andRachel D. Cohen for comments on the manuscript.

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