chapter 13 organelle motilitysun-lab.med.nyu.edu/files/sun-lab/attachments/cpcb... · providing a...

CHAPTER 13Organelle Motility

INTRODUCTION

T he cytoplasm of eukaryotic cells is comprised of a complex network of filaments,including microtubules and actin, that extend throughout the cytoplasm. These fila-

ments are highly dynamic and continuously remodel as the cell changes shape, divides,and interacts with the environment. They play a key role in organizing the cytoplasm byconnecting protein compartments and organelles in different regions of the cell. Micro-tubules, for example, are known to provide communication paths between organelles,controlling the spatial location of these structures. Actin filaments, on the other hand,form contractile bundles with myosin and mediate muscle contraction and contractile-ring formation during cytokinesis. In this chapter, assays are described for examiningthe characteristics and functions of the microtubule and actinomyosin filament systems.These assays are useful for identifying accessory proteins that attach microtubules toorganelles and/or modulate sliding of myosin molecules across actin fibers.

UNIT 13.1 describes an in vitro assay for examining interactions between microtubulesand isolated organelles. Microtubules are polar structures with their plus (i.e., rapidlygrowing) ends extending out to the cell periphery, and their minus ends associated withthe centrosome adjacent to the nucleus. They emanate as a star-like network out from thecentrosome, with new microtubules constantly extending out to replace old ones that havedepolymerized. The endoplasmic reticulum (ER) uses microtubules to move out to the cellperiphery, while membranes of the Golgi complex cluster inwards toward the centrosomealong microtubules. These organelle movements are mediated by microtubule-associatedmotor proteins, including kinesin and cytoplasmic dynein. The in vitro motility assaydescribed in this unit provides a useful system for studying organelle transport alongmicrotubules. It can be used for dissecting the molecular machinery involved in suchmovement, including proteins that regulate microtubule motors or that cross-link motorsto membranes. Effects of pharmacological or biochemical perturbations on microtubulestructure and dynamics can also be investigated. In the assay described in UNIT 13.1,microtubules are nucleated from a stationary point (i.e., the flagellar axoneme), allowingtheir plus ends to grow and shorten in a way analogous to what happens in normalcells. Both motor-driven motility of organelles and microtubule polymerization–drivenmovement of organelles can be studied. The unit also provides detailed methods forpreparation of purified tubulin, cytosol enriched in motor proteins, and axonemes fromsea urchin sperm, as well as for isolation of membrane-bound organelles. The assay isperformed in a simple perfusion chamber and the results are visualized using video-enhanced DIC microscopy.

UNIT 13.2 provides a motility assay for transport of actin by myosin. F-actin filamentslabeled with rhodamine-phalloidin are imaged in an in vitro system that contains puri-fied myosin, an ATP-activated motor protein that hydrolyzes ATP to ADP and Pi whenstimulated by binding to actin filaments. This assay uses two purified proteins, actin andmyosin, to study the contractile forces produced by the sliding of actin filaments alongthe myosin filaments. During muscle contraction, the head regions of myosin moleculesengage in an ATP-driven cycle in which they attach to adjacent actin filaments, undergoa conformational change that pulls the myosin filament, and then detach. This results in

Contributed by Jennifer Lippincott-SchwartzCurrent Protocols in Cell Biology (2005) 13.0.1-13.0.2Copyright C© 2005 by John Wiley & Sons, Inc.

Organelle Motility

13.0.1

Supplement 27

Introduction

13.0.2

Supplement 27 Current Protocols in Cell Biology

the sliding of actin filaments against the myosin, which can be directly visualized andquantified in this assay.

The use of GFP fusion proteins has revolutionized the study of organelle motility byproviding a method to visualize diverse organelles directly under the microscope. Virtu-ally any protein can be tagged with GFP, expressed in cells, then visualized by applyingblue light. UNIT 13.3 describes how to express and image GFP fusion proteins in plants inorder to study organelle dynamics in these cell types. Both transient and viral-mediatedexpression methods are described for expressing GFP fusion proteins. Tips are givenfor imaging GFP proteins targeted to the endoplasmic reticulum and Golgi complex,two organelles that function in secretory transport. Solutions to common problems withexpression and imaging GFP fusion proteins–e.g., misfolding and autofluorescence–arealso discussed.

Nuclear migration (UNIT 13.4) occurs in a wide variety of cell types, including newlyfertilized eggs, muscle, nerves, and dividing cells. It is vital for the cell to properlylocate its nucleus and surrounding endomembrane system. This unit describes an in vitroassay for monitoring nuclear motility of the pronucleus from frog eggs. This motilityis, as are other examples of nuclear motility, driven by microtubule-dependent motility.Purified nuclei are added to a motility extract that includes Xenopus cytosol, centrosomes,purified microtubules, dynein, and regulatory factors. The system is then visualized usingvideo enhanced-differential interference contrast microscopy. Once they associate withmicrotubules, the nuclei normally move toward the centrosomes in microtubule minusend-directed (due the dynein in the extract) motility. There they accumulate over time.Regulators or inhibitors of this movement can be assayed, and the effects on motility canbe quantified using this system.

UNIT 13.5 describes dynamic live cell imaging approaches employing photobleaching tomeasure the kinetics of nuclear proteins. Three major methods of photobleaching mi-croscopy are described fluorescence recovery after photobleaching (FRAP), fluorescenceloss in photobleaching (FLIP), and inverse fluorescence recovery after photobleaching(iFRAP). The unit highlights how each technique differs in permitting the determinationof distinct particular parameters of protein behavior in vivo. In addition to describing thephotobleaching techniques, this unit details transfection methods for introducing GFP-expression vectors into mammalian cells by electroporation or by lipofection. Moreover,it provides a method for the determination of the number of GFP molecules in a single,living cell, which is critical for the quantification of GFP-fusion proteins.

UNIT 13.6 describes a number of in vitro assays for characterizing the function of putativeactin-binding proteins. One assay described uses a change in fluorescence of pyrenyl-labeled actin to assess a protein’s actin capping activity. The read-out of this assay includeskinetic measurements of filament growth or depolymerization at one or the other end of theactin filament. Another assay describes assays of filament growth from actin seeds. Thisallows proteins that interact with G-actin to be analyzed, and permits the determination ofwhether the actin association rate constant for filament assembly is the same or differentfrom that of free G-actin. The unit also provides protocols for measuring the turnover ofa population of filaments in vitro using a fluorescent analog of ATP whose fluorescencechanges when bound to actin. It further describes mathematical modeling approaches forunderstanding the mechanism by which the basic features of actin spontaneous assembly(nucleation-growth process) are modified by the severing or branching agents. Finally,the unit highlights light microscopy approaches that can be used to observe the dynamicsof individual actin filaments, including total internal reflection fluorescence microscopy(TIRF).

Jennifer Lippincott-Schwartz

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UNIT 13.3Organelle Motility in Plant Cells: ImagingGolgi and ER Dynamics with GFP

This unit describes the use of green fluorescent protein (GFP) from the jellyfish Aequoriavictoria as a vital marker for endoplasmic reticulum (ER) and Golgi apparatus (GA) inhigher plant cells. GFP is expressed as a chimeric protein either with selected peptidetargeting sequences or with complete proteins that are resident in the ER or GA.Expression of GFP constructs can be observed in living material using conventional, orpreferably, confocal laser−scanning epifluorescence microscopes. GFP has a number ofintrinsic properties that make it attractive as a marker for cell biological studies. Forinstance, it is highly fluorescent, yet requires no cofactors or substrates. In addition, it isrelatively stable, is pH insensitive, shows no detectable level of cytotoxicity (whenexpressed in organelles), and in many instances, does not interfere with the functioningof native proteins when expressed as chimeric constructs. Moreover, there are geneticallymodified variants of GFP with altered spectral and other properties, such as folding rate(Cubitt et al., 1995).

This unit describes two protocols for rapid transient expression of GFP in the plantendomembrane system using Agrobacterium and virus vectors. These protocols can beused for the expression of GFP targeted to most organelles. Besides Agrobacterium-me-diated transformation (see Basic Protocol), virus transformation (see Alternate Protocol),has been optimized for GFP expression in some Nicotiana species.

BASICPROTOCOL

TRANSIENT EXPRESSION FOR VISUALIZATION OF ER AND GOLGIPROBES IN LEAVES

This protocol describes the production of GFP-transcripts in Nicotiana using an Agro-bacterium tumefaciens–mediated transient expression system. There are a number oftechniques for transient expression of reporter constructs in plant cells, such as electro-poration and PEG-mediated transformation of protoplasts, and microprojectile bombard-ment and virus-mediated expression in tissues; however, transient expression mediatedby Agrobacterium containing a reporter construct in a suitable binary vector (e.g.,pVKH18En6) is one of the easiest, quickest, and more reliable methods. It requiresminimum laboratory equipment, and it allows the study of intact cells and tissues fromleaves still attached to the plant with minimal tissue disruption, although it is not clear towhat extent the bacterial infection alters cell physiology. Moreover, transformed leaftissue may be used to generate stable transformants.

The basic principle of the transformation is infiltration of the intracellular spaces ofNicotiana leaves with a suspension of A. tumefaciens, which is injected into the leaf tissueby pressure though the stomata of the abaxial leaf epidermis. The infectious Agrobac-terium cells then transfer the T-DNA carrying the genes to be transferred to the host cellresulting in a transient build-up of reporter transcripts in the cell. With GFP-basedreporters, fluorescence can be detected in the infected area from abaxial and adaxialepidermis, guard cells, and palisade and spongy mesophylls, but rarely trichomes andvascular tissue. The technique gives high expression levels of GFP constructs, which mayfade in a week to ten days after inoculation of the leaves due to progressive depletion ofthe foreign DNA.

This method offers a number of advantages including speed and relatively little variabilitybetween cells within the infected area in which most, if not all cells, appear to expressthe construct. Moreover, Rossi et al. (1993) have shown that expression from T-DNA

Supplement 9

Contributed by Chris Hawes, Federica Brandizzil, Henri Batoko, and Ian MooreCurrent Protocols in Cell Biology (2001) 13.3.1-13.3.10Copyright © 2001 by John Wiley & Sons, Inc.

13.3.1

Organelle Motility

varies linearly with bacterial concentration over three orders of magnitude offering theopportunity to control GFP expression levels. Finally, the geometry of epidermal cells,which have large vacuoles and a thin layer of cytoplasm immediately below the cuticularwall, greatly facilitates observation by both conventional epifluorescence microscopy(UNIT 4.2) and confocal imaging (UNIT 4.5). For example, the authors have been able todistinguish the distribution of ER- and Golgi-targeted GFP in these cells using a conven-tional epifluorescence microscope.

Materials

Suitable Agrobacterium vector (e.g., pVKH18En6) with multiple cloning site andappropriate selectable marker

Agrobacterium tumefaciens (e.g., GV3101::pMP90)YEB medium containing appropriate selective antibiotic (see recipe)5% (v/v) sodium hypochlorite or 1% (w/v) Virkon (Amtec Int. Ltd.)Infiltration medium (INM; see recipe)Four-week-old greenhouse plants of Nicotiana tabacum, N. clevelandii, and N.

benthamiana

Shaking incubator, 28°C1.5-ml microcentrifuge tubes, sterileSpectrophotometer1-ml disposable plastic syringe, without needle22° to 25°C greenhouse for growing plantsPermanent marker penFine scissorsSlides and coverslips (use thickness 0 for confocal microscopes)Electrical or waterproof tapeConventional epifluorescence microscope, laser-scanning confocal microscope

(preferred), or equivalent, with appropriate filters (e.g., standard FITC filterblock)

Confocal-imaging time-lapse software (Zeiss, Leica, Biorad)

Additional reagents and equipment for vector construction, transformation, andcell culture (see APPENDIX 3) adapted for plants

CAUTION: All solutions used to culture and wash bacteria should be treated with asuitable disinfectant before discarding in an autoclavable waste container. Resulting plantmaterial should be treated as biological hazard and handled accordingly.

NOTE: Sterile conditions are required for culture and handling of Agrobacterium, but notduring tobacco leaf infiltration and subsequent incubation of the plant. All equipmentcoming into contact with bacteria should be autoclaved.

Clone GFP into bacteria and grow Agrobacterium1. Using standard molecular biology techniques (see APPENDIX 3), clone GFP chimera

into a suitable vector with an appropriate selectable marker (e.g., kanr, ampr) andtransform into a suitable Agrobacterium strain.

A number of binary Agrobacterium vectors can be used for the cloning of GFP (Bevan,1984). In the authors’ laboratories a vector with an enhanced 35S promoter, pVKH18En6,is used to drive high levels of GFP expression (Hawes et al., 2000).

2. Pick a single colony of Agrobacterium from a selection plate and inoculate 2 to 5 mlYEB medium containing appropriate selectable antibiotic.


13.3.2

Imaging PlantGolgi and ER

with GFP

3. Culture Agrobacterium at 28°C in a shaking incubator to stationary phase (24 to 48hr).

4. Transfer 1.0 ml of culture to a 1.5-ml sterile microcentrifuge tube and pellet bacteriaby microcentrifuging for 5 min at 4000 rpm.

5. Discard the supernatant in a disinfecting solution of either 5% (v/v) sodiumhypochlorite (NaOCl3) or 1% (w/v) Virkon.

6. Wash the bacterial pellet twice with 1.0 ml INM each.

7. Microcentrifuge for 5 min at 4000 rpm.

8. Resuspend in INM to OD600 0.5 to 0.6 or higher to increase the level of expression.

Inoculate plants 9. Inject bacterial suspension into the abaxial epidermis of plant leaves from a 1-ml

disposable plastic syringe by simply pressing the nozzle against the leaf surface (donot use a needle), holding the leaf on the other side with a gloved finger at the pointof contact to support the pressure.

The infiltration is easier when the leaf stomata are open. Illuminating the plants fromunderneath with a bright light prior to infiltration is recommended.

The spread of liquid entering the leaf via stomata is visualized by a darkening of the leaftissue. The boundaries of the infiltrated area should be outlined with a permanent markerpen.

Expression can also be achieved in whole Arabidopsis plants by immersing plantlets inbacterial suspension under vacuum (Rakousky et al., 1998).

10. Incubate plants for 2 to 3 days under normal growing conditions in a greenhouse,except at 20° to 22°C to optimize infection.

11a. For observation with a UV lamp: Check for GFP fluorescence in leaves with ahand-held long-wavelength UV lamp after 2 days (assuming a GFP variant with aUV-excitation peak).

The time required for the fluorescence to appear may depend on the construct. It is wise tocheck a piece of leaf tissue 2 days after the inoculation of Agrobacterium cells into the leaf,and every 4 to 6 hr thereafter, to establish the optimal time for expression of each constructused.

11b. For observation using a microscope: After 2 days, cut out a segment of leaf tissuewith fine scissors, mount in a drop of water on a microscope slide, and cover witha long coverslip held in position with strips of electrical or waterproof tape at eitherend.

It is important to remember that GFP fluorochrome cyclization will not occur in an anoxicenvironment so it is preferable that microscope preparations are not sealed.

Visualize expression 12. Observe the specimen with a conventional epifluorescence microscope or laser-

scanning confocal microscope (preferred) and appropriate filters.

Most forms of GFP can be observed with a standard FITC filter block; however, the exactfilter configuration will depend upon the excitation and emission wavelengths of the GFP(or its spectral variant) used.

Depending on the levels of expression, for ER- or GA-targeted constructs, image captureby confocal microscopy may require high laser power. This is possible with GFP as it isrelatively resistant to photobleaching. Moreover, a wide pinhole aperture may be required

Current Protocols in Cell Biology Supplement 9

13.3.3

Organelle Motility

to obtain a satisfactory signal-to-noise ratio. Although, this strategy reduces confocality,and the ultimate resolution of the image, it can help overcome problems of image blurringdue to organelle movement during the image capture process.

13. (Optional) To make a confocal video of endomembrane dynamics, select a region ofinterest, increase the laser power to the maximum available, select a wide pinholeaperture, and collect images as fast as possible using confocal-imaging time-lapsesoftware.

If cytoplasmic movement is not dramatic then line (not sequential) image averaging canbe used to improve the signal-to-noise ratio.

ALTERNATEPROTOCOL

VIRUS MEDIATED EXPRESSION OF GFP CONSTRUCTS

Viruses are often used as vectors for transient expression of DNA in foreign organisms,due to the high level of replication of the virus and the high level of protein synthesis(results may be obtained 4 to 10 days after inoculation of leaves of the host plants). Ingeneral the level of protein expression is high because of the activity of the strong viralpromoter. The limitation associated with the virus expression technique resides in the factthat not all cells in plants expressing the viral genome become infected. This may in turnlimit the reproducibility of biochemical assays. Moreover, the high levels of expressionof the desired protein may be a reason for their mistargeting in the cell. Some inserts maybe unstable in the vector, and the likelihood of instability increases with the size of theinsert, such that 3 kb may be close to the practical limit. Finally, the pathogenic effectsof the viral infection must be taken into account.

For plants, potato virus X (PVX) has been used successfully to express the marker proteinsGFP and β-glucuronidase (GUS; Chapman et al., 1992; Baulcombe et al., 1995), andnumerous other foreign proteins, often as fusions with GFP (Blackman et al., 1998;Boevink et al., 1998, 1999). The foreign genes inserted into the multiple cloning site areunder the transcriptional control of the subgenomic promoter. A site at the 3′ end of thePVX sequence is used to linearize the plasmid before infectious run-off transcripts aremade with T7 RNA polymerase (the T7 promoter is present immediately upstream of thePVX sequence).

Additional Materials (also see Basic Protocol)

GFP chimera constructPVX vector (pTXS.P3C2) with multiple cloning site and T7 promoter (available

from various laboratories)Midiprep kit, without RNase (Qiagen)T7 RNA polymerase transcription kit (e.g., Ambion T7 message in machine kit)SpeI or SphI restriction enzymes (see APPENDIX 3)

Aluminum oxideAluminum oxide dispenser: small glass flask containing abrasive, sealed with

miracloth

Additional equipment and reagents for restriction digestion, phenol/chloroformextraction, and ethanol precipitation (see APPENDIX 3), and quantification of DNAconcentration by spectroscopy (see APPENDIX 3D)

IMPORTANT NOTE: In some countries in order to handle plant viruses as vectors, alicense must be obtained from the appropriate authorities.

IMPORTANT NOTE: All solutions used for virus work must be RNase-free.


13.3.4


with GFP

1. Insert the GFP chimera in the multiple cloning site of the PVX vector in the correctorientation with respect to the T7 promoter (see APPENDIX 3).

2 Prepare a high-quality, medium-scale, RNase-free DNA preparation using a midiprepkit. Avoid using RNase in preparation of the DNA (i.e., do not add RNases to solutionP1 and increase the volume of the wash buffer as recommended by the manufacturer).

3. Linearize the vector with SpeI or SphI restriction enzymes to make infectious RNA.

4. (Optional) Clean the DNA by phenol/chloroform extraction and ethanol precipitation(see APPENDIX 3).

5. Synthesize capped transcripts using 0.2 to 1.0 µg linearized DNA for a singleinfection, as template for the T7 RNA polymerase transcription kit, following themanufacturer’s instructions.

The transcripts are generally not phenol/chloroform extracted or precipitated beforeinoculating.

6. Dust host plants lightly with abrasive aluminum oxide using an aluminum oxidedispenser.

7. Drop 5 µl of transcripts (1 to 5 µg/µl) onto four leaves and spread lightly with aglove-protected fingertip.

8. After inoculation wash the abrasive off the leaves. Keep plants at ∼23°C in a cultureroom or sealed greenhouse.

9. Four to seven days post inoculation with PVX-GFP virus, screen plants for smallfluorescent lesions on inoculated leaves with a hand-held long-wavelength UV-lightor an epifluorescence microscope.

10. Analyze as described elsewhere (see Basic Protocol, steps 11 to 13).

REAGENTS AND SOLUTIONS

Use deionized or distilled water in all recipes and protocol steps. For common stock solutions, seeAPPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.

Infiltration medium (INM)In a 10-ml measuring cylinder, add 50 mg glucose, 1 ml 0.5 M MES, pH 5.6 (seerecipe), 1 ml 20 mM Na3PO4 (see recipe), and 5 µl 200 mM acetosyringone (Aldrich)in dimethyl sulfoxide (DMSO; final concentration 100 µM). Bring volume to 10 mlwith water, seal the cylinder with 1 cm2 Parafilm, and mix thoroughly by inversion.Make fresh prior to use.

200 mM acetosyringone in DMSO and 20 mM Na3PO4 can be stored at 4°C up to two weeks,or aliquoted and kept at –20°C for longer (i.e., 6 months) storage.

NOTE: No antibiotic must be added to this medium.

MES, 0.5 M, pH 5.6In a 20-ml beaker add 1.95 g MES and 18 ml water. Adjust the pH to 5.6 with KOHand bring the volume to 20 ml with water.Store up to 2 weeks at 4°C, or for longer storage (i.e., 6 months), aliquot and storeat −20°C.

Na3PO4, 20 mMDissolve 0.152 g Na3PO4⋅12H2O in 20 ml water. Store up to 2 weeks at 4°C, or forlonger storage (i.e., 6 months) aliquot and store at −20°C.


13.3.5

Organelle Motility

YEB mediumIn a 1-liter beaker, dissolve 5 g beef extract (Difco), 1 g yeast extract (Merck), 5 gpeptone (Difco), 5 g sucrose (BDH Laboratory), and 0.5 g MgSO4⋅7H2O, in 0.8 litersof water. Add water to 1 liter, stirring continuously. Autoclave and store up to 6months at room temperature.

It may take some time to dissolve completely.

COMMENTARY

Background InformationWhile many experimental programs may

ultimately require the production of stabletransformants, much useful information can beobtained through the use of transient expressionsystems. The production of transgenic plantsand culture cell lines expressing GFP can beachieved using standard transformation andgene expression technologies. One of the majoradvantages of GFP and its wavelength-shiftedderivatives, when used to tag proteins of inter-est, is that cellular events can be studied in vivoin planta and monitored by conventional epi-fluorescence microscopy. However, with tissuesamples it may be necessary to use either con-focal microscopy or conventional fluorescencemicroscopy combined with low-light-levelcameras and deconvolution software.

The introduction of jellyfish fluorescentprotein has provided cell biologists with a pow-erful new tool with which to study cell structureand function in vivo.

GFP chimeras were first used in plant cellbiology to investigate the movement of virusesby the construction of both fluorescent virusesand fluorescent viral proteins, such as themovement proteins (Baulcombe et al., 1995;Oparka et al., 1995; Itaya et al., 1997). Sub-sequently, most major organelles have beensuccessfully labeled with GFP chimeras, in-cluding the nucleus (Grebenok et al., 1997),vacuole (Di Sansebastiano et al., 1998), mito-chondria (Köhler et al., 1997), plastids (Köhlerand Hanson, 2000), cell plate (Gu and Verma,1997), GA and ER (Boevink et al.,1998, 1999),and cytoskeleton (Köst et al., 1998; Marc et al.,1998).

In plant cells, GFP can be expressed on itsown, fused to targeting peptides, or as a chimerawith a complete protein of interest. Targetingthe protein into the ER can be achieved by theaddition of an appropriate N-terminal signalpeptide (such as sporamin, patatin, or chitinasesignal; Boevink et al., 1999; Haseloff et al.,1997) to translocate the protein into the ERlumen, and a C-terminal His/Lys, Asp, Glu, Leu(H/KDEL) retrieval sequence to maintain the

protein in the ER (see APPENDIX 1C). For GAvisualization, targeting (signal-anchor) se-quences from plant or mammalian transferasescan be spliced onto the C terminus of GFP tolocate the Golgi (Boevink et al., 1998), as cancomplete Golgi enzyme−coding sequences(Nebenführ et al., 1999). For instance the sig-nal-anchor sequence of a rat sialyl transferase,incorporating the transmembrane and cytoplas-mic amino acid domains, targets GFP to thetrans-Golgi, while a soybean mannosidase I-GFP fusion locates towards the cis-face (Ne-benführ et al., 1999). Both ER and Golgi canbe targeted with the Arabidopsis homolog ofthe yeast H/KDEL receptor, aERD2 (Boevinket al., 1998).

The transient expression protocol describedin this unit can be used for species other thanNicotiana. The authors have been able to tran-siently express ER-targeted GFP in Arabidop-sis, Petunia, and cucumber.

Critical ParametersThere are now a number of variants of GFP,

obtained by genetic modification of the wild-type gene, which have been optimized tochange various characteristics of the protein.This reengineering has been aimed at changingthe spectral properties of the protein andimproving the speed of maturation of the chro-mophore, including photoisomerisation andreduction in photobleaching of the protein(Cubitt et al., 1995, 1999); therefore, it is ad-visable to make a careful choice of the GFP tobe used before embarking on extensive trans-formation procedures. For instance, mutationswith enhanced blue excitation peaks may havereduced UV excitation, making rapid screeningof plants with UV lamps impossible.

In order to avoid possible artifacts, it isimportant to ensure that the detected GFP fluo-rescence in transiently expressing plant cells isnot derived from Agrobacteria synthesizing theprotein. With other reporter systems this prob-lem has been overcome by insertion of a plantintron into the reporter gene. Unfortunatelysuch intron-containing GFPs have yet to be


13.3.6


with GFP

constructed. Furthermore, as high expressionlevels may result in mistargeting, it is advisablethat serial dilutions of bacteria be used to opti-mize expression levels.

Due to the availability of the spectral vari-ants of GFP, including yellow (YFP), cyan(CFP) and the newly released red fluorescentproteins (dsRed), co-expression of constructsis possible. Care must then be taken with thechoice of filter sets to prevent bleed through ofsignal when excitation and emission spectra areclose. This of course is not the case for specifictechniques, such as fluorescence resonance en-ergy transfer (FRET; UNIT 17.1), in which theoverlapping spectra are essential parameters(Gadella et al., 1999).

GFP will remain fluorescent after both para-formaldehyde and light glutaraldehyde fixa-tion, thus permitting the use of other fluorescentprobes, such as rhodamine conjugated phal-loidin (Boevink et al., 1998), or standard im-munofluorescence protocols, with red or far-red emitting fluorochromes. GFP will alsoretain antigenicity after fixation and prepara-tion for immunogold labeling using the pro-gressive-lowering-of-temperature techniqueand embedding in acrylic resins (Vanden-Bosch, 1991; Boevink et al., 1998; UNIT 4.7).Thus, it is possible to confirm the location ofGFP fusion proteins at the ultrastructural level.This is important when proteins are targeted tosome of the smaller organelles, such as mito-chondria, plastids, and Golgi. One may encoun-

ter difficulties in obtaining GFP antibodies thatare suitable for immunogold labeling of planttissue. Consequently, it may be advisable toincorporate one of the widely used epitope tagsinto the GFP fusion if at some stage in the futureimmunogold labeling is envisaged.

Finally, with the observation of living plantmaterial, specific problems, such as autofluo-rescence of chlorophyll and rapid cytoplasmicstreaming, or organelle movements, are oftenencountered; these problems can be overcomeby selective use of filter sets and optimizationof microscope settings.

TroubleshootingSometimes a GFP chimera does not fluo-

resce, which can be due to several reasons. Forexample, frame shifts when GFP-fusions areused. If the resulting cDNAs have been checkedand confirmed by sequencing, it is advisable tocheck that the binary plasmid carrying the con-struct is stable in Agrobacterium. Another pos-sible explanation for lack of fluorescence maybe that the GFP chimera is misfolded. Levelsof expression can be checked by conventionalimmunoblot assay of protein extract (UNIT 6.2).This is also useful when with a particular con-struct, one encounters a relatively low-fluores-cence level, which may result from proteininstability or reduced folding kinetics.

On occasion a mislocalization of the fusionprotein can be observed. If the quality of thetargeting signals is trusted, then the phenome-

Table 13.3.1 Optimization of Images of GFP Targeted to GA and ER at the ConfocalMicroscope

Problem Solution Result

Autofluorescence ofchlorophylls

Use a narrow-band emissionfilter set (e.g., 515–525 nm) incombination with optimumbrightness and contrast settings

Reduction of fluorescencefrom chloroplasts

Blurred image due to thedynamics of the ER and GA

Increase laser intensity and/orincrease pinhole aperture incombination with single-linescan or collect data from adefined region of interest (ROI)

Reduction in resolution andconfocality may beexperienced. A collection oftime-resolved data sets forpresentation as movies can beobtained

Low signal from GFP that isnot compensated by changeof brightness/contrastsettings

Increase the laser intensityand/or increase the pinholeaperture

Increase of signal from GFPand reduction of confocalityand resolution

Photobleaching of GFPfluorescence

Reduce the laser intensity, thenumber of image scans foraveraging, and the number ofsections per Z-series

Reduction of resolution andphotobleaching damage


13.3.7

Organelle Motility

non may be due to overexpression of the con-struct. Optimization of the bacterial concentra-tion (as measured by optical density) is thensuggested. Mislocalization can be also due toinstability of the targeting signal within thefusion protein.

During observation of leaf tissue, the auto-fluorescence of the chlorophylls can mask lowlevels of GFP fluorescence. The use of a nar-row-band emission filter set (e.g., 515 to 525nm) in combination with optimum brightnessand contrast settings will reduce the autofluo-

rescence from chloroplasts. Increasing the laserpower and/or altering brightness and contrastsettings can usually compensate for the overallreduction in signal that may result. Movementof labeled organelles in plant cells may resultin a blurred image. Modifying laser and/orpinhole settings in combination with single-line scanning can help limit this problem. Ifnecessary, the authors recommend collectingdata from a defined region of interest (ROI). Areduction in resolution and confocality willoccur, but quick collection of time-resolved

Figure 13.3.1 (A) Agrobacterium-mediated expression of a signal peptide-GFP-HDEL construct in leaf epidermalcells of Nicotiana clevelandii. The low magnification micrograph shows high levels of expression in the endoplasmicreticulum at the cortex of the cells. (B) Potato virus X–mediated expression of a signal peptide-GFP-calreticulinconstruct in a leaf epidermal cell of N. clevelandii. (C) Virus-mediated expression of an ER/Golgi–targeted construct(aERD2-GFP) in a leaf epidermal cell. Note the bright Golgi bodies are associated with the cortical ER tubules. (D)Golgi dynamics. Sequential frame capture of Golgi (arrows) moving over static cortical ER tubules. All micrographstaken with a Zeiss LSM 410 laser scanning confocal microscope.


13.3.8


with GFP

data sets for presentation as movies can beobtained. GFP is an ideal marker for time-lapsestudies because of its relative resistance to pho-tobleaching; however, if a reduction of signalfrom GFP and its variants due to photobleach-ing is observed, the laser intensity, the numberof scans for image averaging, and the numberof sections per Z-series should be reduced.Recommendations for the optimization of theER and GA image at the confocal microscopeare summarized in Table 13.3.1.

Anticipated ResultsTypical results can be seen in Figure 13.3.1.

The results illustrated in the figure are typicalfor those organelles to which GFP has beendeliberately targeted. The percentage of trans-formed cells can vary with the time and withthe concentration of bacteria. With the experi-mental conditions suggested in the Basic Pro-tocol usually, 3 to 4 days after inoculation ofplants, 20% to 40% of the cells are transformedwithin the area of initial bacterial penetration.Similarly, the virus transformation may pro-duce approximately the same yield of trans-formed cells (after 4 to 5 days), and this numberof cells may increase with time due to the virusspread which is not a feature of Agrobacteriumtransformation.

Time ConsiderationAs mentioned above, fluorescence may be

detected 2 days after plant transformation. Inthe authors’ experience, this depends on theparticular construct, but in general, a detectableexpression is obtained from 2 to 4 days and maylast about 7 to 10 days.

Literature CitedBaulcombe, D.C., Chapman, S.N., and Santa Cruz,

S. 1995. Jellyfish green fluorescent protein as areporter for virus infection. Plant J. 7:1045-1053.

Bevan, M. 1984. Binary Agrobacterium vectors forplant transformation. Nucl. Acids Res. 12:8711-8721.

Blackman, L.M., Boevink, P., Santa Cruz, S.,Palukaitis, P., and Oparka, K.J. 1998. The move-ment of cucumber mosaic virus traffics into sieveelements in minor veins of Nicotiana cleve-landii. Plant Cell 10:525-537.

Boevink, P., Oparka, K., Santa Cruz, S., Martin, B.,Betteridge, A., and Hawes, C. 1998. Stacks ontracks: The plant Golgi apparatus traffics on anactin/ER network. Plant J. 15:441-447.

Boevink, P., Martin, B., Oparka, K., Santa Cruz, S.,and Hawes, C. 1999. Transport of virally ex-pressed green fluorescent protein through thesecretory pathway in tobacco leaves is inhibited

by cold shock and brefeldin A. Planta 208:392-400.

Chapman, S.N., Kavanagh, T., and Baulcombe, D.C.1992. Potato-Virus X as a vector for gene expres-sion in plants. Plant J. 2:549-557.

Cubitt, A.B., Heim, R., Adams, S.R., Boyd, A.E.,Gross, L.A., and Tsien, R.Y. 1995. Under-standing, improving and using green fluorescentproteins. Trends Biochem. Sci. 20:448-455.

Cubitt, A.B., Woolenweber, L.A., and Heim, R.1999. Understanding structure-function rela-tionships in the Aequoria victoria green fluores-cent protein. In Green Fluorescent Proteins (K.F.Sullivan and S.A. Kay eds.) pp. 19-30. AcademicPress, San Diego.

Di Sansebastiano, G.-P., Paris, N., Marc-Martin, S.,and Neuhaus, J.-M. 1998. Specific accumulationof GFP in a non acidic vacuolar compartment viaa C-terminal propeptide-mediated sorting path-way. Plant J. 15:449-457.

Gadella, T.W.J., van der Krogt, G.N.M., and Bissel-ing T. 1999. GFP-based FRET microscopy inliving plant cells. Trends Plant Sci. 4:287-291.

Grebenok, R.J., Pierson, E., Lambert, G.M., Gong,F.-C., Afonso, C.L., Haldeman-Cahill, R., Car-rington, J.C., and Galbraith, D.W. 1997. Green-fluorescent protein fusions for efficient charac-terisation of nuclear targeting. Plant J. 11:573-586.

Gu, X.J. and Verma, D.P.S. 1997. Dynamics ofphragmoplastin in living cells during cell plateformation and uncoupling of cell elongationfrom the plane of cell division. Plant Cell 9:157-169.

Haseloff, J., Siemering, K.R., Prasher, D.C., andHodge, S. 1997. Removal of a cryptic intron andsubcellular localisation of green fluorescent pro-tein are required to mark transgenic Arabidopsisplants brightly. Proc. Natl. Acad. Sci. U.S.A.94:2122-2127.

Hawes, C., Boevink, P., and Moore, I. 2000. Greenfluorescent protein in plants. In Protein Local-ization by Fluorescence Microscopy: A PracticalApproach (V.J. Allen, ed.) pp. 163-177. OxfordUniversity Press, Oxford.

Itaya, A., Hickman, H., Bao, Y.M., Nelson, R., andDing, B. 1997. Cell-to-cell trafficking of cucum-ber mosaic virus movement protein: Green fluo-rescent protein fusion produced by biolisticbombardment in tobacco. Plant J. 12:1223-1230.

Köhler, R.H. and Hanson, M.R. 2000. Plastid tu-bules of higher plants are tissue-specific anddevelopmentally regulated. J. Cell Sci. 113:81-89.

Köhler, R.H., Zipfel, W.R., Webb, W.W., and Han-son, M.R. 1997. The green fluorescent protein asa marker to visualise plant mitochondria in vivo.Plant J. 11:613-621.

Köst, B., Spielhofer, P., and Chua, N.-H. 1998. AGFP mouse talin fusion protein labels plant actinfilaments in vivo and visualises the actin cy-


13.3.9

Organelle Motility

toskeleton in growing pollen tubes. Plant J.16:393-401.

Marc, J., Granger, C.L., Brincat, J., Fisher, D.D.,Kao, T.-H., McCubbin, A.G., and Cyr, R.J. 1998.A GFP-MAP4 reporter gene for visualising cor-tical microtubule rearrangements in living epi-dermal cells. Plant Cell 10:1927-1939.

Nebenführ, A., Gallagher, L.A., Dunahay, T.G.,Frohlick, J.A., Mazurkiewicz, A.M., Meehl, J.B.,and Staehelin, L.A. 1999. Stop-and-go move-ments of plant Golgi stacks are mediated by theacto-myosin system. Plant Physiol. 121:1127-1141.

Oparka, K.J., Roberts, A.G., Prior, D.A.M., Baul-combe, D., and Santa Cruz, S. 1995. Imaging thegreen fluorescent protein in plants–Viruses carrythe torch. Protoplasma 189:133-141.

Rakousky, S., Kocabek, T., Vincenciova, R., andOndrej, M. 1998. Transient beta-glucuronidaseactivity after infiltration of Arabidopsis thalianaby Agrobacterium tumefaciens. Biologia Plan-tarum 40:33-41.

Rossi, L., Escudero, J., Hohn, B., and Tinland, B.,1993. Efficient and sensitive assay for T-DNA-dependent transient gene expression. Plant Mol.Biol. Reporter 11:220-229.

VandenBosch, K.A. 1991. Immunogold labelling.In Electron Microscopy of Plant Cells (J.L. Halland C. Hawes, eds.) pp. 181-218. AcademicPress, London.

Key ReferencesBoevink, P., Santa Cruz, S., Hawes, C., Harris, N.,

and Oparka, K.J. 1996. Virus-mediated deliveryof the green fluorescent protein to the endoplas-mic reticulum of plant cells. Plant J. 10:935-941.

This paper illustrates the production of GFP tar-geted to the ER of living plant cells using a PVX-based expression system.

Boevink et al., 1998. See above.

The authors describe the fusion of the wild-typeGFP to the transmembrane domain of a rat sialyltransferase in Nicotiana cells and its localization atthe GA. Moreover, they describe the splicing of thewild-type GFP to the C-terminus of the Arabidopsishomologue of the yeast HDEL receptor, aERD2 andthe localization of the protein chimera at the ER andGA, using the potato virus X expression system.

Haseloff et al., 1997. See above.

In this paper the authors describe how the wild-typeGFP was engineered to be expressed in plants in anon-virus mediated system and to be targeted to theER.

Nebenführ et al., 1999. See above.

The authors describe the fusion of a soybean Gm-Man1, encoding the resident Golgi protein α-1,2mannosidase-1, to the green fluorescent protein, andits targeting to the Golgi of Bright Yellow 2 suspen-sion-cultured cells.

Internet Resourceshttp://www.brookes.uk/schools/bms/research/

molcell/hawes/gfp/gfpold.html

Web site for movies obtained at the confocal lasermicroscope in Nicotiana clevelandii epidermal cellsexpressing GFP fused to the C-terminus of the trans-membrane domain of a rat sialyl transferase (local-ized at the GA) and the GFP spliced to theC-terminus of the Arabidopsis homologue of theyeast HDEL receptor, aERD2 (localized at the ERand GA).

http://www.plantsci.cam.ac.uk/Haseloff/Home.html

Web site for movies of GFP expressed in Arabidopsisroot tips showing different patterns of GFP expres-sion generated by enhancer detection.

http://www.mbg.cornell.edu/kohler/kohler_Trends.html

Web site with movies showing GFP targeting toplastids and mitochondria.

Contributed by Chris Hawes and Federica BrandizzilOxford Brookes UniversityOxford, United Kingdom

Henri Batoko and Ian MooreUniversity of OxfordOxford, United Kingdom


13.3.10


with GFP

UNIT 13.4Movement of Nuclei

This unit presents an in vitro assay for nuclear motility along microtubules (see BasicProtocol). This assay system mimics the movement of the female pronucleus of thefertilized frog egg. Dynamic microtubule asters are assembled from purified centrosomesin Xenopus cytosol. Purified nuclei are then added and the motility is observed usingvideo-enhanced differential interference contrast (VE-DIC) microscopy. This assay hasbeen used to demonstrate that cytoplasmic dynein drives the motility of nuclei alongmicrotubules in this system. Dynein activity causes nuclei to move to microtubuleminus-ends. Therefore, the nuclei move to and accumulate at centrosomes. Other com-ponents of the nuclear motility apparatus can be defined by this assay including regulatoryfactors present in the cytosol, or binding partners for dynein on the nuclear membrane.Pharmacological and biological inhibitors of nuclear motility can be assayed and theeffects on motility can be quantitated.

The assay system requires the following components: purified nuclei assembled inXenopus egg extracts (see Support Protocol 5); purified centrosomes (see SupportProtocols 1 and 2); and Xenopus cytosol that has been clarified by high-speed centrifu-gation (see Support Protocol 4) from fractionated interphase extracts (see Support Protocol3). The clarified extract provides the subunits for microtubule assembly, the dynein motorcomplex that drives motility, and regulatory factors that have not yet been defined.

STRATEGIC PLANNING

These experiments require familiarity with handling Xenopus laevis for obtaining eggsused in this protocol. Protocols for handling Xenopus for egg production and preparationof active cytoplasmic extracts are described elsewhere and should be reviewed (Murray,1991; UNIT 11.10). Included here are those procedures specifically required for nuclearmotility experiments. These experiments require significant microscopy expertise, espe-cially using VE-DIC. VE-DIC theory and techniques are not presented here. Refer to UNIT

4.1 for basic microscopy theory and techniques. Specific techniques for imaging micro-tubules and organelles using VE-DIC are presented elsewhere (Walker et al., 1988;Salmon and Tran, 1998; UNIT 13.1). The support protocols describe the procedures toprepare the different components used in the Basic Protocol.

BASICPROTOCOL

NUCLEAR MOTILITY ASSAY

This protocol describes the set-up and execution of an assay that combines dynamicmicrotubules with synthetic nuclei assembled in Xenopus egg extracts (see Fig. 13.4.1 fora schematic of the assay). All assay components are stored separately in small aliquots at−70°C and recombined during the assay. (Assay components have been stored as long as6 months without any loss of activity. Longer storage is conceivable, but has not beentested.) The use of aliquotted, frozen components allows reproducibility between assaysas well as the possibility to inactivate or modify individual components to study thefunction of individual proteins in microtubule-mediated nuclear motility.

Supplement 10

Contributed by Sigrid ReinschCurrent Protocols in Cell Biology (2001) 13.4.1-13.4.28Copyright © 2001 by John Wiley & Sons, Inc.

13.4.1

Organelle Motility

Materials

Centrosomes (see Support Protocol 1)PE buffer (see recipe)5 mg/ml casein in PE buffer (see recipe)ABC buffer (see recipe)HSS cytosol (see Support Protocol 4)DNA-bead nuclei (see Support Protocol 5)Valap (UNIT 13.1) at 37°CImmersion oilAssay reagent (e.g., drug, antibody, expressed protein)Antibodies bound to beads (for immunodepletion experiments)

tapecoverslipslide

add centrosomes

invert on ice 5 min

wash with casein/PEleave on ice until use

filter

add cytosol (HSS)incubate room temp10 min

add nuclei (HSS)

Figure 13.4.1 Flow chart of nuclear motility assay. Schematic showing the slide/coverslip chamber assembly used in thisassay, and the flow-through technique. The individual steps are shown on the left panels and a representative microscopefield is shown on the right panels. Centrosomes appear as round dots scattered on the field, microtubules are the linesemanating from the centrosomes in the second step, and nuclei moving on the asters are shown in the third step.


13.4.2

Movement ofNuclei

Inverted microscope equipped for high-resolution DIC opticsSimple perfusion chambers made from: clean slide, double-stick tape, and clean

18 × 18–mm coverslips (for preparation see UNIT 13.1, Support Protocol 1)Humid chamber: a covered 10- or 15-cm glass petri dish with strip of moist filter

paper around perimeterMagnetic particle concentrator (Dynal)Inverted microscopeCamera (e.g., Hamamatsu CCD C307, Hamamatsu)Image processor (e.g., Argus 10, Hamamatsu or equivalent)

Assay nuclear motility1. Prior to the experiment, optimize inverted microscope for DIC imaging of micro-

tubules.

2. Assemble perfusion chambers according to the instructions in UNIT 13.1, SupportProtocol 1. Assemble by placing two pieces of double-sided tape 2 mm apart on amicroscope slide and attach an 18 × 18–mm coverslip.

The chambers should have an approximate volume of 7 ��.

3. Thaw an aliquot of centrosomes. Dilute with PE buffer to the optimal concentrationdetermined in Support Protocol 2. Flow 7 µl of diluted centrosomes into eachchamber to be used in the day’s set of experiments. Be careful not to introduce bubblesinto the chamber. Invert perfusion chambers onto ice and allow the centrosomes tosettle onto the coverslip and attach for 5 min.

For all perfusion steps, samples are pipetted at one opening of the perfusion chamber andwicked on the opposite side with a small piece of filter paper.

4. Flow 30 µl of 5 mg/ml casein in PE through perfusion chamber to block binding siteson the glass and rinse out unattached centrosomes. Incubate on ice until use (up to 4hr maximum).

From this point, only one chamber will be used at a time because each must be viewedindividually on the microscope.

5. Ten minutes before the start of the assay, rinse chamber with 30 µl of ABC buffer.Flow 10 µl of HSS cytosol into chamber and incubate for 10 min at room temperaturein the humid chamber.

This allows recruitment of pericentriolar material from the cytosol, nucleation of micro-tubules, and aster formation.

6. During the 10-min incubation, quickly thaw an aliquot of DNA-bead nuclei (seeSupport Protocol 5) and immediately place on ice. Resuspend in 150 µl ABC buffer.Retrieve on a magnetic particle concentrator, carefully remove the ABC buffer, andresuspend in 10 ml HSS cytosol.

7. Flow the nuclear suspension into the chamber to start the assay. Seal the chamberwith Valap, oil top and bottom of chamber with immersion oil, and observe at 20° to22°C. Use DIC optics, a 100× objective, and a high-resolution condenser.

If appropriate, use additional magnification to obtain an optimal on-screen field size ∼50��. Use image background subtraction to enhance the microtubules (Walker et al., 1988;Salmon and Tran, 1998).

Simultaneous imaging of nuclei and microtubules is a bit trickier than imaging onlymicrotubules or other organelles due to the high refringence of the nuclei (especiallymagnetic-bead nuclei). Nuclei move rapidly on the microtubules. By 20 min, most nucleihave stopped moving and reached the centrosomes.


13.4.3

Organelle Motility

Collect images8. To film nuclear movements, collect shuttered images with a camera. Perform rolling

image averaging and background subtraction with an Argus 10 image processor, orequivalent image processing program.

An appropriate interval between shuttered images is 5 sec. Figure 13.4.2 shows severalsequential frames of a movie of a nucleus migrating on a microtubule aster.

Quantitate nuclear motility9. To perform a quantitative assay for accumulation at centrosomes, repeat steps 1

through 8. Starting at 20 min after adding DNA-bead nuclei, for an additional 10 min,count as many fields as possible to determine percentage of nuclei present atcentrosomes versus those at a distance (>5 µm) from centrosomes.

One should be able to count 50 to 100 nuclei in the 10-min interval. Typical values are90% to 95% of the nuclei present at centrosomes. Because many nuclei contain multipleDNA beads, each centrosome containing beads/nuclei should be counted as "1". Thereforenumbers for accumulation are minimal values. Values <90% to 95% indicate difficultieswith the assay that must be addressed before continuing (see Troubleshooting). If the assaylooks optimized, then proceed to the next step.

Perform addition experiments

To assess pharmacological or biological reagents for effect10a. Perform steps 1 through 4.

11a. Add 2 µl of drug, antibody, or expressed protein to 18 µl of HSS cytosol.

12a. Resuspend an aliquot of DNA-bead nuclei in 150 µl of ABC buffer. Retrieve on amagnet particle concentrator and discard supernatant.

13a. Resuspend DNA-bead nuclei in 10 µl of HSS/test compound mixture (step 11a).Use the remaining 10 µl to flow through perfusion chamber after the nucleation step(step 4) to replace the normal HSS cytosol with that containing the test compound.

14a. Flow in the nuclei/HSS/test compound mixture, incubate 20 min at 20° to 22°C andcount as in step 9.

Figure 13.4.2 DIC images of a nucleus moving on a microtubule aster. A sequence of four stills are shown from amotility assay over a period of 10 min. The centrosome is in the lower left corner of each panel, and a robust microtubuleaster emanates from this focus. A synthetic nucleus containing several paramagnetic beads starts in the upper rightcorner and moves to the centrosome during the course of the experiment. The nucleus appears refractory underAVEC-DIC imaging conditions. Image processing, including background subtraction and frame averaging, enhancesthe microtubules for clearer visualization. The time points are not indicated, but would occur during the first 20 minfollowing assembly of the assay. By the 20-min time-point where quantitation is performed, 90% of nuclei will havereached the centrosome.


13.4.4

Movement ofNuclei

Perform each assay three to four times to obtain statistically significant data. The controlmixture should contain 2 �l of appropriate buffer. Figure 13.4.3 shows a titration curve forthe effects of different concentrations of vanadate on the accumulation of nuclei atcentrosomes.

For some reagents, it is important to incubate the cytosol with the reagent for 30 to 60 minbefore the assay. This is especially true for proteins expressed in bacteria, which may needto undergo refolding, or other post-translational modification in the extract to be effective(S. Reinsch, unpub. observ.).

To perform depletion experiments10b. Immunodeplete specific components from HSS cytosol using antibodies bound to

beads. Use depleted HSS cytosol for motility cytosol in the assay.

For examples of successful depletion experiments using Xenopus extracts see Merdes et al.(1996); Walczak et al. (1996); or Tournebize et al. (2000).

These experiments are technically more challenging.

0

20

40

60

80

100

0 20 40 60 80 100Vanadate (µM)

Per

cent

nuc

lei a

t cen

tros

omes

Figure 13.4.3 Effect of vanadate on accumulation of nuclei at centrosomes is an example of howthis assay is used to derive quantitative data. Accumulation of nuclei at centrosomes was comparedin HSS containing varying amounts of sodium orthovanadate, a general ATPase inhibitor. Assayswere performed in triplicate for each vanadate concentration to generate a dose-response curve.The experiment indicates that vanadate significantly inhibits nuclear accumulation at centrosomesat a concentration of 20 mM. This concentration has been shown to be effective in inhibiting themotility of cytoplasmic dynein, while inhibition of kinesins requires concentrations of vanadate inthe 100 mM range.


13.4.5

Organelle Motility

SUPPORTPROTOCOL 1

CENTROSOME ISOLATION FROM LYMPHOCYTES

Centrosomes in fibroblastic and lymphoid cells are anchored in the cytoplasm bymicrotubules and interact with the nucleus in an unknown way. The centrosome-nucleusassociation is dynamic in vivo but becomes irreversible in vitro if cells are lysed atphysiological ionic strength. Most procedures for making nuclei lead to the preparationof nucleus-centrosome complexes that are almost impossible to dissociate. Therefore, inthis protocol, the nucleus-centrosome interaction in vivo is first disrupted, then cells arelysed under conditions where centrosomes can be purified away from the nucleus.

Human lymphoid cells in culture are used here because they grow in suspension and havea low cytoplasm/nucleus ratio. Evidence indicates that both the microtubule cytoskeletonand the actin network are involved in centrosome anchoring in the cytoplasm (Euteneuerand Schliwa, 1985; Buendia et al., 1990; Mack and Rattner, 1993). To purify centrosomes,cells are first preincubated with cytochalasin and nocodazole to disrupt connectionsbetween centrosomes and nucleus (Bornens et al., 1987). Extremely fast lysis of cells atvery low ionic strength in the presence of Triton X-100 leads to chromatin dispersion andrelease of centrosomes in suspension. The addition of a low concentration of magnesiumions to the lysis buffer leads to partial chromatin stabilization and still allows centrosomerelease. This reduces contamination of centrosomes by chromatin components. Centro-somes are rapidly sedimenting particles. Their purification is therefore achieved byseparation on sucrose gradients.

It takes ∼1 week to grow enough cells for the preparation. The preparation itself takes 1day. One or two people can do the preparation, and a minimum of two people are neededto perform steps 6 to 7, which need to be done very rapidly. It is important to have highlyconcentrated centrosomes (∼2 × 108/ml) for this assay. The yield is typically 30% to 50%,with several fractions containing centrosomes at the optimal concentration.

Materials

KE37 human lymphoblastic cells (ACC46; DSMZ German Collection ofMicroorganisms and Cell Cultures)

RPMI-10: RPMI 1640 medium with 10% (w/v) FBS10 mM nocodozole stock solution in DMSO (see recipe)10 mg/ml cytochalasin D stock solution in DMSO (see recipe)PBS (see recipe), ice coldPBS/10 with 8% sucrose (see recipe)Lysis buffer (see recipe)4 ml 0.5 M K-PIPES (pH 7.2)/1 mM EDTA (see recipe)1 mg/ml DNase I (see recipe)40%, 50%, and 70% sucrose in gradient buffer (see recipe)Liquid nitrogenPE buffer (see recipe)Methanol, −20°CPBS/0.1% Triton X-100Monoclonal anti-tubulin antibody (Amersham)Secondary antibodiesHoechstPolyclonal anti-pericentriolar antibody (e.g., γ-tubulin, pericentrin; optional)

250-ml plastic flask2-liter spinner culture flask500-ml centrifuge bottlesClinical centrifuge or equivalent with swinging bucket rotor for 50-ml tubes


13.4.6

Movement ofNuclei

Super-speed centrifuge (e.g., Sorvall RC-26 or equivalent)Large volume rotor (GSA or equivalent)50-ml capped conical tubes10-ml plastic pipet125-µm nylon mesh (Millipore)Beckman SW28 and SW28.1 centrifuge tubesUltracentrifugeUltracentrifuge SW28 rotor with SW28 and SW28.1 buckets18-G needlesRefractometerModified Corex tubes (Evans et al., 1985; UNIT 11.13)11- to 12-mm diameter coverslips (acid-washed)HB-4 or HB-6 rotorForcepsFixing jar with coverslip holder

Additional reagents and equipment for immunofluorescence staining (UNIT 4.3) andanalyzing centrosomes by spinning onto coverslips (UNIT 11.13)

Grow KE37 cells1. Grow 1 liter of KE37 cells in four 250-ml plastic flasks in RPMI 1640-10.

2. One day before the centrosome preparation, transfer 1-liter cells to a 2-liter spinnerculture bottle and add 1 liter RPMI 1640-10.

3. On the day of purification have 2 liters of cells at a density close to 1.5 × 106 cells/ml(3 × 109 cells total).

It is important that the cells are maintained in exponential phase at densities between 1and 4 × 106 cells/ml.

4. Add 60 µl of 10 mM nocodazole to a final concentration of 33 µM and 200 µl of 10mg/ml cytochalasin D to 1 µg/ml final. Incubate 1 hr at 37°C with continuous stirring.

5. Transfer to 500-ml centrifuge bottles and centrifuge for 15 min at 650 × g (2000 rpmin GSA rotor), 4°C.

During all subsequent centrifugations, do not decant supernatant, but rather aspiratesupernatant.

6. Resuspend cells in ice-cold PBS to a maximum total volume of 160 ml.

7. Transfer to four 50-ml capped conical tubes. Centrifuge for 5 min at 500 × g (in aclinical centrifuge), 4°C. Repeat PBS wash and centrifugation once.

8. Resuspend cells in ≥25 ml/tube PBS/10 with 8% sucrose. Centrifuge for 5 min at 250× g (1000 rpm in a clinical centrifuge), 4°C.

Do this step and step 9 as quickly and gently as possible.

9. Lyse cells in each tube by adding 10 ml of lysis buffer to pellet. Disperse pellet bypipetting up and down with a 10-ml plastic pipet without generating bubbles. Bringvolume in each tube up to 20 ml with lysis buffer. Invert 2 to 3 times slowly. Incubate5 min on ice.

Cells should not form an aggregate. They should lyse immediately. The maximum totalvolume for the preparation should not exceed 90 ml at this point.

10. Centrifuge 10 min at 2000 × g (3000 rpm in a clinical centrifuge), 4°C.


13.4.7

Organelle Motility

11. Filter supernatant through 125-µm nylon mesh. Remove a 300-µl sample for count-ing.

12. To the pooled 90 ml supernatants add 1.8 ml of 0.5 M K-PIPES (pH 7.2)/1 mM EDTAand 90 µl DNase I stock solution.

Purify centrosomes on sucrose gradients13. Load 30 ml of lysate into three Beckman SW28 ultracentrifuge tubes and using a

10-ml plastic pipet underlay with 5 ml of 50% sucrose in gradient buffer.

14. Ultracentrifuge 20 min at 20,000 × g (11,100 rpm in SW28 rotor), 4°C.

15. Aspirate supernatant leaving ∼2 ml of lysate above the cushion in each tube.

To estimate the 2-ml volume above the cushion, prepare a blank tube containing the cushionvolume and 2 ml above this volume. Mark tube at the appropriate height to compare withthe tubes containing the centrosome samples.

16. Take the 2 ml of remaining supernatant and 2 ml of 50% sucrose cushion. Pool 12-mltotal volume from the three step gradients. Keep a 100-µl aliquot for counting.

17. Mix well and layer over a discontinuous sucrose gradient in a SW28.1 ultracentrifugetube.

2.0 ml 70% sucrose1.5 ml 50% sucrose1.5 ml 40% sucrose12.0 ml of centriole suspension (∼25% sucrose: pooled volume from thethree-step gradients).

18. Ultracentrifuge 75 min at 110,000 × g (25,000 rpm in SW28.1 rotor), 4°C.

19. Eliminate top of the gradient to the 40% solution. Pierce bottom of tube with an 18-Gneedle and collect 0.4-ml fractions manually.

Analyze gradient fractions and count centrosomes20. Read the fractions with a refractometer. Take 5-µl aliquots from fractions between

40% and 70% sucrose to count centrosomes by immunofluorescence. Snap freezethe remaining volume of these fractions in liquid nitrogen without further aliquotting.Store at −70°C. Discard fractions at <40% and >70% sucrose.

21. Prepare one modified Corex tube for each 5-µl aliquot and one each for the samplesfrom steps 11 and 16. Place the removable chuck on top of the bottom spacer. Placea clean 11-mm diameter coverslip on top of the chuck.

22. Mix the 5-µl aliquots each with 5 ml of PE buffer by inverting several times in a 15-mlcapped tube. Pipet into the prepared Corex tube. Centrifuge 10 min at 23,600 × g(12,000 rpm in HB4 rotor), 4°C.

23. Carefully remove coverslips from the tube: use a thin spatula with a bent tip to liftthe chuck out of the tube, then use forceps to transfer coverslip into cold methanol.Fix coverslips 5 min in methanol at −20°C.

It is important to remember which side of the coverslip the centrosomes are on; maintainall of the coverslips in the same orientation.

24. Remove coverslips from fix and place in PBS/ 0.1% Triton X-100.

For a more detailed description of the proecdure in steps 21 through 25, see UNIT 11.13,Support Protcol 4.


13.4.8

Movement ofNuclei

25. Perform immunofluorescence using a monoclonal anti-tubulin antibody, followed byappropriate secondary antibodies (UNIT 4.3). Add Hoechst to secondary antibodies tocheck for DNA contamination.

If possible, also use a polyclonal antibody against pericentriolar material (anti-γ-tubulin).This helps to confirm that the dots are centrosomes. Santa Cruz Biotechnology offerspolyclonal anti-γ-tubulin antibody, but it has not been tested in this protocol.

26. Observe samples at a magnification where centrosomes are easily visualized (usuallyas pairs of dots, with some single dots). Count the total number of centrosomes inseveral microscope fields to get an average number. Use a stage micrometer tomeasure the size of the field. Extrapolate to the total cross-sectional area of the Corextube to get the total number of centrosomes in a particular 5-µl aliquot or fromintermediate steps 11 and 16 (see Fig. 13.4.4 for a typical determination of centro-some concentration from a sucrose gradient).

27. Maintain the sucrose gradient fractions of centrosomes frozen at −80°C.

Centrosomes can be stored this way for ≥1 year. See Support Protocol 2 for subaliquottingthe centrosomes and for titration in the motility assay.

SUPPORTPROTOCOL 2

TITRATION OF CONCENTRATED CENTROSOMES

The assay presented in the Basic Protocol requires standardized components so that assaysperformed on different days can be compared. In particular, the same concentration ofcentrosomes must be used from one assay to the next as the plating density of centrosomesaffects assay results.

4

3

2

1

70

60

50

402

04 6 8 10

Per

cent

suc

rose

(do

tted

line)

Fraction number

No.

of c

entr

osom

es (

× 10

8; f

illed

circ

les)

Figure 13.4.4 Fractionation of centrosomes in sucrose step gradient. A typical graph obtainedafter fractionation of centrosomes on a sucrose step gradient. The sucrose concentration (dottedline) of each fraction was determined using a refractometer, while the centrosome concentration(solid line) was determined as in Support Protocol 1. The peak fractions, containing centrosomessufficiently concentrated for use in this assay, typically sediment between 45% and 50% sucrose.


13.4.9

Organelle Motility

Additional Materials (also see Basic Protocol)

Large aliquot of centrosomes (see Support Protocol 1)Liquid nitrogen0.5-ml microcentrifuge tubes for aliquotting

Prepare centrosome aliquots1. Prepare about one hundred 0.5-ml microcentrifuge tubes on ice.

2. Rapidly thaw one large 400- to 500-µl aliquot of centrosomes. Mix the aliquot wellby inverting tube several times or pipetting up and down without introducing bubbles.Make 5-µl aliquots. Snap freeze in liquid nitrogen.

It is important that the large aliquot is well mixed before dividing into smaller aliquots sothat the concentration of centrosomes is the same in each tube. Since centrosomes are largeparticles they will sediment in the tube over time.

Titrate centrosomes3. Prepare 8 perfusion chambers according to UNIT 13.1, Support Protocol 1.

4. Thaw four 5-µl aliquots of centrosomes. Resuspend each in a different volume tobracket the range of final concentrations of 5-20 × 106 centrosomes/ml.

5. Flow each centrosome sample into two perfusion chambers to serve as duplicates.

6. Perform Basic Protocol, steps 2 through 7 on each sample to determine whichcentrosome concentration yields the highest value for number of nuclei that accumu-late at the centrosomes.

100

5 10 15 20

Centrosomes/ml (× 106)

90

80

70

60

50

40

Per

cent

nuc

lei a

t cen

tros

omes

Figure 13.4.5 Titration of centrosomes in motility assay. An assay showing how centrosomeconcentration affects the accumulation of nuclei at centrosomes. If centrosomes are plated tosparsely, microtubules form spontaneously rather than at centrosomes preferentially, and nuclei willmove in a random fashion on these microtubules rather than accumulating at centrosomes. At highcentrosome concentrations, elevated concentrations beyond a certain value do not enhanceaccumulation of nuclei at centrosomes. Steps for titrating centrosomes for this assay are describedfor Support Protocol 2.


13.4.10

Movement ofNuclei

A typical plot is shown in Figure 13.4.5.

Empirically, a final concentration in the range of 7.5 × 106 centrosomes/ml gives consis-tently good values. This corresponds to a plating density of 2 to 3 centrosomes/50 �m2

microscope field. The optimum value is a result of the competition between free nucleationof microtubules and that nucleated by centrosomes. If there are too few centrosomes, thereis a lot of spontaneous nucleation of microtubules that generates random arrays. With toomany centrosomes, there is little or no spontaneous microtubule assembly, but microtubuleswill have a shorter overall length at the 20-min time point.

SUPPORTPROTOCOL 3

PREPARING FRACTIONATED INTERPHASE EXTRACTS FOR NUCLEARASSEMBLYFunctional nuclei can be assembled in vitro using Xenopus extracts (for review seeNewmeyer and Wilson, 1991; Gant and Wilson, 1997; UNIT 11.10). Freshly made 16,000 ×g interphase extracts generally give the best results for nuclear assembly and can be usedwith the DNA-magnetic beads (stop at step 11). However, extracts that have beenfractionated into membraneous and cytosolic components are useful for many types offunctional studies and are therefore presented here. The cytosolic and membrane fractionsare stored frozen and are recombined with template DNA to assemble chromatin andnuclei. Many published protocols for nuclear assembly use sperm nuclei. These are notappropriate for studies of nuclear motility that mimic movement of the female pronucleusalong microtubules. Sperm nuclei have an associated centrosome, and microtubulesnucleated from this centrosome will generate motility events in their own right (reviewedin Reinsch and Gonczy, 1998). For motility assays one can use nuclei assembled withmagnetic beads coupled with plasmid DNA, or nuclei assembled around larger purifiedDNA such as lambda DNA. The use of magnetic beads as substrates for nuclear assemblyallows rapid and gentle purification of the nuclei.

For handling of frogs and general methodologies for preparing Xenopus egg extracts seeMurray (1991) and Newmeyer and Wilson (1991); also see UNIT 11.10. The followingprotocol uses eggs that have been collected after overnight stimulation of the frogs withhuman chorionic gonadotropin (HCG), and is based on a published protocol (Hartl et al.,1994; also see UNIT 11.10).

The general principles of this procedure are illustrated in Figure 13.4.6.

NOTE: All protocols using live animals must first be reviewed and approved by anInstitutional Animal Care and Use Committee (IACUC) or must conform to governmentalregualtions regarding the care and use of laboratory animals.

Materials

5 to 7 frogs100 U pregnant mare serum gonadotropin (PMSG; see recipe)500 U HCG (see recipe)MMR (see recipe)Dejellying solution (see recipe)S-lysis buffer (see recipe)S-lysis-plus buffer (see recipe)10 mg/ml cytochalasin D stock solution in DMSO (see recipe)1 M DTT (see recipe)Protease inhibitors (LPC; see recipe)2.5 M sucrose (see recipe)GlycerolLiquid nitrogen for freezing aliquotsS-lysis-plus/500 mM sucrose solution (see recipe)


13.4.11

Organelle Motility

1-ml syringes and 27-G needles400-ml beakersSW50 ultraclear centrifuge tubes (Beckman)Pasteur pipet cut to a wide-mouth bore with a file, fire polishedSarstedt 13-ml adaptor tubes2-ml syringe and 18-G needleClinical centrifugeSorvall RC5 centrifugeHB-4 rotor with rubber adapters (Sorvall)5-ml polyproponate tubes with capsUltracentrifugeSW55 rotor500-ml microcentrifuge tubes for aliquotting cytosol and membranes

cytosol-200

membranes (M-200)mitochondria

glycogen/ribosomes

dilute 1:1in ABC buffer100,000 × g

nuclei

cytosol-200M-200glycogen

cytosol-200glycogen

chromatin

Dynabeads coupled withbiotinylated DNA

10,000 × g 200,000 × g

Figure 13.4.6 Nuclear assembly from fractionated extracts. Schematic showing the basic stepsrequired to assemble synthetic nuclei using fractionated extracts. In the protocol presented here,generation of the cytosol-200 and M200 fractions are described in Support Protocol 3. These arefrozen separately and then recombined with DNA-coupled Dynabeads as described in SupportProtocol 5. Preparation of DNA-coupled Dynabeads is described in UNIT 11.13 (Heald et al., 1998).If the M200 membranes are omitted during the assembly process, interphase chromatin assembleson the DNA, but full nuclear assembly requires the M200 fraction. Synthetic nuclei often containmultiple Dynabeads which behave very similarly in the assays presented here.


13.4.12

Movement ofNuclei

NOTE: Make sure that all glassware is clean and rinsed with distilled water before use.Additionally, wet all glassware with buffer before contact with the eggs as they will stickto the glassware and activate or lyse.

Collect the eggs1. Inject 5 to 7 frogs subcutaneously into the dorsal lymph sac with 0.5 ml of 100 U

PMSG, using a 1-ml syringe and 27-G needle. Perform the injection ≥4 days (and≤10 days) before the extract is to be prepared. Do not feed frogs after PMSG injection.

2. Twelve to 18 hr before use, inject each frog subcutaneously with 0.5 ml of 500 UHCG. Place frogs in individual containers with 500 ml MMR and maintain the frogsovernight at 16°C.

3. Collect eggs into 400-ml beakers using a separate beaker for each frog. Discardbatches of eggs containing >5% of lysed, mottled, or stringy eggs.

Keep batches of eggs separate through dejellying and even through the 16,000 × gcentrifugation, if possible. Note that there is no need to activate the eggs to send them intointerphase. They will spontaneously enter interphase due to lysis in the presence of Ca2+

ions. Cycloheximide added to the buffers will maintain the interphasic state of the extract.Any batches of eggs that lyse during dejellying, loading into tubes, or packing in thelow-speed centrifuge should be discarded.

Dejelly the eggs4. Pour off MMR and add 50 to 100 ml dejellying solution to each beaker. Swirl beaker

gently and change the solution several times until the eggs start to pack (usually ∼5min).

The eggs should be left in dejellying solution for the minimal amount of time required tocompletely remove the jelly coat. Further incubation seriously compromises egg quality.

5. Rinse three times with 50 to 100 ml MMR. Work quickly through the next steps, untillysis/centrifugation, as the eggs are quite fragile once the jelly coat is removed.Ensure that the eggs are always covered with buffer. Remove bad eggs (white andpuffy, or dark pigment retracted or mottled).

Lyse the eggs6. Pour off as much MMR as possible while keeping eggs immersed in a minimal

quantity. Rinse two times with S-lysis buffer in the same manner.

7. Rinse once with S-lysis-plus buffer. Use a wide-bore Pasteur piper to load eggs intoSW50 ultraclear centrifuge tubes containing 1 ml S-lysis-plus buffer and 10 µl of 10mg/ml cytochalasin D stock (to 100 µg/ml final concentration). Fill tubes as full aspossible and remove any buffer on top of eggs.

Always place the tip of the pipet containing the eggs into the buffer in the tube beforeexpelling the eggs. This ensures that the eggs do not contact air which would cause lysis.

8. Place each filled SW50 tube into a Sarstedt 13-ml adapter tube containing ∼0.5 mlwater in the bottom. Pack eggs by centrifuging 30 sec at low speed, 150 × g, roomtemperature, in a clinical centrifuge, then 30 sec at 700 × g. Remove all the bufferabove the eggs and discard any tubes with significant lysis.

9. Lyse eggs by centrifuging 15 min at ∼16,000 × g (10,000 rpm in HB-4 rotor containingrubber adapters), 4°C. Remove tubes from adapters and place upright on ice.


13.4.13

Organelle Motility

Collect cytoplasm10. Attach an 18-G needle to a 2-ml syringe. Move syringe plunger back and forth once

to loosen before piercing tube. Wipe sides of tube with a tissue and then pierce thetube just above the bottom of the clear cytoplasmic layer. Make sure the opening ofthe needle faces upwards. Withdraw the clear straw-colored cytoplasm avoiding theyellow lipid layer above and the heavier layers containing yolk, mitochondria, andcortex fragments below. Remove the needle from the syringe and expel the cytosolfrom the syringe into an appropriate sized polyproponate tube with a cap.

Bracing the centrifuge tube against the inside of an ice bucket will help avoid accidentsduring piercing.

One 5-ml tube of eggs will yield 1 to 2 ml of cytosol.

11. Pool cytosols that look good. Add 1/1000 volume each of cytochalasin D (final 10µg/ml), DTT (to 1 mM) and protease inhibitors.

12. If there is any contamination of the cytosol, then repeat the 16,000 × g centrifugationto clarify cytosol of any residual yolk or heavy cortex fragments. Remove clearcytosol as above and add 2.5 M sucrose to 125 mM to the cytosol.

This helps separate the membranes out of the cytosol in the following centrifugation.

Isolate membrane and cytosol fractions13. Ultracentrifuge cytosol for 60 min at 200,000 × g (555,000 rpm in SW55 rotor), 4°C.

There will be the following fractions from top to bottom: lipid, clear cytosol, goldenmembranes, grayish mitochondrial membranes, then translucent pellet consisting of gly-cogen and ribosomes.

14. Carefully aspirate clear lipid layer and discard. Remove the clear cytosol to a separatetube using a 200-µl tip (do not use a 1000-µl tip).

From a 5-ml tube one should collect ∼2.5 ml. It is difficult to remove the clear cytosolwithout pulling up some of the golden membranes. Go slowly when near the membranelayer.

15. Remove and save golden membrane fractions (they are the nuclear assembly mem-branes) and follow with them at step 17. Carefully avoid the gray mitochondrial layer.

16. Recentrifuge cytosol 30 min at 200,000 × g (55,000 rpm in TLS55 rotor), 4°C, toremove residual membranes and lipid. Remove cytosol as in step 14. To cytosol, addglycerol to a final concentration of 3%. Snap freeze in liquid nitrogen in 50-µlaliquots. Store aliquots at −70°C.

This cytosol fraction is called cytosol-200.

17. Resuspend the golden membranes in excess S-lysis-plus (≥10 vol) and centrifugeover a 0.5-ml cushion of S-lysis-plus/500 mM sucrose for 20 min at 26,000 × g(14,000 rpm in SW55 rotor), 4°C. Resuspend as a 10× stock in S-lysis-plus/500mMsucrose solution and freeze in 10-µl aliquots (10× stock means 1/10th the volume ofcytosol).

This membrane stock is called M-200.


13.4.14

Movement ofNuclei

SUPPORTPROTOCOL 4

PREPARATION OF HIGH-SPEED SUPERNATANT (HSS)The cytosol used for the motility assay is clarified of all membranes by dilution ofconcentrated 16,000 × g cytosol followed by centrifugation. The concentrated cytosol isprepared, dispensed into aliquots, and stored frozen until the day of the experiment, whenthe dilution and centrifugation steps are performed.

Additional Materials (see Support Protocol 3)

ABC buffer (see recipe)Table top ultracentrifuge and TLA-100 rotor or Airfuge (Beckman) and rotor

1. Perform Support Protocol 3, steps 1 through 11. However, in step 11 lower the sucroseconcentration to 100 mM rather than 125 mM.

2. Pipet the cytosol into 50-µl aliquots and snap freeze in liquid nitrogen. Store aliquotsat −80°C until use.

3. On the day of the assay, thaw an aliquot of cytosol and dilute with 2 vol ABC buffer.Centrifuge for 20 min at 100,000 × g (15 min, 30 psi in Beckman Airfuge or 550,000rpm in TLA-100 rotor, ), 4°C, to pellet membranes.

4. Pipet the cytosol to a fresh tube and maintain on ice until use (∼4 hr max).

SUPPORTPROTOCOL 5

NUCLEAR ASSEMBLY USING DNA-COATED MAGNETIC BEADS ASTEMPLATEThis protocol describes the assembly of nuclei around DNA-coated magnetic beads. Theprotocol for binding of DNA to the beads is presented in UNIT 11.13 (Heald et al., 1998).The nuclei can be assembled by simply adding DNA beads to fresh 16,000 × g cytosol(see Support Protocol 2, step 11). However, to perform manipulations of the nuclei to testspecific components of the nucleus for a role in nuclear motility, use this alternateapproach. Each batch of fractionated cytosol has to be titrated for nuclear assembly usingDNA-magnetic beads. Given below is a starting protocol. One may have to vary theglycogen, amount of membranes, and/or dilution of cytosol with ABC buffer to get betterassembly. Assembly using beads is much trickier than assembly with either sperm nucleior lambda DNA. The beads must have a high concentration of DNA. Only one end of theDNA should be attached to the beads (i.e., fill in only one end of the DNA with biotinylatednucleotides), and the plasmid cannot be too short. (The MCP plasmid is ∼8 kb andassembles nuclei much better than a 6-kb plasmid. Much shorter plasmids do not assembleat all.) Once the parameters for assembly with the extract preparation has been optimized,then scale up to make a nuclear prep for dispensing aliquots and freezing.

Materials

Cytosol-200 (see Support Protocol 2)ABC buffer (see recipe)DNA-Dynabeads (for preparation see UNIT 11.13, Heald et al., 1998)PBS/1% (w/v) BSA (see recipe)150 mg/ml glycogen stock (see recipe)M-200 (see Support Protocol 2)50 mM Mg-ATP (see recipe)0.5 M creatine phosphate (see recipe)8 mg/ml creatine kinase (see recipe)1.4 mg/ml TRITC-BSA-NLS (transport substrate; UNIT 11.7, Support Protocol 2)Fix solution (see recipe)

Beckman table top ultracentrifuge and TL100 rotor (or Beckman airfuge and rotor)Magnetic particle concentrator


13.4.15

Organelle Motility

Microscope slides and clean coverslipsNail polishMicroscope equipped with epifluorescence optics

1. Thaw one 50-µl aliquot of cytosol-200.

2. Add 50 µl ABC buffer and centrifuge for 30 min at 100,000 × g (55,000 rpm inTL100rotor or 15 min, 30 psi in airfuge), 4°C.

3. During spin, prepare DNA beads. Use 10 µl beads (70 µg/ml) for a 100-µl reaction.Resuspend the DNA-bead stock well before pipetting. Pipet 10 µl beads to a cleantube. Add 200 µl PBS/1% BSA and pipet beads up and down several times. Retrievebeads on the magnetic particle concentrator, carefully remove the buffer and repeatwashing step.

4. Resuspend washed beads in the diluted and centrifuged ceptosol (step 2, ∼100 µl).Pipet thoroughly but carefully to disrupt any aggregates. Do not introduce air bubblesinto the cytosol.

Failure to disrupt aggregates results in large masses of beads that do not properly assembleinto nuclei.

5. Add and mix by pipetting:

10 µl 150 mg/ml glycogen10 µl membranes (M-200)2 µl 50 mM Mg-ATP2 µl 0.5 M creatine phosphate1 µl 8 mg/ml creatine kinase.

Incubate 2 to 3 hr at 20°C.

Fully formed nuclei are first apparent at 40 min and increase progressively until 3 hr.

6. To assay for functional nuclei, remove a 10-µl aliquot and add 0.5 µl of 1.4 mg/mlTRITC-BSA-NLS. Incubate 30 min at 4°C. Pipet 2 µl of this reaction onto a cleanslide. Add 5 µl of fix solution and apply a clean coverslip. Seal the edges with nailpolish if retention of the sample is desired. Observe nuclei with a microscopeequipped with epifluorescence optics to detect nuclei that have accumulated thetransport substrate (see Fig. 13.4.7 for images of nuclear uptake).

Since Dynabeads are autofluorescent with emissions in the red wavelengths, a nuclearuptake substrate that will emit in the green wavelengths is often preferable, such as GFPor fluorescein-labeled. If desired, prepare FITC-BSA-NLS as in UNIT 11.7, Support Protocol2, by substituting fluorescein-5-EX succinimidyl ester (Molecular Probes) for TRITC insteps 2 to 6 of the protocol. Follow manufacturer’s recommendations for coupling bufferand pH conditions. The rest of the protocol remains the same. Alternatively, prepare GFP-GST-NLS (UNIT 11.7, Support Protocol 3). Use both as indicated here for TRITC-BSA-NLS.

7. When functional nuclei are observed, then scale up the reaction 10- to 20-fold andrepeat all steps including the assay step. To freeze, add glycerol to 10% (v/v), dispense10-µl aliquots, and freeze in liquid nitrogen. Store at −70°C.

Nuclei can be stored for several years in this state. Frozen nuclei can be thawed on ice andwashed in ABC buffer, retrieved on a magnet for several minutes (on ice) and resuspendedin cytosol, or other buffers. Typically, they retain ∼90% activity or higher after freezing.One 10-�l aliquot of nuclei is used for each motility assay.

If the membranes are omitted from the reaction, interphase chromatin assembles onto theDNA, but no nuclear envelope forms. The motility of the chromatin can be compared withthat of bona fide nuclei (Reinsch and Karsenti, 1997).


13.4.16

Movement ofNuclei

REAGENTS AND SOLUTIONS

Use deionized or distilled water in all recipes and protocol steps. For common stock solutions, seeAPPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.

ABC (complete acetate buffer)Acetate buffer (see recipe) supplemented with:7.5 mM creatine phosphate (see recipe for 0.5 M)1 mM MgATP80 µg/ml creatine kinase (see recipe for 8 mg/ml)10 mg/ml protease inhibitorsStore as 1- to 2-ml aliquots 1 year at −20°CThaw once and discard

Acetate buffer

To prepare 1 liter:9.8 g potassium-acetate (100 mM final)0.64 g magnesium-acetate (3 mM final)1.9 g EGTA (5 mM final)2.38 g HEPES (10 mM final, pH 7.4 )51.34 g sucrose (150 mM final)H2O to 950 mlAdjust the pH to 7.4H2O to 1 literAliquot in 50-ml tubesStore 1 year at −20°C

Figure 13.4.7 Nuclei assembled around magnetic DNA-beads are functional for nuclear transportand contain nuclear antigens. Confocal fluorescence images of synthetic nuclei assembled usingthe protocols described here. Autofluorescence of the beads is shown below (rhodamine filter),while the upper panels show the same beads in the fluorescein channel. FITC-BSA-NLS: typicalfluorescence following uptake of the nuclear transport substrate FITC-BSA NLS in an unfixedsample. Staining for lamins and nuclear pores gives the typical peripheral staining pattern for theseantigens.


13.4.17

Organelle Motility

Casein in PE100 mg caseinAdd 20 ml PE (see recipe)Vortex vigorouslyIncubate at 37°C with intermittent vortexing to aid solubilization of caseinAliquot and store 1 year at −20°C

Creatine kinase, 8 mg/ml8 mg creatine kinaseAdd 475 µl distilled waterAdd 20 µl 1 M HEPES-KOH, pH 7.5Add 5 µl 1 M DTTAdd 500 µl glycerolSnap freeze in 10-µl aliquots in liquid nitrogenStore 1 year at −70°C

Creatine phosphate, 0.5 M127.55 mg creatine phosphate1 ml H2OStore 1 year at −20°C in 50-µl aliquots

This solution can be thawed and refrozen

Cytochalasin D, 10 mg/ml10 mg of cytochalasin1 ml DMSOStore 1 year at −20°C in 50-µl aliquots

Dejellying solution (2% L-cysteine, pH 7.8)20 g cysteineH2O to 970 mlAdjust pH to 7.8 with 6 M NaOHPrepare fresh just before use

DNaseI, 1 mg/ml10 mg DNase I10 ml H2OStore 1 week at 4°C

DTT, 1 M0.77 g of DTT5 ml H2OKeep on ice. Dissolve the DTT precipitate by moderate heating just before use.Prepare fresh.

Fix solution600 µl 80% glycerol300 µl 37% formaldehyde100 µl 10× MMR (see recipe)0.5 µl 10 mg/ml Hoechst dyeAlways prepare fresh on day of use

Glycogen, 150 mg/ml750 mg glycogenDissolve in 5 ml acetate buffer (see recipe)Aliquot and store 1 year at −20°C

Aliquots can be repeatedly thawed and refrozen


13.4.18

Movement ofNuclei

Gradient buffer3.02 g PIPES (10 mM final)H2O to 950 mlAdd:2 ml 0.5 M EDTA stock solution (1 mM final)1 ml pure 2-β-mercaptoethanol (1% final)10 ml of 10% Triton X-100 stock solution (0.1% final)Bring to pH 7.2 with KOHH2O to 1 literUse immediately to prepare sucrose gradient solutionsStore remainder 1 year at −20°C

HCG, 500 U/ml10,000 U human chorionic gonadotropin (Sigma)Add 10 ml sterile distilled waterKeep sterileStore at 4°CUse within 10 days

HEPES/KOH, pH 7.5 (1 M)59.6 g HEPES200 ml H2OAdjust pH to 7.5 with concentrated KOHH2O to 250 mlFilter sterilizeStore 1 year at −20°C in 5-ml aliquots

K-PIPES, pH 7.2, 0.5 M, 1 mM EDTA7.55 g PIPES40-ml H2OAdd 0.1 ml 0.5 M EDTA stock solutionBring pH to 7.2 with KOHH2O to 50 mlStore 1 year at −20°C in 2-ml aliquots

Lysis buffer0.5 ml Tris⋅HCl, pH 8.0, 1 M stock (APPENDIX 2A; 1 mM final)490 ml H2O0.5 ml pure β-mercaptoethanol (0.1% v/v final)2.5 ml pure NP-40 (0.5% v/v final)51 µl 4.9 M MgCl2 (Sigma; 0.5 mM final)87.1 g PMSF0.5 ml 10 mg/ml stock aprotinin50 µl 10 mg/ml stock leupeptin50 µl 10 mg/ml stock pepstatinH2O to 500 ml

This buffer can be freshly prepared or stored in aliquots at −20°C.

NOTE: NP40 is chemically identical to IGEPAL CA-630 (Sigma).


13.4.19

Organelle Motility

Mg-ATP, 50 mM0.275 mg ATP9 ml H2O0.1 g MgCl2

Adjust pH to 7.5 with NaOHH2O to 10 mlStore 1 year at −20°C as 500-µl aliquots

MMR, 10×58.4 g NaCl (100 mM final)1.49 g KCl (20 mM final)2.04 ml of 4.9 M MgCl2 stock solution (Sigma) (10 mM final)2.94 g CaCl2 (20 mM final)372 mg EDTA (0.1 mM final)11.9 g HEPES (50 mM final, pH 7.8)900 ml H2OAdjust pH to 7.8 with 6 M NaOHH2O to 1 literAutoclave and store 6 months at room temperature

For MMR, dilute 10× stock 1:10 with water.

Nocodazole, 10 mMTo prepare 1 ml:3 mg nocodazole1 ml DMSOStore 1 year at −20°C in 50-µl aliquots

PBS, 10×14.2 g Na2HPO4 dissolved in 500 ml H2O2.67 g NaH2PO4 dissolved in 100 ml H2OMix the 2 solutionsAdd 90 g NaClH2O to 1 literCheck that solution is at pH 7.4Store 1 year at −20°C

For PBS: Dilute 10× PBS 1:10 with water.

PBS/10 with 8% sucrose10 ml 10× PBS (see recipe) diluted to 1 liter with H2O (PBS/10)80 g sucroseBring weight to 1 kg with PBS/10Store 1 year at −20°C

PBS/1% (v/v) BSAWeigh 500 mg BSADissolve in 50 ml PBS (see recipe)Filter and store 1 week at 4°C or aliquot and store 1 year at −20°C


13.4.20

Movement ofNuclei

PE bufferTo prepare 500 ml:1.51 g K-PIPES (10 mM final; see recipe)480 ml H2OAdd 1 ml 0.5 M EDTA stock solution (final 1 mM)Bring to pH 7.2 with KOHH2O to 500 mlFilterStore 3 months at 4°C

PMSG, 200 U/mlAdd 5 ml sterile water to vial containing 1000 U pregnant mare serum

gonadotropinStore 2 weeks at 4°C

Protease inhibitor stock (LPC, 10 mg/ml)10 mg each of leupeptin, pepstatin, and chymostatinDissolve in 1 ml DMSOStore 1 year at −20°C in 50-µl aliquots

S-lysis buffer50 ml 2.5 M sucrose (250 mM final)255 µl 4.9 M MgCl2 (Sigma; 2.5 mM final)12.5 ml 2M KCl (50 mM final)5 ml 1 M HEPES-KOH, pH 7.5 (10 mM final)H2O to 500 mlAdjust pH to 7.5 with KOHPrepare fresh

S-lysis-plus buffer150 ml S-lysis buffer (see recipe)150 µl protease inhibitor stock (LPC; see recipe)150 µl 1 M DTT (see recipe)15 mg cycloheximide (final 100 µg/ml)Stir until dissolved and store on ice until use

Wear gloves when handling cycloheximide.

S-lysis-plus/500 mM sucrose3.4 g sucroseS-lysis buffer (see recipe) to 20 mlIncubate 37°C with intermittent vortexing until dissolved20 µl protease inhibitor stock (LPC; see recipe)20 µl 1 M DTT (see recipe)20 mg cycloheximideStore on ice until use

Wear gloves when handling cycloheximide.

Sucrose, 2.5 M (85% w/v)85 g sucroseAdjust to a total volume of 100 ml with H2OAllow overnight for dissolving (at 60°C)FilterStore 6 months at 4°C


13.4.21

Organelle Motility

Sucrose in gradient buffer, 40% (w/w)40 g sucroseGradient buffer (see recipe) to 100 gPrepare fresh or store 6 months at −20°C



COMMENTARY

Background InformationThe microtubule cytoskeleton plays a cru-

cial role in positioning of the nucleus (reviewedin Reinsch and Gonczy, 1998). Nuclear migra-tion processes occur in numerous cell typessuch as muscle, nerve, and somatic dividingcells. In newly fertilized eggs, long-range mi-crotubule-dependent movements of the maleand female pronuclei are essential for the twonuclei to meet and form the zygotic nucleus(Schatten, 1994).

During oogenesis in metazoan organisms,including Xenopus, the centrosome of the eggdegenerates (Schatten, 1994). At fertilization,the basal body of the sperm flagellum convertsto the centrosome by recruiting pericentriolarmaterial from the egg cytosol, and nucleates alarge microtubule aster. The male pronucleus(Fig. 13.4.8, open circle) remains at the centerof the aster, which grows to fill the egg, thustransporting the male pronucleus to the centerof the egg. The female pronucleus (Fig. 13.4.8,shaded circle) has no associated centrosome or

microtubule-nucleating activity. Instead, the fe-male pronucleus translocates along the micro-tubules of the sperm aster from the cell cortexto the centrosome located in the center of theaster. Thus, the two nuclei meet at the center ofthe aster prior to the first cleavage. Pronucleiof some species traverse hundreds of micronsin a fraction of the first cell cycle (Stewart-Sav-age and Grey, 1982). The motility of the twonuclei differs in that the female pronucleustranslocates along microtubules towards themicrotubule minus-end while the movement ofthe entire sperm aster drives the movement ofthe male pronucleus.

The assay presented here recapitulates themovement of the female pronucleus in vivo andis the first system to demonstrate the transloca-tion of nuclei along microtubules in vitro. Us-ing this extract-based assay, it has been deter-mined that cytoplasmic dynein is required fornuclear movements along microtubules (Re-insch and Karsenti, 1997). This system can befurther used to determine how cytoplasmic

Figure 13.4.8 Movements of the male and female pronuclei in newly fertilized eggs. Schematicshowing the microtubule-mediated positioning events of the male and female pronuclei in a typicalmetazoan early embryo. See text for details. Microtubules are represented by lines, and the maleand female pronuclei by open and shaded circles respectively.


13.4.22

Movement ofNuclei

dynein is targeted to the nuclear membrane tomediate the organelle-like movement along mi-crotubules.

Until the development of this assay, micro-tubule-dependent nuclear motility had onlybeen studied in vivo because of the complexityof the process. Perturbation studies in vivo haveused pharmacological agents to demonstratethe role of the microtubule cytoskeleton inpronuclear movement (Aronson, 1971; Schat-ten, 1982; Rouviere et al., 1994). Genetic stud-ies in C. elegans have only very recently begunto address this process (Gonczy et al., 1999b).Recent experiments using RNA-mediated inhi-bition of expression in C. elegans embryos haveconfirmed the role of dynein and associatedproteins in pronuclear migration (Gonczy et al.,1999a). These genetic experiments, thus, vali-date the biochemical approach presented hereto demonstrate that dynein is required for themotility of synthetic nuclei along microtubules(Reinsch and Karsenti, 1997), and indicate thatthis approach is a valid means to identify can-didate molecules involved in complex proc-esses in the early embryo.

There were two important constraints on thechoice of nuclei for the motility assay. First, itis important to use nuclei lacking an associatedcentrosome to resemble the female pronucleus.Nuclei purified from many tissues and fromcells in culture generally have an associatedcentrosome whose activity could potentiallygenerate motility events resembling those ofthe male rather than the female pronucleus(Reinsch and Gonczy, 1998). Second, it is im-portant to choose a source of nuclei that wouldbe competent for motility. Nuclei from manytissue sources may not be functional for motil-ity if nuclear migration events are not occurringin the selected tissue at the time of purification.

Interphase Xenopus egg extracts spontane-ously assemble nuclei from soluble and mem-brane components stored in the egg (Forbes etal., 1983; Newport, 1987; Newmeyer and Wil-son, 1991). Upon the addition of DNA or chro-matin to a lysed extract, nuclei containing dou-ble nuclear membranes, a nuclear lamina, andnuclear pores are quickly assembled (UNIT 11.10).These nuclei are capable of nuclear import,DNA replication, and normal mitotic disassem-bly. The most commonly used and convenientsubstrate for nuclear assembly studies is de-membranated sperm nuclei. However, thesperm has an associated basal body that con-verts into a centrosome in the egg (or extract).Sperm nuclei are therefore not appropriate sub-strates for this assay. Recently, magnetic beads

coupled to plasmid DNA have been demon-strated to function as artificial chromosomescapable of inducing nuclear assembly in inter-phase extracts and bipolar spindles in mitoticextracts (Heald et al., 1996; Reinsch andKarsenti, 1997; UNIT 11.13). Since DNA beadsare paramagnetic, they can be simply retrievedfrom the extract. Centrosomes do not sponta-neously assemble in these extracts and eggscontain no centrosomes. Therefore, the nucleiassembled around magnetic beads in this ex-tract lack an associated centrosome. These syn-thetic nuclei provide a novel approach to study-ing nuclear motility since they allow rapid andgentle isolation of large numbers of nuclei in amotility competent state.

An alternative to magnetic bead nuclei, is touse DNA purified from phage lambda as asubstrate for nuclear assembly. The DNA issimply added to a 16,000 × g fresh extract andallowed to assemble (see Support Protocol 3,steps 1 through 11). However, the nuclei mustthen be purified over sucrose gradients (S. Re-insch, unpub. observ.). The advantage of thelambda DNA–nuclei is that they are muchsmaller than the DNA-bead nuclei used hereand more suitable for certain analyses such aselectron microscopy.

The benefits of using this extract-based as-say system are numerous. This is an ideal sys-tem for the identification of the molecular com-ponents involved in nuclear motility as it isamenable to biochemical dissection of bothnuclear and cytoplasmic components and al-lows high-resolution microscopic observationof nuclear motility.

This assay allows pronuclear migration tobe studied in isolation of other simultaneous orupstream processes. Both microtubules and cy-toplasmic dynein function in numerous proc-esses that occur both prior to and after fertili-zation. By using this assay, the molecular com-ponents that are specific to this process as wellas those which are shared with other processescan be identified. The ease of biochemical ma-nipulation of these extracts allows experimentsthat are impossible in intact cells or organisms.

Nuclear motility can be observed usingmuch cruder preparations from Xenopus eggsthan those described here (S. Reinsch, unpub.observ.; Murray et al., 1996). However, thisassay provides a means to define the minimalessential components required for nuclear mo-tility along microtubules.


13.4.23

Organelle Motility

Critical Parameters andTroubleshooting

The system presented here uses largely ho-mologous components. Only the centrosomesare purified from humans rather than frog. Intheory, purified centrosomes from any sourcecan be used. The use of homologous cytosoland nuclei is critical. Cytosol from othersources (e.g., bovine brain, HeLa cells, rat liver)does not support the movement of nuclei as-sembled Xenopus egg extracts (S. Reinsch, un-pub. observ.). The use of heterologous nucleihas not been rigorously tested to determinewhether heterologous nuclei will move on mi-crotubules in Xenopus egg extracts. However,nuclei purified from other sources may undergo"repair" when incubated in Xenopus egg ex-tracts and acquire components from the cytosol(Leno and Munshi, 1994) that may enable themto use the Xenopus cytosolic machinery formotility. It is conceivable then, that they willdemonstrate motility events in this system.However, it is important to realize that nucleipurified from tissues or cultured cells may haveassociated centrosomes. Microtubules nucle-ated from these associated centrosomes maygenerate motility events in their own right thatare independent of the translocation of nucleialong microtubules (reviewed in Reinsch andGonczy, 1998).

As stated previously, nuclear motility per secan be visualized using less purified extract-based components. Freshly prepared low-speed (10,000 × g) extracts support both nu-clear assembly and motility. Visualization ofnuclear motility in this simpler system is a bitmore challenging and requires the use of fluo-rescent components to label microtubules andnuclei. The assay presented here has been de-veloped to allow for biochemical manipula-tions of both the cytosol and the nuclei to dissectthe motility apparatus while simultaneouslyallowing visualization of motility events. It hasalso been optimized so that only frozen aliquot-ted components are used. This allows repro-ducibility from one assay to the next so that theeffects of drugs or other added components canbe determined.

For the motility assay, each component canpresent specific difficulties as outlined below.Once the assay has been optimized and is work-ing well, then it is quite reproducible. To mini-mize difficulties, prepare larger stocks withaliquots that are the appropriate size for indi-vidual experiments. Never refreeze aliquots.Prepare a new stock (centrosomes, nuclei,HSS) well before the last aliquots are used. Try

to switch out only one component at a time.Test the new batch of the component alongsidethe old batch to ensure that it functions properly.

Careful biochemical preparation of the indi-vidual components of the Basic Protocol isabsolutely critical for the success of the quan-titative motility assay. Several steps are criticalin preparing extracts of Xenopus eggs. Pleaseread the general references for handlingXenopus and extract preparation (Murray,1991; Newmeyer and Wilson, 1991; UNIT 11.10).It is also worthwhile to understand the scientificliterature on using Xenopus extracts for nuclearassembly (for review see Lohka, 1998 and UNIT

11.10). Eggs for preparation of nuclear assemblyextracts and for HSS must be of top quality.Carefully read each support protocol and haveall materials on hand before starting the prepa-rations. Do not let the preparations sit on ice forlong periods. Proceed immediately through thesteps to the finished frozen product.

One of the most critical parameters in pre-paring fractionated extracts for nuclear assem-bly is to avoid contaminating the nuclear mem-branes with mitochondria. The mitochondriacontribute to apoptotic events (Newmeyer etal., 1994; UNIT 11.12). The mitochondrial layerappears as grayish as opposed to the nuclearmembranes, which have a golden tint. Duringthe 200,000 × g centrifugation, sometimes themembranes do not band optimally and multiplebands are visible. Recentrifuge for a longertime to allow better separation. Too low a su-crose concentration in the extract can also contrib-ute to this. The prep can be done on a smaller scaleusing a TLS-55 rotor instead of an SW-55 rotor.

For nuclear assembly using magnetic beads,the quality of DNA-beads is very important. Besure to precisely follow the instructions in UNIT

11.13 for bead preparation. By Hoechst staining,the DNA should make a uniform bright rimaround the bead. There should not be a punctatepattern around the bead. Make sure to use longDNA fragments. An 8-kb fragment works verywell. Shorter fragments do not assemble wellinto nuclei. The DNA must be filled in withbiotin at only one end so that only one endattaches to the bead. This assures good chro-matin assembly on the DNA bead. DNA-beadscan be stored for months at 4°C. Do not freeze.

When assembling the bead-nuclei fromfractionated extracts make sure that DNA-beads are well suspended in extract so that theydon’t clump too much. Otherwise all the nucleiwill contain many beads. An average of 1 to 3beads per nucleus is optimal for the motilityassay. Glycogen is essential for assembly of


13.4.24

Movement ofNuclei

chromatin onto naked DNA (Hartl et al., 1994).The source of the glycogen is not important.Commercial oyster glycogen works quite well.The length of time for the nuclear assemblyreaction is not critical and can be from 1 to 3hr. The longer the nuclei are left in the assemblyreaction, the larger they tend to get. They donot acquire more beads, but they will importproteins from the extract and become moreswollen. This is not really a problem and oftenis helpful for visualizing uptake of the transportsubstrate TRITC-BSA-NLS. When perform-ing the transport reaction, one should be ableto clearly see uptake of nuclear transport sub-strate into 90% to 95% of nuclei. After assem-bly, nuclei can be stored overnight at 4°C with-out loss of function. Longer storage at thistemperature is not advised. If one encountersdifficulties in assembling nuclei from fraction-ated components, then try a different substrate,such as sperm nuclei (see UNIT 11.13 for prepara-tion) or lambda DNA to be convinced that theproblem is not the DNA-bead substrate.

The centrosome preparation presented herehas been optimized for high yield and highconcentration of centrosomes. It is a variationof a published method (Moudjou and Bornens,1998). Other preparations of centrosomes havenot been tested for use in this assay, but wouldbe interesting to compare (e.g., Blomberg-Wir-schell and Doxsey, 1998). In theory, any cen-trosome preparation should be acceptable foruse in this assay as long as robust microtubuleasters are generated upon addition of HSS. Forthis particular preparation, it is essential thatthe lymphocytes be grown in exponentialphase. The treatments with the cytoskeletalinhibitors, nocodozole and cytochalasin, sepa-rate the centrosome from the nuclear membraneso that centrosome and nucleus do not copurify.The incubation time for this treatment allowsdepolymerization of microtubules and actinfilaments without causing cells to accumulatein mitosis. The washes with high osmoticstrength buffers cause the cells to be very frag-ile. These steps should be done as rapidly aspossible to minimize cell lysis. However, do notpipet so vigorously that bubbles are produced.The centrifugation steps are straightforwardand can be adapted for other rotors and vol-umes. Do not leave the centrosomes unfrozenfor long periods. Freeze the large aliquots asquickly as possible. Thaw only once to re-ali-quot and freeze again. The size for the smallaliquots should be dictated by experiment de-sign. Do not refreeze the small aliquots.

Preparations of HSS can vary in activitylargely due to egg quality. Cycloheximide isadded to ensure that the extract stays in inter-phase. It is important to add sucrose as a cryo-protectant to the 16,000 × g extract beforefreezing. The prep is frozen as a 16,000 × gextract for several reasons. First, this takes upless freezer space, and the subsequent dilutionand centrifugation steps to be done on the dayof preparation are very reproducible. Second,preps that were carried through the subsequentdilution and centrifugation and then frozen asaliquots did not function as well in the motilityassay. On the day of the motility assay, thedilution before high-speed centrifugation en-sures that all of the membranes are sedimented.This allows optimal DIC imaging of nucleiwithout extraneous membranes. The dilutionrecommended here has been optimized for mi-crotubule aster assembly. Too low a dilutioncauses a lot of spontaneous microtubule assem-bly. Sometimes asters generated in HSS be-come fragmented before the 20-min time point.Throw these batches of cytosol away since theywill consistently give the same result, and pre-pare a new batch of 16,000 × g extract. Thesepoor-quality batches often also show highapoptotic activity. Try keeping egg batchesfrom different frogs separate all the waythrough the preparation if poor quality cytosolis a problem.

The quantitative assay for nuclear motilityrequires nuclei to accumulate at centrosomes.Therefore, robust aster morphology with nicespacing between the asters is crucial. Humanlymphocytes assemble asters with only a fewmicrotubules in the presence of pure tubulin.Incubating centrosomes in HSS allows pericen-triolar material to be recruited to the centro-somes. After incubation in Xenopus egg extract,their nucleation capacity increases signifi-cantly (Buendia et al., 1992) so that robustmicrotubule asters are generated. Plating den-sity of the centrosomes can affect the outcomeof the assay. Steps for optimizing this parameterare outlined in Support Protocol 3. The astersshould be spaced so that there are not more than1 to 3 centrosomes/50-µm field. This allowsnuclei to reach centrosomes so that quantitationis possible. If asters are too widely spaced, thenthere will be a lot of free microtubule nucleationand random movement of the nuclei rather thannuclei moving to and accumulating at centro-somes. Make sure the incubation to allow cen-trosomes to attach to the glass coverslip isconsistently 5 min. Several blocking agentshave been tested; casein gives satisfactory re-


13.4.25

Organelle Motility

sults to prevent nuclei from sticking. The caseinalso inhibits dynein present in HSS from bind-ing to the coverslip. This decreases the amountof microtubule gliding that occurs.

If robust, well-spaced asters are not ob-tained, the problems can either be in the densityof the centrosomes or more likely in the prepa-ration of HSS as described above. Addingagents such as drugs and antibodies can alsoseriously affect the ability of asters to be nucle-ated and should only be added after the nuclea-tion step. For example, addition of agents thatinhibit dynein activity cause random micro-tubules to assemble, rather than preferentialassembly off centrosomes (S. Reinsch, unpub.observ.). If one does not plan to use the motilityassay as a quantitative assay, but only for visu-alizing nuclear movements without perturbingagents, then the nucleation step can be com-pletely omitted. Many agents also compromisemicrotubule assembly. For example, BAPTA,a calcium chelator, completely inhibits micro-tubule assembly in this assay (S. Reinsch, un-pub. observ.). Since the microtubules are dy-namic in this assay, any factor that affects mi-crotubule dynamics may compromise thesuccess of the assay.

Anticipated ResultsThe nuclear motility assay allows clear

documentation of nuclear motility events.When bead-nuclei are purified on the magnet,they do come with a considerable amount ofassociated endoplasmic reticulum (ER). There-fore, both nuclear movements and ER tubuleelongation will be visible. In HSS, the nucleimove exclusively towards the minus-ends ofthe microtubules, that is, towards the centro-some. The average rate of movement of nucleiassembled in Xenopus extracts is 1 µm/sec witha range from 0.2 to 1.8 µm/sec. Neither DNA-beads (unassembled) nor chromatin-beads (as-sembled in cytosol without adding membranes)will move to the microtubule minus-ends. Inthe presence of inhibitors of dynein (e.g., >15µM vanadate) minus-end directed movementsare blocked. The nuclei then are moved to theaster periphery due to the action of growingmicrotubules. This is not plus-end directedmovement. Instead nuclei appear to be pulledout by growing microtubules due to ER tubuleattachment to the tips of growing microtubules(Waterman-Storer et al., 1995; Reinsch andKarsenti, 1997).

For the quantitative assay, 90% to 95% ofthe nuclei should accumulate at centrosomesby the 20-min time point under control condi-

tions. One should be able to easily count be-tween 50 and 100 nuclei in the 10-min countinginterval. Dynein inhibitors significantly inhibitaccumulation at centrosomes (see Fig. 13.4.3;Reinsch and Karsenti, 1997). An antibodyagainst the intermediate chain of cytoplasmicdynein (mAb 70.1, Sigma) also inhibits nuclearmotility and accumulation of nuclei at centro-somes when directly added to HSS (Reinschand Karsenti, 1997). Not all antibodies againstcytoplasmic dynein successfully inhibit nu-clear motility (S. Reinsch, unpub. observ.).

Each assay should be performed at leastthree times to generate statistically significantdata. Once a control reaction has been per-formed on a given batch of centrosomes/nu-clei/HSS, the control reaction should be con-sistent from day to day and should always give90% to 95% accumulation. This includes add-ing buffer to account for added experimentalagents. Therefore, on a given experimental day,the control reaction can be performed once orpossibly twice, to confirm that the system looksthe same as on the previous experimental day,rather than having to perform multiple controlexperiments on each day an experiment is per-formed. This gives more time for experimentalmanipulation (see below).

The HSS should contain no membraneousmaterials. When incubated at room tempera-ture, microtubule assembly should be visiblewithin several minutes using VE-DIC optics.In the absence of centrosomes, only randommicrotubule assembly occurs. In the presenceof centrosomes, microtubules first assemblerandomly, but over the 10-min nucleation pe-riod, the centrosomes acquire pericentriolarmaterial and nucleate more microtubules. By10 min almost all microtubules are nucleatedby centrosomes. Microtubule dynamics do oc-cur and by VE-DIC both growing and shrinkingmicrotubules should be visible.

The typical yield for this centrosome prepa-ration is 30% to 50%. Therefore, a 2-liter prepa-ration of lymphocytes (3 × 109 cells) generallyyields ∼109 centrosomes with several fractionscontaining centrosomes at or above the optimalconcentration (∼2 × 108/ml). A single centro-some preparation generates enough centro-somes for at least 1000 motility assays.

Eggs from one frog (5 ml dejellied eggs)generally yields 1 ml of 16,000 × g cytosol.Each milliliter of cytosol generates ∼3 ml ofHSS. Each assay uses 20 µl of HSS. Therefore,one frog yields enough HSS for ∼150 assays.

Eggs from one frog (5 ml dejellied eggs) willgenerally yield ∼300 µl of cytosol-200 (and


13.4.26

Movement ofNuclei

membranes). This will make a ∼700-µl nuclearassembly reaction, which is aliquotted by 10 µlat the end. Each 10-µl aliquot contains nucleifor one motility assay. Therefore, eggs from onefrog generate nuclei for ∼70 motility assays.

Time ConsiderationsOnce all of the support protocols have been

followed to generate the components of themotility assay and the learning period has beenovercome, then each motility assay takes ∼40min to perform. One to 2 hr of preparation timeis required before each assay period. Therefore,only a few experimental samples can be com-pared on 1 day if assays are performed intriplicate with appropriate buffer controls.

The centrosome preparation requires mini-mal time for 10 to 14 days to grow the lympho-cytes, 1 day to prepare the buffers, and 1 day todo the actual preparation. Titration of the cen-trosomes in the motility assay requires anotherfull day, but should only be done once all theother components have been prepared.

Preparation of fractionated extracts for nu-clear assembly requires 1 full day. Frogs mustbe primed 4 to 10 days in advance with PMSGand the night before the prep with HCG.

For time considerations in preparing mag-netic DNA-beads, see UNIT 11.13. Working outthe conditions for optimal nuclear assemblyusing DNA beads and fractionated extracts canoptimistically be done in an afternoon, but oftencan take several tries. Actual preparative scalenuclear assembly takes half a day.

Preparation of 16,000 × g extracts for gen-eration of HSS takes half a day once frogs areprimed as above.

Preparation of slide chambers for motilityassays takes parts of a day, during which manyother activities can be undertaken simultaneously.

Time considerations for familiarization withVE-DIC and optimization for visualization ofmicrotubules are beyond the scope of this pro-tocol. Please refer to UNITS 4.1 & 13.1 and to Salmonand Tran (1998) and Walker et al. (1988).

Literature CitedAronson, J.F. 1971. Demonstration of a colcemid-

sensitive attractive force acting between the nu-cleus and a center. J. Cell Biol. 51:579-583.

Blomberg-Wirschell, M. and Doxsey, S.J. 1998.Rapid isolation of centrosomes. Methods Enzy-mol. 298:228-238.

Bornens, M., Paintrand, M., Berges, J., Marty, M.C.,and Karsenti, E. 1987. Structural and chemicalcharacterization of isolated centrosomes. CellMotil. Cytoskeleton 8:238-249.

Buendia, B., Bre, M.H., Griffiths, G., and Karsenti,E. 1990. Cytoskeletal control of centrioles move-ment during the establishment of polarity inMadin-Darby canine kidney cells. J. Cell Biol.110:1123-1135.

Buendia, B., Draetta, G., and Karsenti, E. 1992.Regulation of the microtubule nucleating activ-ity of centrosomes in Xenopus egg extracts: Roleof cyclin A-associated protein kinase. J. CellBiol. 116:1431-1442.

Euteneuer, U. and Schliwa, M. 1985. Evidence foran involvement of actin in the positioning andmotility of centrosomes. J. Cell Biol. 101:96-103.

Evans, L., Mitchison, T., and Kirschner, M. 1985.Influence of the centrosome on the structure ofnucleated microtubules. J. Cell Biol. 100:1185-1191.

Forbes, D.J., Kirschner, M.W., and Newport, J.W.1983. Spontaneous formation of nucleus-likestructures around bacteriophage DNA microin-jected into Xenopus eggs. Cell 34:13-23.

Gant, T.M. and Wilson, K.L. 1997. Nuclear assem-bly. Annu. Rev. Cell. Dev. Biol. 13:669-695.

Gonczy, P., Pichler, S., Kirkham, M., and Hyman,A.A. 1999a. Cytoplasmic dynein is required fordistinct aspects of MTOC positioning, includingcentrosome separation, in the one cell stageCaenorhabditis elegans embryo. J. Cell Biol.147:135-150.

Gonczy, P., Schnabel, H., Kaletta, T., Amores, A.D.,Hyman, T., and Schnabel, R. 1999b. Dissectionof cell division processes in the one cell stageCaenorhabditis elegans embryo by mutationalanalysis. J. Cell Biol. 144:927-946.

Hartl, P., Olson, E., Dang, T., and Forbes, D.J. 1994.Nuclear assembly with lambda DNA in fraction-ated Xenopus egg extracts: An unexpected rolefor glycogen in formation of a higher-order chro-matin intermediate. J. Cell Biol. 124:235-248.

Heald, R., Tournebize, R., Blank, T., Sandaltzopou-los, R., Becker, P., Hyman H., and Karsenti, E.1996. Self-organization of microtubules into bi-polar spindles around artificial chromosomes inXenopus egg extracts [see comments]. Nature382:420-425.

Heald, R., Tournebize, R., Vernos, I., Murray, A.,Hyman, T., and Karsenti, E. 1998. In vitro assaysfor mitotic spindle assembly and function. InCell Biology. A Laboratory Handbook, Vol. 2.(J.E. Celis, ed.) pp. 326-335. Academic Press,San Diego.

Leno, G.H. and Munshi, R. 1994. Initiation of DNAreplication in nuclei from quiescent cells re-quires permeabilization of the nuclear mem-brane. J. Cell Biol. 127:5-14.

Lohka, M.J. 1998. Analysis of nuclear envelopeassembly using extracts of Xenopus eggs. Meth-ods Cell Biol. 53:367-395.

Mack, G. and Rattner, J.B. 1993. Centrosome repo-sitioning immediately following karyokinesisand prior to cytokinesis. Cell Motil. Cytoskeleton26:239-247.


13.4.27

Organelle Motility

Merdes, A., Ramyar, K., Vechio, J.D., and Cleve-land, D.W. 1996. A complex of NuMA and cy-toplasmic dynein is essential for mitotic spindleassembly. Cell 87:447-458.

Moudjou, M. and Bornens, M. 1998. Method ofcentrosome isolation from cultured animal cells.In Cell Biology: A Laboratory Handbook, Vol. 2.(J. Celis, ed.) pp. 111-119. Academic Press, SanDiego.

Murray, A.W. 1991. Cell cycle extracts. In Xenopuslaevis: Practical Uses in Cell and Molecular Bi-ology, Vol. 36, Methods in Cell Biology (B.K.Kay and H.B. Peng, eds.) pp. 581-604. AcademicPress, San Diego.

Murray, A.W., Desai, A.B., and Salmon, E.D. 1996.Real time observation of anaphase in vitro. Proc.Natl. Acad. Sci. U.S.A. 93:12327-12332.

Newmeyer, D.D. and Wilson, K.L. 1991. Egg ex-tracts for nuclear import and nuclear assemblyreactions. In Xenopus laevis: Practical Uses inCell and Molecular Biology, Vol. 36, Methods inCell Biology (B.K. Kay and H.B. Peng, eds.) pp.607-634. Academic Press, San Diego.

Newmeyer, D.D., Farschon, D.M., and Reed, J.C.1994. Cell-free apoptosis in Xenopus egg ex-tracts: inhibition by Bcl-2 and requirement for anorganelle fraction enriched in mitochondria. Cell79:353-364.

Newport, J. 1987. Nuclear reconstitution in vitro:Stages of assembly around protein-free DNA.Cell 48:205-217.

Reinsch, S. and Gonczy, P. 1998. Mechanisms ofnuclear positioning. J. Cell Sci. 111:2283-2295.

Reinsch, S. and Karsenti, E. 1997. Movement ofnuclei along microtubules in Xenopus egg ex-tracts. Curr. Biol. 7:211-214.

Rouviere, C., Houliston, E., Carre, D., Chang, P.,and Sardet, C. 1994. Characteristics of pronu-clear migration in Beroe ovata. Cell Motil. Cy-toskeleton 29:301-311.

Salmon, E.D. and Tran, P. 1998. High resolutionvideo enhanced differential interference contrastmicroscopy. Methods Cell Biol. 56:153-184.

Schatten, G. 1982. Motility during fertilization. Int.Rev. Cytol. 79:59-163.

Schatten, G. 1994. The centrosome and its mode ofinheritance: The reduction of the centrosomeduring gametogenesis and its restoration duringfertilization. Dev. Biol. 165:299-335.

Stewart-Savage, J. and Grey, R.D. 1982. The tempo-ral and spatial relationships between corticalcontraction, sperm trail formation, and pronu-clear migration in fertilized Xenopus eggs. Wil-helm Roux’s Arch 191:241-245.

Tournebize, R., Popov, A., Kinoshita, K., Ashford,A.J., Rybina, S., Poznikovsky, A., Mayer, T.U.,Walczak, C.E., Karsenti, E., and Hyaman, A.A.2000. Control of microtubule dynamics by theantagonistic activities of XMAP215 and

XKCM1 in Xenopus egg extracts. Nat. Cell Biol.2:13-19.

Walczak, C.E., Mitchison, T.J., and Desai, A. 1996.XKCM1: A Xenopus kinesin-related protein thatregulates microtubule dynamics during mitoticspindle assembly. Cell 84:37-47.

Walker, R.A., O’Brien, E.T., Pryer, N.K., Soboeiro,M.F., Voter, W.A., Erickson, H.P., and Salmon,E.D. 1988. Dynamic instability of individualmicrotubules analyzed by video light micros-copy: Rate constants and transition frequencies.J. Cell Biol. 107:1437-1448.

Waterman-Storer, C.M., Gregory, J., Parsons, S.F.,and Salmon, E.D. 1995. Membrane/microtubuletip attachment complexes (TACs) allow the as-sembly dynamics of plus ends to push and pullmembranes into tubulovesicular networks in in-terphase Xenopus egg extracts. J. Cell Biol.130:1161-1169.

Key ReferencesReinsch and Gonczy, 1998. See above.

This review describes the different mechanisms thatdrive microtubule-mediated nuclear motility eventsin different cell types and different organisms. Nu-clear motility following fertilization is particularlyemphasized.

Reinsch and Karsenti, 1997. See above.

This work demonstrates that nuclei can translocatealong microtubules in Xenopus egg extracts similarto other organelles, and that cytoplasmic dyneincytoplasmic dynein drives the translocation of nu-clei along microtubules.

Internet Resourceshttp://current-biology.com/supmat/cub/bb7325s1

.mov

http://current-biology.com/supmat/cub/bb7325s2.mov

http://current-biology.com/supmat/cub/bb7325s3.mov

These movies are supplemental material for Reinschand Karsenti (1997). They are videos of nuclei mov-ing along microtubule asters as described in theBasic Protocol. The nuclei used in these movies wereassembled as described in Support Protocols 3 and 5.

http://www.indiana.edu/∼elegans/

This is the webpage for the laboratory of SusanStrome at Indiana University. Movies within this siteshow pronuclear migration in vivo in the nematodeC. elegans.

Contributed by Sigrid ReinschNASA-Ames Research CenterMoffett Field, California


13.4.28

Movement ofNuclei

UNIT 13.5Measuring Dynamics of Nuclear Proteins byPhotobleaching

Photobleaching techniques offer the possibility of obtaining information on molecularmotion and interactions in a specific part of a cell (McNally and Smith, 2002). The threemain advantages of photobleaching techniques are fast experimental turn around, goodspatial and temporal resolution, and the ability to measure kinetics inside of living cells.The main disadvantage of these techniques is the requirement for fluorescently taggedproteins (in this case by creating a gene fusion with GFP). Alternatively, other geneticallyengineered fluorescent tags may be used for a fusion with the protein of interest—e.g.,cyan fluorescent protein (CFP) or yellow fluorescent protein (YFP). Also, in some cases,addition of this moiety can functionally alter the protein. Thus, prior to proceeding withphotobleaching studies, the fusion protein must be rigorously tested to ensure it has thesame properties and function as its native counterpart.

Ideally, a fusion protein is tested in a functional in vivo assay. The best is a functionalcomplementation assay, for example, in a knock-out cell line or yeast. However, for manymammalian proteins such assays are not readily available. A minimal requirement fortesting GFP-fusion proteins is their stability and proper localization in cells. The stabilityof a GFP-fusion protein can be tested by immunoblot (UNIT 6.2), using either a specificantibody against the endogenous protein of interest or an antibody against GFP. Animmunoblot ensures that the protein is of the expected molecular weight and that it is notdegraded upon expression. Furthermore, the GFP-fusion protein must also colocalize withits endogenous counterpart. If the immunoblot shows degradation products of the proteinchimera or the protein is mislocalized, the GFP-fusion protein is likely not fully func-tional. In this case, it is worth trying to introduce the GFP tag at the opposite end of theprotein molecule. In some cases, the introduction of a longer linker between the proteinand GFP is also helpful.

Three major methods of photobleaching microscopy are commonly used: fluorescencerecovery after photobleaching (FRAP; see Basic Protocol), fluorescence loss in pho-tobleaching (FLIP; see Alternate Protocol 1), and inverse fluorescence recovery afterphotobleaching (iFRAP; see Alternate Protocol 2). As summarized in Table 13.5.1, each ofthese techniques has specific characteristics permitting the determination of distinct particularparameters of protein behavior in vivo. In addition, transfection methods are given forintroducing GFP-expression vectors into mammalian cells by electroporation (see SupportProtocol 1) or by lipofection (see Support Protocol 2). Finally, since quantification ofGFP-fusion proteins is critical, a method for the determination of the number of GFPmolecules in a single, living cell is also described (see Support Protocol 3).

NOTE: The reader is referred to UNIT 4.5 for a general discussion of confocal microscopy.

Supplement 18

Contributed by Miroslav Dundr and Tom MisteliCurrent Protocols in Cell Biology (2003) 13.5.1-13.5.18Copyright © 2003 by John Wiley & Sons, Inc.

Table 13.5.1 Comparison of Photobleaching Methods for Detecting Movement of NuclearProteinsa

Mobility Diffusionconstant

Immobilefraction Distinct pools Compartment

continuity

FRAP +++ +++ +++ + +FLIP +++ + + +++ +++iFRAP − − + +++ −a+++ optimal, + suitable, −not suitable.

13.5.1

Organelle Motility

BASICPROTOCOL

FLUORESCENCE RECOVERY AFTER PHOTOBLEACHING (FRAP)

In a FRAP experiment, an intense focused laser beam of the appropriate wavelengthbleaches a relatively small region of interest. The laser light irreversibly bleaches thefluorescent signal of molecules in the region of interest and, immediately followingbleaching, the recovery of the fluorescence signal in the bleached area is monitored usingan attenuated laser beam. The signal in the region of interest is measured and a fluores-cence recovery curve Frec(t) (fluorescence intensity as a function of time after pho-tobleaching) is generated (Fig. 13.5.1). Acquisition of a FRAP data set involves threephases: prebleach images, bleach pulse, and postbleach monitoring of fluorescencerecovery.

FRAP experiments are most easily performed on any modern confocal microscope (e.g.,Zeiss 510 and newer, Leica SP and newer), as these microscopes have integrated FRAProutines. Alternatively, it is possible to custom write macros for many older microscopesthat will automatically perform FRAP routines (Ellenberg et al., 1998). This protocol, aswell as its alternatives (see Alternate Protocols 1 and 2), are designed for the Zeiss LSM

ROI

bleachpre-bleach monitoring

T

B

prebleach

bleach

Flu

ores

cenc

e in

tens

ity

Recovery time

immobile fraction

mobile fraction

highly mobile

Recovery Recovery

intermediate immobile

Recovery

Figure 13.5.1 General scheme of a FRAP experiment. A cell is imaged before a short bleachpulse is applied to a defined region of interest (ROI). The recovery of the average fluorescencesignal in the ROI is then monitored. To quantify and normalize FRAP data, the average fluorescencesignal in the ROI, average total nuclear signal (T), and average signal in a random area outside ofthe cell for background correction (B) are measured during the entire experiment. The FRAP curveshows the prebleach intensity, depth of bleach, and recovery of the signal. The immobile fraction isdetermined as the difference between the prebleach signal and the signal after complete recovery.Based on different profiles of recovery curves, proteins can be considered highly mobile withvirtually no immobile fraction, intermediately mobility with an immobile fraction, or immobile.


13.5.2

MeasuringDynamics of

Nuclear Proteinsby Photobleaching

510 confocal microscope, but similar protocols can be applied to other confocal micro-scopes.

When FRAP experiments are carried out on confocal microscopes, the fluorescencerecovery in the region of interest can be considered largely the result of two-dimensionalexchange of fluorescent protein molecules within the same focal plane as the bleachedarea. This condition is approximately valid when objectives with high-numerical-aperture(N.A. of 1.2-1.45) lenses are used; a large cone of out-of-focus light bleaches thefluorescent molecules above and below the focal plane so that only nonbleached mole-cules from the focal plane participate in the recovery in the region of interest (Ellenberget al., 1998).

There are several essential conditions that must be fulfilled for a successful FRAPexperiment. (1) The fluorescent signal to be bleached must be clearly detectable over anybackground signal. (2) The photobleaching must be fast relative to the period of recoveryto provide sufficient temporal resolution for analysis of the recovery curve and to allowmeasurement of the half-time of recovery. (3) The monitoring beam must be of lowintensity to minimize photobleaching. To obtain a sufficiently high signal during therecovery period, the sensitivity of the photomultiplier tube may be increased.

Materials

2- or 4-chambered Lab-Tek II coverglass (Nalgene Nunc) containing cells, 50% to70% confluent

Complete normal growth medium with 25 mM HEPES, but without phenol red(UNIT 1.2)

Confocal microscopeAir stream incubator (Nevtek)

1. On the stage of a confocal microscope, place a 2- or 4-chambered Lab-Tek IIcoverglass containing cells in complete normal growth medium with 25 mM HEPES,but without phenol red. Maintain growth temperature with a Nevtek air incubator.Examine the cells with fluorescent light of the appropriate wavelength.

2. Choose the appropriate lens (at least 63× magnification) and zoom setting to clearlyobserve the region of interest.

If the same zoom is used for all observed cells, one can easily compare individual data setswithout correction for bleach-area size.

3. Image a cell of interest using a low level of laser power to prevent photobleaching.

On the Zeiss LSM 510 microscope, 20% to 50% output from a 40-mW laser with the beamattenuated to 0.1% is routinely used for the image acquisition.

4. Choose a small pinhole diameter.

The authors recommend using a pinhole diameter in the range of 1 Airy unit (minimalresolvable distance which corresponds to the diameter of the peak of the bright spot of thepoint source of light; UNIT 4.1). A larger pinhole diameter improves the signal intensity, butincreases the optical thickness of the collected section. This could affect monitoring of thesignal in the region of interest by collecting data from a thicker optical section includingunbleached fluorescent molecules. When a smaller pinhole diameter is used, the monitoringof signal recovery is restricted to the optical section most completely bleached. Therefore,it is advisable to use the smallest pinhole diameter which gives a sufficient signal-to-noiseratio.


13.5.3

Organelle Motility

Setup imaging sequence5. Adjust the intensity of fluorescence signal to slightly below the saturation level by

adjusting the gain on the photomultiplier tube.

Use at least 8- but preferably 12-bit imaging. It is important to ensure that no pixels aresaturated since the change in fluorescence intensity in these pixels cannot be measuredaccurately.

6. Adjust laser power to 100% for bleaching.

The bleach time should be minimized. The efficiency of the bleach should be tested on afixed sample. The average intensity in a bleached spot in a fixed sample should be reducedby at least 70% compared to its surroundings. If the bleach depth is insufficient, use several,rapid, consecutive bleach pulses. If the bleach spot in a living cell is distinct from the bleachspot in a fixed cell, do not increase the bleach time or intensity. The discrepancy is animportant indication that the observed protein contains a very rapidly moving fraction.These highly mobile molecules enter the bleached spot between the time the bleach pulseis terminated and the acquisition of the first image. Recovery of freely mobile proteins inthe nucleus can occur on the time scale of ≤50 msec.

7. Adjust the number of prebleach images for five to ten images and acquire five to tenprebleach images.

It is important to acquire more than one prebleach image, as many fluorescent moleculesare disproportionately bleached when first excited.

8. Adjust the number of postbleach intervals and the time between images.

The number of intervals is determined by the resolution needed. The length of intervalsshould cover the curve until it reaches a plateau. Be careful not to collect an unnecessarilyhigh number of images as this will bleach the sample.

9. Define the region to be bleached.

The size of the bleach region is critical for quantitative analysis of FRAP recovery. Theauthors recommend using a small circular spot as the bleach region over the desired regionof interest in the nucleus. The size of the bleach region should be small compared to thesize of the nucleus. The advantage of a circular bleach spot is its symmetry, which permitsfinding it even when the cell moves. To check the dimensions of the bleach region, it isnecessary to fix the sample, bleach the region of interest, and collect a series of opticalsections through the cell. When the bleached region is monitored for signal recovery, thesame size area or slightly smaller can be used for measurement of signal recovery. Usinga larger monitoring area allows one to obtain the recovery data even if the bleached areamoved slightly during monitoring; however, the relatively unchanged fluorescence intensityin the neighborhood of the bleached region reduces the sensitivity of the measurement.

10. Initiate data collection (i.e., prebleach images, bleach, postbleach images) andmonitor the recovery of the fluorescence signal. Check the position of the cell andthe focal plane to ensure that the region of interest is not moving from its originalposition due to focal-plane movement.

It is essential to determine that recovery is complete—i.e., the curve should reach a clearplateau. The shape of a recovery curve should resemble the one in Figure 13.5.1. If therecovery is not complete, it is essential to increase the monitoring time of recovery byincreasing the number of acquired images during monitoring or the time between images(step 8).

Perform quantitative analysis of FRAP11. Measure the average intensity of the region of interest at each time point (It).

Most confocal microscope software packages allow the researcher to perform this meas-urement on a whole stack of time-lapse data.


13.5.4



12. To correct for signal bleaching during monitoring, measure the average intensity ofthe entire nucleus at each time point (Tt).

13. Measure the background intensity (Bt) in a randomly selected region outside of thecell at each time point.

14. Calculate the relative intensity for each time point:

Irel =(T0 − B0) × (I(t) – B(t))/(T(t) − B(t)) × (I0 – B0)

where T0 is the average intensity of the entire nucleus during prebleach and I0 is theaverage intensity of the region of interest during prebleach (step 6).

This formula is derived in three steps. First, background is subtracted from all measuredvalues, then the fluorescence loss due to monitor bleaching is normalized, and finallyfluorescence intensity is normalized to one.

15. Plot Irel as function of time.

For convenient calculation, it is useful to import FRAP data sheets from the microscopesoftware to a spread sheet program such as Microsoft Excel.

16. Evaluate the individual curves and determine the average and standard deviations foreach time point.

It is advisable to plot all acquired curves before determining the average. Jumps in therecovery curves or declining curves are frequently caused by cell or focal plane movement.These curves can either be discarded from analysis, or if possible, they should be manuallyremeasured. For many applications, it is sufficient to collect data for 10 to 20 individualcells. Typical error bars are on the order of 5% to 10% of the measured value.

Alternative normalization procedures have been published. For example, some investiga-tors assign the value 1 to the prebleach intensity and the value 0 to the postbleach imagein the bleached spot. (Kruhlak et al., 2000). This method is only appropriate for relativelyslow-moving molecules where the signal in the bleached region is reduced by ≥75%. Forfast moving molecules where the reduction of signal might only be ≤50%, this normaliza-tion method overestimates the recovery kinetics.

In the case of FRAP analyses of proteins which are enriched in small nuclear compartmentssuch as the nucleolus or dot-like nuclear bodies and present at only low concentrations inthe surrounding nucleoplasm, the relative fluorescent intensity in the bleached nuclearcompartment can be normalized against the nonbleached nuclear compartment in the samenucleus (Chen and Huang, 2001). For each time point, the relative intensity is determinedby the formula Irel = (NetN10)/(Ne0N1t), where Net and Nlt are the average intensities ofthe bleached compartment and control nonbleached compartment at each time point,respectively, and Ne0 and Nl0 are the average intensities of the bleached compartment andcontrol nonbleached compartment in the same nucleus during prebleach, respectively.

Determine the immobile fraction and diffusion constant17. Calculate the immobile fraction of fluorescent molecules as the difference between

the relative fluorescence intensity in the region of interest after the recovery curvehas reached plateau, and the prebleach fluorescence signal intensity (normalized as100%; Fig. 13.5.1).

Failure to recover to 100% prebleach fluorescence signal intensity even for a proteinwithout immobile fraction such as GFP alone may represent the loss of total nuclearfluorescence due to bleaching during monitoring.

18. Determine the diffusion constant.

The mobility of a protein is characterized by its diffusion constant (D). FRAP experimentsallow the determination of D assuming lateral diffusion in the focal plane (Axelrod et al.,


13.5.5

Organelle Motility

1976); however, it is essential to understand that the measured diffusion constant is onlyan apparent diffusion constant. The overall mobility of a protein is not only determined byits diffusional properties, but more significantly, by its functional properties such asincorporation into multiprotein complexes and binding to relatively immobile structuressuch as chromatin or a karyoskeleton (Misteli, 2001). Therefore, although a diffusionconstant is a convenient method to compare the recovery kinetics of various proteins, onemust be cautious in interpreting diffusion constants as a true indicator of the translationalmobility of a protein. For discussion of diffusion constants see Lippincott-Schwartz et al.(2001). For determination of diffusion constants using a bleached strip of defined size seeEllenberg et al. (1998) Alternatively, diffusion constants can be determined directly fromfitting the experimental data to the theoretical curve by computer simulation (Phair andMisteli, 2000; Houtsmuller and Vermeulen, 2001; Reits and Neefjes, 2001).

ALTERNATEPROTOCOL 1

FLUORESCENCE LOSS IN PHOTOBLEACHING (FLIP)

In FLIP, a region in the nucleus is repeatedly bleached and the loss of fluorescence in amonitoring region of interest some distance away from bleached region is recorded (Fig.13.5.2). The basis of FLIP is the bleaching of mobile molecules as they pass through ableached region. If all or a fraction of the fluorescently-tagged molecules in the nucleusare immobile, they will never enter or exit the repeatedly bleached region; therefore, theywill not be bleached and cannot distribute the bleached signal elsewhere. As a conse-quence, the fluorescence signal in the regions surrounding the bleached spot will remainconstant. In contrast, if molecules are highly mobile, they will eventually pass throughthe bleached region. The fluorescence signal in the nucleus will decrease. If all fluores-cently-tagged molecules inside of the monitoring region of interest move freely into thebleached region, repeated bleaching will reduce the fluorescent signal in the monitoringregion of interest at a constant rate. On the other hand, if multiple, kinetically distinctfractions of molecules exist, the loss curve will contain multiple distinct slopes. FLIP isalso a particularly effective technique for testing continuity between nuclear compart-ments such as nucleoli, splicing factor compartments, Cajal bodies, or PML bodies.

FLIP is complementary to FRAP. Since in FLIP the signal in an unbleached region ismeasured, the technique is often used to ensure that the mobility of a protein of interestobserved in FRAP experiments is not due to photodamage of the protein at the bleached spot.

See Basic Protocol for materials.

1. Set up the cells on the confocal microscope as described (see Basic Protocol, steps 1and 2).

2. Image two cells of interest close to each other using a low level of laser power toprevent photobleaching.

On the Zeiss LSM 510 microscope, 20% to 50% output from a 40-mW laser with the beamattenuated to 0.1% is routinely used for the image acquisition.

3. Adjust settings as described for FRAP (see Basic Protocol, steps 4 to 9).

A successful FLIP experiment depends on the successful loss of fluorescence in regionsoutside the bleached region. It is therefore more practical to use bleach areas that are largerin size than the ones used in the FRAP experiments.

4. Initiate data collection (i.e., prebleach images, bleach, postbleach images) andmonitor fluorescence loss. Check the position of the cell and the focal plane to besure that the monitoring spot is not moving from the original position.

The acquisition routine should run in a loop between bleach pulse and single imageacquisition separated by the predetermined time interval.


13.5.6



It is essential that the level of fluorescence signal at the end of the experiment be below15% of the prebleach level. The fluorescence loss curve should resemble the FLIP curvein Figure 13.5.2. If the loss of fluorescence is not below 15%, it is essential to increase themonitoring time. Note that the slope of the loss curve depends on the size, intensity, andduration of the bleach pulse.

Perform quantitative analysis of FLIP5. Measure the average intensity of the monitoring region of interest as a function of

time It.

6. Measure the average intensity of the total nuclear area of a neighbor cell at each timepoint (Tt).

7. Measure the background intensity in a random region outside of the cells at each timepoint (Bt).

8. For each time point, calculate the relative intensity:

Irel = (It − Bt)/[(I0 − B0) × (Tt − Bt)]

B T ROI

pre-bleach bleachimage

monitoring bleach

repeat

immobile fraction

Time

Log

(flu

ores

cenc

e in

tens

ity)

multiple fractions

single fraction

Figure 13.5.2 General scheme of a FLIP experiment. A cell is imaged before a short bleach pulseis applied to a defined bleach region. The image-bleach routine is repeated with the two stepsseparated by a predetermined time interval. To quantify and normalize FLIP data, measure theaverage fluorescence signal in a region of interest (ROI) distinct from the bleached region, measurethe average total nuclear fluorescence signal (T) in a neighboring cell, and the average signal in arandom region outside of the cell for background correction (B), for the duration of the experiment.When the FLIP data are plotted on a semilog scale, the shape of the curve indicates whether theGFP-fusion protein exists in the cell in a single fraction, in multiple fractions or whether it is immobile.


13.5.7

Organelle Motility

where I0 is the average intensity of the region of interest during prebleach.

This formula is derived in three steps. First, background is subtracted from all measuredvalues, then fluorescence loss due to monitor bleaching is normalized, and finally fluores-cence intensity is normalized to one.

9. Plot Irel as a function of time and determine average and standard deviations asdescribed for FRAP (see Basic Protocol, steps 16 and 17).

If all fluorescently tagged proteins in the compartment are kinetically identical, the regionof interest will lose its fluorescence linearly. If the nuclear compartment contains kineticallydistinct pools of proteins (e.g., due to binding of one fraction of the protein to chromatinor incorporation of a fraction into a large complex), the loss of fluorescence will bemultiphasic. The number of distinct fractions can be determined by plotting the loss curveas a function of time in a semilog plot. Each kink in the semilog plot corresponds to akinetically distinct fraction of molecules (Fig. 13.5.2).

As a negative control, fixed cells should be used. No significant loss of signal should beobserved outside of the bleached region. The amount of loss that is observed correspondsto bleaching due to imaging.

ALTERNATEPROTOCOL 2

INVERSE FLUORESCENCE RECOVERY AFTER PHOTOBLEACHING (iFRAP)

iFRAP is a hybrid method between FRAP and FLIP. In iFRAP, the entire nucleus excepta small region of interest is bleached using a single bleach pulse. The loss of fluorescencesignal from the unbleached region of interest is then monitored over time (Fig. 13.5.3).Because the number of unbleached molecules in the region of interest is small comparedto the number of bleached molecules outside the region of interest, the unbleachedmolecules do not contribute to new binding events because their pool is proportionally toosmall in comparison to the large pool of bleached molecules. Thus, iFRAP is ideally suitedto provide information about off-rates of proteins from binding sites (e.g., those on chromatin)or from nuclear compartments. iFRAP is particularly well suited for proteins which areenriched in relatively small structures (e.g., replication sites, intranuclear compartments) andfor proteins which are bound to a substrate for relatively long periods of time.

See Basic Protocol for materials.

1. Set up cells on the confocal microscope and image as described for FLIP (seeAlternate Protocol 1, steps 1 to 2). Adjust the microscope settings as described forFRAP (see Basic Protocol, steps 4 to 6).

2. Adjust laser power to 100% for bleaching.

The bleach time should be minimized and the efficiency of the bleach should be tested ona fixed sample. The average intensity in a bleached region in a fixed sample should bereduced by at least 70% compared to the same region of the neighboring cell. If the bleachdepth is insufficient, use several, rapid, consecutive bleach pulses.

3. Adjust the number of prebleach images for five to ten images.

4. Adjust the number of postbleach intervals and the time intervals.

The number of intervals is determined by the resolution needed and the length of intervalsshould cover the curve until it reaches a plateau. Be careful not to collect an unnecessarilyhigh number of images as this will bleach the sample.

5. Define the region to be bleached excluding the region of interest.

This must be performed quickly as the region of interest can move while the bleached regionis being outlined. The bleach has to be as fast as possible to obtain information aboutrapidly moving molecules in the unbleached region of interest.


13.5.8



6. Initiate data collection (i.e., prebleach images, bleach, postbleach images) andmonitor fluorescence loss in the region of interest. Check the position of the cell and thefocal plane to be sure that the region of interest is not moving from the original position.

The shape of the fluorescence loss curve should resemble the iFRAP curve in Figure 13.5.3.The region of interest has to be monitored until the loss of fluorescence signal reaches a plateau.

Perform quantitative analysis of iFRAP7. Measure the average intensity of the region of interest at each time point (It).

8. To correct for the photobleaching effect due to monitoring, measure the averageintensity of the entire nucleus of a neighboring cell at each time point (Tt).

9. Measure the background intensity in a randomly selected region outside of the cellat each time point (Bt).

10. Calculate the relative intensity (Irel) each time point:

( )− × −=

− × −0 0

0 0 )

( )

( ) (t t

relt t

I B T BI

I B T B

B T ROI

pre-bleach bleach monitoring

Flu

ores

cenc

e in

tens

ity

Time

single fraction multiple fractions

Time

Figure 13.5.3 General scheme of an iFRAP experiment. A cell is imaged before a short bleach pulseis applied to the whole area of the nucleus except a small region of interest (ROI). The loss of fluorescencesignal in the ROI is then monitored. To quantify and normalize iFRAP data, measure the averagefluorescence signal in the ROI, the average total nuclear fluorescence signal (T) in a neighboring cell,and the average signal in random place outside of the cell for background correction (B), for theduration of the experiment. If the GFP-fusion protein in the ROI exists as a single population, the iFRAPcurve should resemble the iFRAP curve on the left. If the GFP-fusion protein exists in the region ofinterest in multiple populations, the iFRAP curve should resemble the curve on right.


13.5.9

Organelle Motility

where I0 is the average intensity of the region of interest during prebleach (step 1)and T0 is the average intensity of the total nuclear area of a neighbor cell duringprebleach.

This formula is derived in three steps: first background is subtracted from all measuredvalues, then fluorescence loss due to monitor bleaching is normalized, and finally fluores-cence intensity is normalized to one.

11. Plot Irel as a function of time and determine average and standard deviations asdescribed for FRAP (see Basic Protocol).

It is advisable to plot all acquired curves before determining the average. Jumps in therecovery curves or declining curves are frequently caused by cell or focal plane movement.These curves can either be discarded for analysis or, if possible, they should be manuallyremeasured. For many applications it is sufficient to collect data for 10 to 20 individualcells. Typical error bars are on the order of 5% to 10% of the measured value.

If all GFP-fusion proteins in the compartment are fast moving components from the samepool, the fluorescent signal in the region of interest after the bleach will drop dramaticallyand then reach a plateau immediately (Fig. 13.5.3). If the nuclear compartment containskinetically distinct pools of proteins (due to binding of one fraction of the protein tochromatin or substrate for longer time or incorporation of a fraction into a large complex),the curve of loss of fluorescence in the region of interest will drop dramatically and thenshow a linear decline until it finally reaches a plateau (Fig. 13.5.3).

As a control, fixed cells should be used. In this case the loss of fluorescence in the regionof interest should show no significant loss of signal after the bleach over the monitoring.

SUPPORTPROTOCOL 1

TRANSIENT TRANSFECTION OF MAMMALIAN CELLS BYELECTROPORATION

To study the dynamics of nuclear proteins in living cells, the specific protein-GFP chimeraneeds to be expressed. Ideally, a cell line stably expressing the GFP fusion protein froman integrated transgene is generated. The use of stable cell lines and inducible promoterssuch as the tetracycline- or ecdysone-inducible expression systems (No et al., 1996) arerecommended when the overexpression of a fusion protein might be toxic or interferewith essential biological functions (Freundlieb et al., 1998). Detailed protocols forestablishing stable cell lines are presented elsewhere (e.g., Stenmark and Zerial, 1998).

A quicker and often sufficient option for expression of GFP-fusion proteins in living cellsis the introduction of a GFP-fusion protein by transient transfection encoded on a vectorwhich does not integrate into the cell’s genome. These vectors result in short-term,high-level expression of the fusion protein, but they will eventually be eliminated fromthe cell and expression will cease. Regardless of whether GFP fusion proteins are stablyor transiently expressed, a vector containing the GFP-fusion protein cDNA must beintroduced into cells by transfection. Two of the most effective and reliable methods oftransfection are transfer of DNA via electrical current (presented here) or through one ofthe commercially available lipid-mediated delivery protocols (see Support Protocol 2).These protocols are for transfection of adherent cells, but both methods can also be usedfor transfection of suspension cells.

This protocol has been tested for the following adherent cell lines: HeLa, COS, CMT3,NIH 3T3, BHK, and CHO cells. The protocol must be optimized for other cell types. Theimportant parameters for optimal results are the strength of applied electric field, durationof the electrical pulse, number of pulses applied, and concentrations of cells and plasmidDNA.


13.5.10



Materials

CellsComplete growth medium (e.g., UNIT 1.2)Plasmid DNACarrier DNA: sheared salmon sperm DNA1× PBS (APPENDIX 2A)0.25% (w/v) trypsin/1 mM EDTA, 37°CComplete growth medium with 25 mM HEPES, but without phenol red

10-cm Petri dish37°C warming tray15-ml test tubesElectroporation cuvettes with 2-mm gapBTX ECM830 electroporator (BTX, a division of Genetronics, Inc.) or equivalentCulture dish or incubation chamber (e.g., 2- or 4-chambered Lab-Tek II

coverglass; Nalgene Nunc)

Additional reagents and equipment for mammalian cell culture (UNIT 1.1)

NOTE: All solutions and materials coming into contact with cells must be sterile, andproper aseptic technique must be used.

NOTE: All cell culture incubations should be performed in a humidified 37°C, 5% CO2

incubator unless otherwise indicated.

NOTE: Avoid repeated freezing and defrosting of plasmid DNA stock solution. It maycause nicks in the plasmid DNA which leads to linearization of supercoiled DNA. Preparealiquots of plasmid DNA and store them at −20°C.

1. Prepare an ∼70% confluent monolayer of cells in a 10-cm Petri-dish (UNIT 1.1) incomplete growth medium.

2. Add 3 to 7 µg plasmid DNA to a 1.5-ml test tube. Add sufficient carrier DNA to bringthe total amount of DNA to 20 µg. Adjust the volume to 30 µl with water.

3. Wash cells once with 1× PBS.

4. Add enough 0.25% (w/v) trypsin/1 mM EDTA, 37°C to the culture to cover theadhering cell layer. Place the plate on a 37°C warming tray 1 to 2 min.

5. When cells round up, but before they detach from the substratum, add 7 ml completegrowth medium. Harvest by collecting the cells and transferring them to a 15-mlcentrifuge tube. Pellet cells by centrifuging 2 min at 1000 rpm in a benchtopcentrifuge, room temperature.

6. Decant the aqueous layer and resuspend cells in 200 µl fresh complete medium. Pipetthe cell suspension into the 1.5-ml test tube containing the prepared DNA solution(step 2). Mix well by pipetting up and down three times, let stand 2 min, and transferto an electroporation cuvette with 2-mm gap.

7. For HeLa, COS, CMT3, NIH 3T3, BHK, or CHO cells, electroporate on a BTXECM830 electroporator using the following settings: 150 V, 1-msec pulse, 4 pulses,and 0.5-sec interval.

For BTX electroporator settings for other cell types refer to http://www.btxonline.com/btxand the manufacturer’s lab manual at http://www.btxonline.com/products/pdfs/ECM_830/ECM_830_Manual.pdf. For electroporators from other manufacturers seethe appropriate company’s web pages.


13.5.11

Organelle Motility

8. Using a Pasteur pipet, transfer the cells from the cuvette to a well in a culture dish orto an incubation chamber.

For HeLa, COS, CMT3, NIH 3T3, BHK, or CHO cells, one drop of cell suspension in a35-mm well gives a confluent layer after overnight incubation. For other cell types, theamount of cells must be optimized.

9. Change the medium 8 to 10 hr after transfection.

10. Before imaging living cells (see Basic Protocol and Alternate Protocols 1 and 2), replacethe medium with complete medium containing 25 mM HEPES, but no phenol red.

Phenol red increases background fluorescence.

SUPPORTPROTOCOL 2

TRANSIENT TRANSFECTION OF MAMMALIAN CELLS USING FuGENE 6

FuGENE 6 is a transfection reagent that highly efficiently transfects a wide variety ofcells including primary cultures and hard-to-transfect cell lines. The advantage ofFuGENE 6 is that it demonstrates virtually no cytotoxicity even with primary cell cultures.The authors have successfully tested several adherent transformed cell lines—HeLa,COS, CMT3, NIH 3T3, CHO—and mouse primary fibroblasts. The FuGENE 6 reagentweb page—http://biochem.roche.com/techserv/fugene.htm—has a current list of 250successfully transfected cell lines.

Materials

CellsSerum-containing and serum-free medium (UNIT 1.2)FuGENE 6 reagent (Roche)Plasmid DNA solutionSerum-containing medium (UNIT 1.2)Serum (optional)Glass coverslips or Lab Tek II incubation chambers (Nalgene Nunc)

Additional reagents and equipment for mammalian cell culture (UNIT 1.1)

NOTE: All solutions and materials coming into contact with cells must be sterile, andproper aseptic technique must be used.

NOTE: All cell culture incubations should be performed in a humidified 37°C, 5% CO2

incubator unless otherwise indicated.

NOTE: Avoid repeated freezing and defrosting of plasmid DNA stock solution. It maycause nicks in the plasmid DNA which leads to linearization of supercoiled DNA. Preparealiquots of plasmid DNA and store them at −20°C.

1. Prepare ∼50% to 70% confluent monolayers on glass coverslips or in Nalgene LabTek II incubation chambers (UNIT 1.1) in serum-containing medium.

2. In a 1.5-ml sterile test tube, add 97 µl serum-free medium, then add 3 µl FuGENE6reagent directly into this medium.

The order of addition is critical. The serum-free medium must be aliquoted into the testtube first to avoid adversely affecting transfection efficiency by contact of the undilutedFuGENE with plastic surfaces.

3. Add 1 to 2 µg plasmid DNA solution in a volume of 0.5 to 50 µl to the diluted FuGENE6 reagent.

The total volume of DNA solution has to be in this range.


13.5.12



4. Gently tap the test tube to mix the contents, but do not vortex. Incubate a minimumof 15 min at room temperature.

Continued incubation for up to 45 min will not affeect the transfection efficiency in mostcell types.

5. Dissolve the mixture into 2 ml serum-containing medium. Add to the cells dropwise,distributing it around the well or incubation chamber. Swirl the wells or incubationchamber to ensure even dispersal.

6. Return the cells to the incubator. After 3 to 8 hr, replace the medium with serum-con-taining medium or add serum directly to wells. Continue incubating (12 to 20 hr)until imaging is performed (see Basic Protocol and Alternate Protocols 1 and 2).

SUPPORTPROTOCOL 3

QUANTIFICATION OF FLUORESCENTLY TAGGED PROTEINMOLECULES IN NUCLEAR COMPARTMENTS OF SINGLE LIVING CELLS

The number of fluorescently tagged molecules in nuclear compartments in single livingcells can be determined using individual fluorescently labeled virus-like particles (VLP)as a standard for the observed signals. VLPs contain a precise number of GFP moleculesand the number of GFP molecules in a test volume can therefore be determined bycomparing the total fluorescent intensity of VLP and the total fluorescent intensity of anuclear volume. Since VLPs are small spherical particles, this technique is especiallyuseful for the determination of the GFP content of small spherical cellular compartmentsand organelles such as Cajal bodies, PML bodies, fibrillar centers within the nucleoli,mitochondria, or vesicles.

VLPs are in vitro–assembled rotavirus particles (Charpilienne et al., 2001). Rotavirusesare large icosahedral particles, which contain three concentric capsid layers. While theoutermost layer is composed of the VP4 and VP7 proteins, the intermediate capsid layeris composed of trimers of VP6. The innermost layer contains exactly 120 molecules ofVP2. When VP2 fused to GFP is coexpressed with the intermediate capsid protein VP6in a baculovirus-insect cell system, double-layered icosahedral VLP are completelyassembled. Each such VLP measures ∼100 nm in diameter and contains exactly 120VP2-GFP molecules (Charpilienne et al., 2001).

Materials

Cells expressing GFP-tagged protein of interest (see Support Protocols 1 and 2)Complete growth medium (UNIT 1.2)1× PBS (APPENDIX 2A)GFP-labeled VLP particles (Charpilienne et al., 2001; Dundr et al., 2002)

22 × 22–mm glass coverslipsConfocal microscope with 100× objective

Additional reagents and materials for mammalian cell culture (UNIT 1.1)

1. Culture cells expressing the GFP-tagged protein of interest on 22 × 22–mm glasscoverslips for 16 to 20 hr (UNIT 1.1) in complete growth medium.

2. Wash cells twice with 1× PBS for 5 min each time.

3. Dilute purified GFP-labeled VLP particles to a concentration of 1 to 10 µg/ml in PBS.

4. Place ∼30 to 40 µl diluted VLP onto the coverslip (step 1).

It is advisable to perform a pilot experiment in which VLP particles are imaged in solutionwithout cells. This allows for their easy detection later when they are mixed with the cells.


13.5.13

Organelle Motility

5. Examine cells using a confocal microscope with 100× objective.

The powerful objective is needed because VLPs are difficult to see at low magnifications.

Alternatively, a conventional wide-field microscope fitted with a cooled CCD camera canbe used.

6. Adjust the photomultiplier tube intensity such that the fluorescence signal of the GFPfusion protein is slightly below the saturation level.

Use at least 8- but preferably 12-bit imaging. It is important to ensure that no pixels aresaturated since the fluorescence intensity in these pixels cannot be measured accurately.

7. Collect a series of 50 single images of the cells expressing the GFP-fusion protein.

8. Using identical settings, collect 50 single images of the VLP-GFP particles in adifferent field of view in the same specimen.

9. Measure the average area and the average intensity of at least fifty compartments ofinterest containing the GFP-fusion protein and at least fifty VLPs.

10. Measure the background intensity in a randomly selected area that does not containcells or VLPs.

11. Subtract the background intensity from the average intensity value of compartmentsand VLPs.

12. Multiply the average area with the background-corrected average intensity to obtainaverage fluorescence intensities for the compartment and VLPs.

13. Calculate the number of GFP molecules per compartment:

no. GFP molecules/compartment = (TIC × 120)/(TIVLP)

where TIC is the average intensity of the compartments and TIVLP is the averageintensity of the VLPs.

When using a confocal microscope, not all of the imaged VLP particles will fall within thefocal plane and a range of signals will be observed. To determine the average intensity ofa single VLP particle, it is helpful to perform three-dimensional reconstruction of VLPswith a distance of 100 nm between optical sections. Collect the maximum number ofsections possible and observe them for quantification as a maximum projection. In thiscase, the total intensities of VLPs should fall within a 10% range of intensities.

COMMENTARY

Background InformationIt has been established that the eukaryotic

cell nucleus is not a homogeneous organelle butcontains several specific membrane-less com-partments. Recent observations using pho-tobleaching methods have demonstrated thatmany nuclear proteins are highly mobile anddo not permanently reside in any specific nu-clear compartment but are continuously ex-changed between the compartment and the sur-rounding nucleoplasm (Kruhlak et al., 2000;Phair and Misteli, 2000; Chen and Huang,2001). In this view, nuclear compartments arethe consequence of the steady-state dynamicbehavior of their components. Nuclear proteins

are not primarily targeted to the nuclear com-partments by specific targeting signals butrather retained there as a result of binding orcollision with other nuclear components (i.e.,protein-protein interactions; DNA-protein in-teractions: engagement in replication, DNA re-pair or RNA-protein interactions, and engage-ment in transcription etc.). The exchange rateof the nuclear proteins is strongly determinedby the roles they play in nuclear function.

The localization of proteins within cells hastraditionally been studied by indirect im-munofluorescence microscopy using fluores-cently labeled antibodies. This method is lim-ited by the requirement for chemical fixation


13.5.14



of cells and yields only static snapshots of aprotein’s distribution. The development of thegreen fluorescent protein (GFP) as a geneticallyencoded fluorescent protein reporter has revo-lutionized the study of protein localization inliving cells.

Apart from the use of GFP as a convenientand rapid method to determine where a knownprotein or one encoded by a newly clonedcDNA is localized, GFP has also been exten-sively used to visualize proteins in living cellsby time-lapse microscopy. These latter ap-proaches allow the study of dynamic aspects ofprotein function. GFP fusion proteins havebeen particularly insightful in the study of thecell nucleus. Using time-lapse-microscopy ap-proaches, it has become clear that many nuclearcompartments are highly dynamic both in in-terphase and during mitosis, and that manyproteins continuously shuttle between the nu-cleus and the cytoplasm. (Ellenberg et al., 1997;Misteli et al., 1997; Boudonck et al., 1999;Dundr et al., 2000; Platani et al., 2000;Tsukamoto et al., 2000; Kamath et al., 2001).More recently, nuclear GFP fusion proteinshave also been used in combination with pho-tobleaching techniques to study the dynamicorganization of nuclear compartments in vivo.

Fluorescence photobleaching techniqueswere originally developed in the 1970s to studythe mobility of lipids and proteins in the lipidbilayer of the plasma membrane (Axelrod et al.,1976). They have now become a standardmethod for studying the dynamics of nuclearproteins in living cells (Ellenberg et al., 1997;Houtsmuller et al., 1999; Kruhlak et al., 2000;Phair and Misteli, 2000; Snaar et al., 2000;Misteli, 2001; Chen and Huang, 2001; Boisvertet al., 2001). The noninvasive nature of thephotobleaching approach allows one to labelnuclear compartments in living cells with highspecificity, but often without functional inter-ference. The GFP-protein marker is particu-larly useful for photobleaching experiments,because the signal is stable and does not bleachsignificantly at the low-intensity levels used tomonitor bleach recovery. In addition, thebleaching of GFP in living cells can be consid-ered irreversible and does not cause any detect-able damage to the cell (but see Verkman,2002).

What are the advantages of using pho-tobleaching techniques for studying the dy-namics of nuclear proteins in living cells? Re-cent observations using photobleaching tech-niques have demonstrated that many nuclearproteins are highly mobile and do not perma-

nently reside in any specific domain of thenucleus (Kruhlak et al., 2000; Phair and Misteli,2000; Chen and Huang, 2001). The variousavailable photobleaching techniques can pro-vide information about the in vivo kinetics ofthe protein of interest and since the kineticbehavior of a protein is often directly related toits functional status, information about a pro-tein’s biological role may also be obtained(Phair and Misteli, 2001). Furthermore, pho-tobleaching methods can be used to determinehow many different pools of a protein are pre-sent in a cell and whether an immobile staticallybound fraction exists.

Critical Parameters

Transfection efficiencyTransfection efficiency plays an important

role in photobleaching experiments. Very lownumbers of transfected cells can affect theevaluation of the pattern of GFP-fusion proteinlocalization and selection of optimal cells forbleaching. In the case of FLIP or iFRAP ex-periments, high transfection efficiency is essen-tial so that images of two neighboring cellsexpressing the GFP protein can be obtained,which is required for correction of the signaldue to photobleaching. Therefore special atten-tion should be paid to optimization of transfec-tion efficiency.

Cell typeA cell type that tolerates introduction of

plasmid DNA should be selected. If there is anoption, select preferentially transformed cellsover primary cells, which are usually moredifficult to optimize for transfection. Cellsshould be in logarithmic growth when used fortransfection.

Quality of plasmid DNAOne critical parameter of good transfection

efficiency is the quality of plasmid DNA. Itshould be pure and the stock DNA solutionshould not be repeatedly frozen and defrosted.When transfection efficiency is low, a higherconcentration of plasmid DNA can be used.

ElectroporationWhen electroporation is used, three parame-

ters should be varied for optimization: electri-cal field, duration, and number of pulses. Whenthe electrical field is too high, the survival rateof the cells is lowered. When it is too low, theefficiency of transfection is usually poor. When


13.5.15

Organelle Motility

a low electric field is used, longer pulses shouldbe applied and vice versa.

Expression levelsFor photobleaching experiments, it is advis-

able to select only cells expressing low or mod-erate levels of GFP fusion protein. Overexpres-sion of a protein of interest can dramaticallyaffect its localization and may change its be-havior as well as influence the overall metabo-lism of the cell.

Critical elementsSpecial attention must be paid to the viabil-

ity of the cells during photobleaching measure-ments. Cells in a chambered coverglass shouldnot be on the microscope stage for more than 1hr. Longer exposure of cells can affect cellmetabolism and can cause changes in proteinmobility. After 1 hr cells should be replacedwith another sample or put back in the incubatorfor at least 30 min.

ControlsFixed cells should be used as a control for

all photobleaching experiments. The fixed sam-ple permits determination of the x, y, and zdimensions of the bleach region and depth ofbleach, if the series of optical sections throughthe sample is collected. The size of the bleachspot should be small relative to the size of thenucleus. Typically a bleach spot 1 µm in diame-ter works for most applications. Bleaching aregion that is larger than the compartment ofinterest will affect the accuracy of measurement

by mixing the populations of GFP-moleculesoutside and inside of the compartment. There-fore, it is advisable to adjust the size of com-partment relative to the size of the bleach spotby using the appropriate zoom.

Each individual recovery curve should beevaluated. Any dramatic change in the shape ofthe curve may indicate that the monitoringregion of interest changed position or focalplane due to cell movement. These data shouldbe discarded.

Anticipated ResultsIn a typical transient transfection, a fluores-

cent signal should be detected 12 to 20 hr aftertransfection, but weak signals can often beobserved as early as 4 to 6 hr. The differencesin expression level can vary depending on thepurity of DNA, the vector and promoter used,and the cell type used in the study.

The recovery curve of the GFP fusion pro-tein of interest contains information about thedegree of mobility of a protein of interest withinthe nucleus. Generally, qualitative analysis ofany photobleaching curves allows one to con-clude whether a protein is completely mobile,mobile with a fraction of immobile molecules,or almost completely immobile. In many casesthis basic information has significant ramifica-tions for protein function. Note that coinci-dence of recovery curves for multiple proteinsdoes not necessarily mean that these proteinsare found in a complex and move together.Conversely, however, distinct curves of severalproteins are generally a good indication that the

Table 13.5.2 Comparison of the Mobility of Nuclear Proteins

GFP-fusion protein FRAP recovery(sec)

Diffusioncoefficient (µm2/s)

Reference

EGFP ∼0.5 27 Swaminathan et al., 1997580 kDa dextran 7 0.95 Calapez et al., 2002GFP-ASF/SF2 20 0.24 Phair and Misteli, 2000GFP-HMG17 30 0.45 Phair and Misteli, 2000GFP-PABP2 7 0.6 Calapez et al., 2002GFP-TAP 3 1.2 Calapez et al., 2002Fibrillarin-GFP 30 0.53 Phair and Misteli, 2000GFP-UBF1 60 0.14 (nucleolus)

0.57 (nucleoplasm)Chen and Huang, 2001

GFP-Nucleolin 60 0.14 (nucleolus)1.15 (nucleoplasm)

Chen and Huang, 2001

TFIIH-GFP 30 5.1 Hoogstraten et al., 2002ERCC1-GFP/XPF 8

—

15 (absence ofDNA damage)(immobile withDNA damage)

Houtsmuller et al., 1999


13.5.16



majority of the proteins of interest are notpresent in a complex in vivo.

Examples of recovery times and diffusioncoefficients are given in Table 13.5.2.

As a positive control for the evaluation ofthe mobility of GFP protein of interest, GFPalone can be used as a standard for relativelyfreely mobile protein. The recovery of the fluo-rescent signal of the GFP alone is very fast(within 1 sec) and complete recovery with vir-tually no immobile fraction should be ob-served. Convenient negative controls are GFPfusions with the core histones (H2A-GFP, H3-GFP, H4-GFP; Kimura and Cook, 2001). In thiscase, there should be no recovery over themonitoring period. If recovery of fluorescenceis observed using either fixed cells or corehistone fusion proteins, it should be ensuredthat the correct excitation wavelength of thelaser (488 nm for GFP) is used. Similarly, if nobleaching is observed, the laser settings shouldbe checked.

In FLIP experiments, the kinetic status ofspecific nuclear compartments is evaluated.Specifically it is tested whether a GFP-fusionprotein resides statically or moves rapidly inand out of a compartment. If two or morecompartments are in physical continuity,bleaching one will result in loss of fluorescencein the other(s).

In iFRAP experiments, loss of fluorescence inregion of interest reflects predominantly how theGFP-fusion protein is lost from its binding site.

Most photobleaching experiments shouldbe completely reproducible with a typical erroramong cells in a population on the order of 5%to 10% of measured values. To achieve this goal,each acquired curve has to be evaluated inde-pendently, and every curve with jumps due to cellor focal plane movement should be discardedbefore final averaging. Since these are single-cellexperiments, variations in cell populations mightbe observed as relatively large errors in a popula-tion measurement. For many proteins the positionof the cell in the cell cycle can affect their behaviorand mobility. If large fluctuation in measurementsis observed, it is recommended to use cell-cycle-synchronized cells.

Special care must be applied to minimizephotobleaching due to imaging during moni-toring. The number of images collected duringmonitoring should be a compromise betweenthe resolution of measurements needed and thelength of the interval. The decline of the signalintensity due to photobleaching during moni-toring should generally not exceed ∼5% to 10%of the prebleach value. If bleaching is a prob-

lem, the laser power or the interval betweenimages can be reduced.

GFP levels can be quantified with VLPs bycomparing total fluorescent intensities of VLPswith a defined number of GFP molecules andfluorescently labeled specific small nuclearcompartments of interest (see Support Protocol3). This measurement provides the averagenumber of GFP-fusion protein molecules in thecompartments in single living cells. Since in12-bit imaging, 4096 gray levels can be distin-guished and the typical background intensity is∼100 to 200 units, the observable ratio of VLPto compartment signal is ∼20-fold. Given thateach virus particle contains 120 GFP mole-cules, only compartments which contain ∼2500molecules of interest can be measured usingthis method. The level of GFP molecules foundin many nuclear compartments is well withinthis range.

Time ConsiderationsA transient transfection can be done in 30 to

45 min and the cells can be used for microscopy12 to 20 hr after transfection. The time for aFRAP and iFRAD experiment depends on themobility of the GFP fusion-protein, but is typi-cally on the order of seconds to <10 min. AFLIP experiment takes on the order of minutesup to 1 hr. Each GFP fusion protein should betested on 15 to 20 cells. The normalization ofone sheet of FRAP data using Microsoft Exceltakes several minutes. Electroporation takes∼20 min and the transfection procedure usingFuGENE 6 takes 25 min to 1 hr. Acquiring asufficient number of images of cells expressingprotein-GFP and VLPs requires 0.5 to 1.0 hr.Measuring the average area and the averageintensity of at least fifty compartments of inter-est and fifty VLPs using microscope or Meta-morph software requires ∼2 to 3 hr.

Literature CitedAxelrod, D., Koppel, D.E., Schlessinger, J., Elson,

E., and Webb, W.W. 1976. Mobility measure-ment by analysis of fluorescence photobleachingrecovery kinetics. Biophys. J. 16:1055-1069.

Boisvert, F.M., Kruhlak, M.J., Box, A.K., Hendzel,M.J., and Bazett-Jones, D.P. 2001. The transcrip-tion coactivator CBP is a dynamic component ofthe promyelocytic leukemia nuclear body. J. CellBiol. 152:1099-1106.

Boudonck, K., Dolan, L., and Shaw, P.J. 1999. Themovement of coiled bodies visualized in livingplant cells by the green fluorescent protein. Mol.Biol. Cell 10:2297-2307.

Calapez, A., Pereira, H.M., Calado, A., Braga, J.,Rino, J., Carvalho, C., Tavanez, J.P., Wahle, E.,


13.5.17

Organelle Motility

Rosa, A.C., and Carmo-Fonseca, M. 2002. Theintranuclear mobility of messenger RNA bind-ing proteins is ATP dependent and temperaturesensitive. J. Cell Biol. 159: 795-805.

Charpilienne, A., Nejmeddine, M., Berois, M.,Parez, N., Neumann, E., Hewat, E., Trugban, G.,and Cohen, J. 2001. Individual rotavirus-likeparticles containing 120 molecules of fluores-cent protein are visible in living cells. J. Biol.Chem. 276:29361-29367.

Chen, D., and Huang, S. 2001. Nucleolar compo-nents involved in ribosome biogenesis cycle be-tween the nucleolus and nucleoplasm in inter-phase cells. J. Cell Biol. 153:169-176.

Dundr, M., Misteli, T., and Olson, M.O.J. 2000. Thedynamics of postmitotic reassembly of the nu-cleolus. J. Cell Biol. 150:433-446.

Dundr, M., McNally, J.G., Cohen, J., and Misteli, T.2002. Quantification of GFP fusion proteins insingle living cells. J. Struct. Biol. 140:92-99.

Ellenberg, J., Siggia, E.D., Moreira, J.E., Smith,C.L., Presley, J.F., Worman, H.J., and Lippincott-Schwartz, J. 1997. Nuclear membrane dynamicsand reassembly in living cells: Targeting of aninner nuclear membrane protein in interphaseand mitosis. J. Cell Biol. 138:1193-1206.

Ellenberg, J., Chazotte, B., and Lippincott-Schwartz, J. 1998. Fluorescence photobleachingtechniques. In Cells, A Laboratory Manual.(D.L. Spector, R.D. Goldman, L.A. Leinwand,eds). pp. 79.1-79.23. Cold Spring Harbor Press,Cold Spring Harbor, N.Y.

Freundlieb, S., Baron, U., and Bujard, H. 1998.Controlling gene activities via the tetracyclineregulatory systems. In Cell Biology: A Labora-tory Handbook, 2nd Edition, Vol. 4 (J.E. Celis,ed.) pp. 230-238. Academic Press, San Diego.

Hoogstraten, D., Nigg, A.L., Heath, H., Mullenders,L.H., van Driel, R., Hoeijmakers, J.H., Ver-meulen, W., and Houtsmuller, A.B. 2002. Rapidswitching of TFIIH between RNA polymerase Iand II transcription and DNA repair in vivo. Mol.Cell. 10: 1163-1174.

Houtsmuller, A.B., Rademakers, S., Nigg, A.L.,Hoogstraten, D., Hoeijmakers, J.H.J., and WimVermeulen, W. 1999. Action of DNA repair en-donuclease ERCC1/XPF in living cells. Science284:958-961.

Houtsmuller, A.B. and Vermeulen, W. 2001. Macro-molecular dynamics in living cell nuclei revealedby fluorescence redistribution after photobleach-ing. Histochem. Cell Biol. 115:13-21.

Kamath, R.V., Leary, D.J., and Huang S. 2001.Nucleocytoplasmic shuttling of polypyrimidinetract-binding protein is uncoupled from RNAexport. Mol. Biol. Cell 12: 3808-3820

Kimura, H. and Cook, P.R. 2001. Kinetics of corehistones in living human cells: Little exchangeof H3 and H4 and some rapid exchange of H2B.J. Cell Biol. 153:1341-1353.

Kruhlak, M.J., Lever, M.A., Fischle, W., Verdin, E.,Bazett-Jones, D.P., and Hendzel, M.J. 2000. Re-

duced mobility of the alternative splicing factor(ASF) through the nucleoplasm and steady statespeckle compartments. J. Cell Biol. 150:41-51.

Lippincott-Schwartz, J., Snapp, E., and Kenworthy,A. 2001. Studying protein dynamics in livingcells. Nature Rev. Mol. Cell Biol. 2:444-456.

McNally, J.G. and Smith, C.L. 2002. Photobleach-ing by confocal microscopy. In Confocal andTwo-Photon Microscopy: Foundations, Applica-tions, and Advances, (A. Diaspro, ed.) pp. 525-538. Wiley-Liss, New York.

Misteli, T. 2001. Protein dynamics: Implications fornuclear architecture and gene expression. Sci-ence 291:843-847.

Misteli, T., Caceres, J.F., and Spector, D.L. 1997.The dynamics of a pre-mRNA splicing factor inliving cells. Nature 387:523-527.

No, D., Yao, T.-P., and Evans, R.M. 1996. Ecdysone-inducible gene expression in mammalian cellsand transgenic mice. Proc. Natl. Acad. Sci.U.S.A. 93:3346-3351.

Phair, R.D. and Misteli, T. 2000. High mobility ofproteins in the mammalian cell nucleus. Nature404:604-609.

Phair, R.D. and Misteli, T. 2001. Kinetic modellingapproaches to in vivo imaging. Nature Rev. Mol.Cell Biol. 2:1-10.

Platani, M., Goldberg, I., Swedlow, J.R., andLamond, A.I. 2000. In vivo analysis of Cajalbody movement, separation, and joining in livehuman cells. J. Cell Biol. 151:1561-1574.

Reits, E.A. and Neefjes, J.J. 2001. From fixed toFRAP: Measuring protein mobility and activityin living cells. Nature Cell Biol. 3:E145-E147.

Snaar S., Wiesmeijer K., Jochemsen A.G., TankeH.J., and Dirks R.W. 2000. Mutational analysisof fibrillarin and its mobility in living humancells. J. Cell Biol. 151:653-662.

Stenmark, H. and Zerial M. 1998. Lipofection. InCell Biology: A Laboratory Handbook, 2nd Edi-tion, Vol. 4 (J.E. Celis, ed.) pp. 141-144. Aca-demic Press, San Diego.

Swaminathan, R., Hoang, C.P., and Verkman, A.S..(1997) Photobleaching recovery and anisotropydecay of green fluorescent protein GFP-S65T insolution and cells: Cytoplasmic viscosity probedby green fluorescent protein translational androtational diffusion. Biophys. J. 72: 1900-1907.

Tsukamoto, T., Hashiguchi, N., Janicki, SM, Tum-bar, T., Belmont, A.S., and Spector, D.L. 2000.Visualization of gene activity in living cells. Nat.Cell Biol. 2:871-878.

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Contributed by Miroslav Dundr and Tom MisteliNational Cancer InstituteBethesda, Maryland


13.5.18



UNIT 13.6Functional Characterization of ProteinsRegulating Actin Assembly

The actin cytoskeleton is a dynamic array of polar helical filaments that allows a eukaryoticcell to organize its intracellular space and the traffic of organelles, as well as to react tosignals from the outside world by changing shape or migrating. Directed assembly ofactin filaments is at the heart of motile processes involved in cell migration, oogenesis,tissue morphogenesis and repair, embryonic development, and the immune response.

A very large repertoire of actin-binding proteins regulates the assembly dynamics andthe spatial organization of actin filaments, thus orchestrating the motile behavior of thecell (Small et al., 2002). At least 50 to 60 regulatory proteins have been identified so far,yet regulatory factors endowed with new specific properties continue to be discovered.The goal of this unit is to describe a logical method and derived in vitro assays for thecharacterization of the function of a putative actin-binding protein. The classical basicregulatory activities are presented here as dogma, or basic themes, on which variationsare played by nature to generate the full complexity of the actin response in physiologicalsituations.

In the physiological context, actin is in a dynamic steady state between the monomeric (G,for globular; see Support Protocol 1) and polymeric (F, for filamentous) forms. A filamenthas two ends that differ both structurally and dynamically: a barbed end (the fast end) anda pointed end (the slow end). Actin is an ATPase: it binds MgATP, and hydrolysis of ATPis associated with incorporation of ATP-G-actin in the filament. Actin assembly at steadystate is described by an ATPase cycle featuring the energetic imbalance between the twoends, which is linked to the irreversible hydrolysis of ATP. This cycle comprises threeelementary steps: net depolymerization from the pointed end releases ADP-G-actin, ex-change of ATP for bound ADP restores ATP-G-actin, which then undergoes net assemblyat the barbed end (Fig. 13.6.1). The rate constants of all these steps determine both theconcentrations of each species at steady state and the turnover of actin filaments. It is note-worthy that the concentration of unpolymerized ATP-G-actin that is maintained in thiscycle is intermediate between the critical concentrations at the barbed and pointed ends.

Individual actin regulatory proteins affect the kinetic and thermodynamic parameters ofactin assembly at one or the other end of the filament in various fashions. Actin exposesmany functionally relevant sites for regulating assembly: side binding or end binding to theF-actin filament and binding to G-actin in ways that allow, favor, or inhibit incorporationof actin in filaments or that favor or inhibit nucleation of filaments.

A primordial question is whether a given actin-binding protein binds with a higher speci-ficity to G-actin or F-actin. A protein that binds F-actin better than G-actin stabilizesthe filaments (i.e., shifts the monomer-polymer equilibrium toward the polymer form),thus causing a decrease in G-actin concentration at steady state. Conversely, a proteinthat binds G-actin preferentially, also called a G-actin-sequestering factor, shifts the equi-librium toward the monomer, thus causing depolymerization of F-actin. Sedimentationassays (see Basic Protocol 1), which allow separation of F-actin (in the pellet) fromfree and liganded G-actin (in the supernatant) are useful to address this issue and mayprovide quantitative information on the protein’s affinity for and molar binding ratio toactin. Changes in fluorescence of pyrenyl-labeled actin or other fluorescently labeledforms of actin (see Support Protocols 2 and 3) also provide accurate measurements ofthe concentration of F-actin and G-actin at the steady state of assembly.

Contributed by Maud Hertzog and Marie-France CarlierCurrent Protocols in Cell Biology (2005) 13.6.1-13.6.23Copyright C© 2005 by John Wiley & Sons, Inc.

OrganelleMotility

13.6.1

Supplement 26

ProteinsRegulating Actin

Assembly

13.6.2


Figure 13.6.1 Treadmilling of the actin filament. This diagram emphasizes that the actin filamenthas a polar structure, with a barbed end and a pointed end. Under physiological ionic conditions,F-actin is at a steady state with ATP-G-actin monomers. An energetic imbalance occurs betweenthe two ends, because of ATP hydrolysis linked to F-actin assembly. Monomer-polymer exchangesat the two ends do not sum up to zero at each end; instead the barbed ends undergo net growth,balanced by equal depolymerization at the pointed end. This flux of subunits through the filamentis called treadmilling. Css, steady-state (critical) concentration; D, ADP-actin; DPi, ADP-Pi-actin; T,ATP-actin.

Because ATP hydrolysis accompanies filament assembly, a relevant question is whetheran actin-binding protein shows specificity for ATP-actin or ADP-actin (in either the G orF form). Direct binding assays (solid phase or fluorometric assays; see Basic Protocols2 and 3) are recommended. The change in tryptophan fluorescence (actin has four tryp-tophans) may be used as a probe of the binding of a protein to ATP- or ADP-G-actin.Alternatively, actin can be labeled on specific residues (e.g., Cys374) with a variety offluorophores that may react to ligand binding, thus providing information on bindingaffinity and the mechanism of interaction through the use of rapid kinetics. Labelingmay alter the affinity of the protein for actin, but this can easily be checked by compe-tition studies with unlabeled actin. Similarly, the interaction with an actin filament canbe studied in the standard ADP-F-actin state and also using the nonhydrolyzable analogsBeF3

− and AlF4− in the ATP- or ADP-Pi-F-actin state (Combeau and Carlier, 1988,

1989).

Regulation of filament turnover is crucial in motility and is elicited by changing thekinetic parameters for actin association-dissociation at the two ends of the filament. Alarge class of actin regulatory proteins, called capping proteins, binds tightly to the barbedend of the actin filament, thus blocking all association-dissociation processes at this end.Appropriate kinetic measurements of filament growth or depolymerization at one endor the other using the change in fluorescence of pyrenyl-labeled actin is described totest the capping activity (see Basic Protocols 5 and 6). Blockage of barbed ends alsoresults in a shift in the steady state critical concentration (Css) for assembly, increasingit up to the critical concentration at the pointed end (Cc) (see Basic Protocol 4). Mostcapping proteins simply block barbed ends, but leaky cappers that alter the associationor dissociation of actin without blocking the end may exist.

OrganelleMotility

13.6.3


Another way to affect the rate of filament elongation is provided by proteins that interactwith G-actin to form a complex that has an association rate constant for filament ends thatdiffers from that of free G-actin. Appropriate assays of filament growth from seeds helpto evaluate this possibility. Seeds are preformed filaments with a few subunits, which areused to initiate actin elongation from the barbed or pointed end of the actin filament.

The turnover of a population of filaments in vitro is monitored using a fluorescent analogof ATP whose fluorescence changes when bound to actin. The rate at which F-actinbecomes nonfluorescent following an ATP chase is an indication of the filament turnoverrate (see Basic Protocol 7).

BASICPROTOCOL 1

CO-SEDIMENTATION ASSAY FOR MEASURING BINDING OF A PROTEINTO F-ACTIN (OR G-ACTIN)

F-actin is easily sedimented by centrifuging 20 min at 400,000 × g (20◦C). G-actin andsmall complexes of G-actin with a G-actin-binding protein remain in the supernatant.

Materials

50 µM G-actin in G-buffer (see Support Protocol 1), store on ice2 M KCl20 mM MgCl2 (APPENDIX 2A)100 µM protein of interest in 10 mM Tris·Cl (pH 7.5)/1 mM dithiothreitol (DTT)10 mM Tris·Cl (pH 7.5)/1 mM DTTF-buffer (see recipe)G-buffer (see recipe)

0.5-ml polycarbonate centrifuge tubesBeckman TL100 tabletop ultracentrifuge (or equivalent)Densitometer

Additional reagents and equipment for SDS-polyacrylamide gel electrophoresis(UNIT 6.1)

1. To 1 ml of 50 µM G-actin, add 100 µl of 2 M KCl (0.2 M final) and 100 µl of 20mM MgCl2 (2 mM final). Allow to polymerize 15 min at room temperature.

2. Dilute 360 µl polymerized G-actin (i.e., F-actin) to 1.5 ml with F-buffer. Mix thor-oughly and prepare ten samples of 100 µl F-actin (10 µM final) in 0.5-ml polycar-bonate centrifuge tubes.

3. Centrifuge 200 µl of 100 µM protein of interest at 400,000 × g, 20◦C, for 20min. Prepare ten dilutions of the protein of interest ranging from 0 to 100 µM finalconcentration in a final volume of 100 µl using 10 mM Tris·Cl/1 mM DTT.

4. Add 100 µl protein of interest dilutions (0 to 50 µM final) to each of ten 100-µlF-actin samples (step 2; 5 µM final) and incubate 15 min at room temperature.Divide each sample between two tubes (100 µl each). Process one sample from eachpair immediately (step 5). Incubate the other sample from each pair for 18 hr at roomtemperature before processing.

With the longer incubation, one can monitor possible shifts in the steady state of actinassembly (changes in CSS) that are induced by the protein.

5. Centrifuge 20 min at 400,000 × g, 20◦C, using a tabletop ultracentrifuge. Removeand save supernatant, wash pellets and tube walls three times with 150 µl F-buffer,and resuspend pellets in 150 µl G-buffer.


Assembly

13.6.4


6. Perform SDS-PAGE of the pellets and supernatants on a 12% gel to visualize full-sizeactin. Run samples in parallel with appropriate molecular mass standards for actinand the protein of interest (15,000 to 100,000 Da). Identify actin by its molecularweight (42 kDa).

Under denaturing conditions, F- and G-actin migrate at the same level, but F-actin willbe found in the pellet and G-actin will be in the supernatant.

7. Scan the gel and use a densitometer to evaluate the amount of protein bound toF-actin.

If the protein binds to G-actin, it should cause depolymerization; the amount of G-actinwill be greater in the supernatant after overnight incubation of the samples.

BASICPROTOCOL 2

DIRECT BINDING TO G-ACTIN: FLUORESCENCE MEASUREMENTS

Actin can be fluorescently labeled using a variety of probes, such as 7-chloro-4-nitrobenzofurazane (NBD-Cl), pyrene, BODIPY, and N-iodoacetyl-N′-5sulfo-1-naphtyl-ethylenediamine (IAEDANS), that are differentially sensitive to local environments.Thus, binding of an actin-binding protein to the fluorescently labeled actin can leadto a change in fluorescence. The change in fluorescence (�F) reflects the fraction of actinin complex with the protein (Y) as follows:

Equation 13.6.1

where �Fmax is the maximal extent of change in fluorescence that is caused by the proteinat a saturating actin concentration, [PA] is the concentration of the protein-actin complex,and [A]0 is the total actin concentration.

Several fluorescent labels should be tried (see Support Protocols 2 and 3 for two ex-amples), to determine which is best suited to the purpose (see Troubleshooting). Thisassay may be performed with ATP-G-actin or ADP-G-actin to determine which actinspecies is preferentially bound by the protein. For example, β-thymosins and profilinprefer ATP-actin, whereas ADF/cofilin and twinfilin prefer ADP-actin.

Materials

1.5 µM 100% fluorescently labeled G-actin in G-buffer (see Support Protocols 2and 3)

Actin-binding protein of interest in G-buffer (see recipe)Spectrofluorometer (time base mode) at wavelengths appropriate for label.

1. Using a spectrofluorometer, record emission and excitation spectra of 1.5 µM fluo-rescently labeled G-actin in the absence and in the presence of a saturating amountof an actin-binding protein of interest in G-buffer. Choose the set of excitation andemission wavelengths at which the largest change is observed upon association ofactin with the protein.

A protein is at a saturating amount when its concentration is about ten times its Kd.

For NBD-labeled actin, λex = 475 nm and λem = 530 nm; for IDEDANS-labeled actin,λex = 340 nm and λem = 410 nm; and for prenyl-labeled actin, λex = 366-nm and λem =407.

This protocol can also be carried out in F-buffer (see recipe) to determine the effect ofionic strength on binding.

2. Measure the fluorescence of 1.5 µM fluorescently labeled G-actin alone and in thepresence of increasing amounts of the actin-binding protein across a range of 0 to100 µM.

OrganelleMotility

13.6.5


This protocol is suitable if the Kd is in the 0.5- to 10-µM range. If the Kd is much lower,the experiment should be carried out at a lower actin concentration (∼0.1 µM) and witha lower range of protein concentrations (0 to 1 µM). For proteins with an unknown Kd,the entire range should be tested at first, with a subsequent finer resolution of solutionconcentrations tested once an approximate Kd is known.

3. Plot the change in fluorescence versus the protein concentration and perform dataanalysis.

If a simple 1:1 complex (PA) is formed between actin (A) and the protein (P), then thevalue of the equilibrium dissociation constant for the PA complex, Kp, can be derived fromanalysis of the fluorescence curve:

Equation 13.6.2

where [P]0 is the total protein concentration.

This method is useful to measure Kp in the range of 0.1 to 10 µM. To obtain a reliablevalue of Kp, the concentration of G-actin in the assay should be as close as possible tothe value of Kp. A few preliminary trials at different actin concentrations are advisable.

BASICPROTOCOL 3

BINDING TO ACTIN DERIVED FROM A CHANGE IN THE RATE OFNUCLEOTIDE DISSOCIATION FROM G-ACTIN

Many G-actin-binding proteins affect the rate of nucleotide exchange on G-actin. Forinstance, profilin increases the rate of nucleotide dissociation from G-actin, whereasβ-thymosins and cofilin slow it down. On the condition that the protein (P) shuttlesmore rapidly between actin molecules (A) than nucleotide dissociates from either actinor the PA complex, the change in the observed first-order rate constant for nucleotidedissociation upon increasing the concentration of protein reflects the saturation of actinby the protein.

Dissociation of G-actin-bound ATP can be monitored by the increase in etheno-ATP(ε-ATP) fluorescence that takes place upon addition of a 10-fold molar excess of ε-ATP to the ATP-G-actin 1:1 complex. The fluorescence of ε-ATP increases 6-fold uponbinding to G-actin. Because ATP dissociation from G-actin is rate limited by dissociationof the bound metal ion (the Ca2+ ion in G0-buffer, which is G-buffer without ATP) andis affected by ionic strength, all measurements performed in the absence and presence ofthe actin-binding protein should be carried out under identical ionic conditions (i.e., theprotein to be tested must be equilibrated in the G0-buffer used for the assays).

Materials

50% suspension of Dowex-1-X8 (Sigma) or AG-1-X8 (Bio-Rad) strong anionexchange resin in G0-buffer

50 µM G-actin in G-buffer (see Support Protocol 1)G0-buffer: G-buffer (see recipe) without ATP1 mM etheno-ATP (ε-ATP), pH adjusted to 7.2 with NaOH, stock solution (Sigma)100 µM protein of interest in 10 mM Tris·Cl (pH 7.5)/1 mM dithiothreitol (DTT)

Microcentrifuge, 4◦CSpectrophotometer and quartz cuvettesSpectrofluorometer (time base mode): λex = 350 nm, λem = 410 nm, slits = 5- to

10-nm band width


Assembly

13.6.6


1. Add 150 µl of a 50% Dowex-1-X8 suspension in G0-buffer to 1 ml of 50 µM G-actinin a 1.5-ml microcentrifuge tube. Incubate 30 sec on ice while rotating the tube tomix actin and Dowex-1-X8. Microcentrifuge 1 min at 10,000 × g, 4◦C, and transfersupernatant to a clean tube.

2. Repeat step 1 with 150 µl fresh Dowex-1-X8.

The supernatant should contain ATP-G-actin in a 1:1 complex.

3. Measure the UV spectrum in a spectrophotometer to determine whether free ATPhas been removed.

The maximum of the UV spectrum should shift from 260 nm (typical of ATP) to 280 nm(typical of protein) and should show a minimum at 250 nm.

Most generally no dilution is required, even when using a 1-cm path cuvette. If the ab-sorbance of the sample is larger than 3.0 at the peak, the authors generally use a smallerpath cuvette (e.g. 0.2 cm) rather than diluting, and thus losing, an aliquot of the sample.

If ATP has not been totally removed, the Dowex treatment should be repeated.

4. Use the absorbance at 290 nm to measure the concentration of ATP-G-actin 1:1complex (ε = 0.617 cm2/mg at 290 nm).

The absorbance at 278 nm is 1.97-fold higher.

5. Dilute ATP-G-actin (1:1 complex) to 2 µM in G0-buffer. Mix together 50 µl of 2 µMATP-G-actin and 49 µl G0-buffer in a 1.5-ml microcentrifuge tube. Transfer solutionto a microcuvette. At time zero, add 1 µl of 1 mM ε-ATP (10 µM final) to start theexchange reaction. Monitor and record the increase in fluorescence (∼50% increase)at 410 nm (excitation wavelength, 350 nm) in a spectrofluorometer.

For G-actin in Ca2+-containing buffer at a low ionic strength, dissociation of ATP followsthe rate-limiting dissociation of bound metal ion. The rate of dissociation therefore varieswith the concentration of free Ca2+ (Nowak et al., 1988; Valentin-Ranc and Carlier, 1989),and the measurement period will need to be adjusted accordingly.

6. Prepare 10 dilutions of the protein of interest using G0-buffer to final concentrationsof 0 to 50 µM in a final volume of 50 µl.

7. Repeat step 5, replacing G0-buffer with the protein of interest diluted in G0-buffer.

An actin-binding protein that binds ATP-G-actin would be expected to cause an increaseor a decrease in the rate of ATP association.

8. To measure the change in rate of ATP dissociation, derive the first-order rate constant(kobs) at different concentrations of the protein.

The time course of ATP dissociation is a monoexponential process. The change in kobs

versus the total concentration of the protein reflects the formation of the PA complex.

9. Determine Y, the fraction of actin in complex with the protein, as follows:

Equation 13.6.3

where k0obs and k∞

obs are the rate constants measured in the absence of protein andin the presence of a saturating amount of protein, respectively, and [P]0 is the totalconcentration of protein. To derive KP, plot Y versus [P]0 as described in Equation13.6.2 (Perelroizen et al., 1995).

OrganelleMotility

13.6.7


BASICPROTOCOL 4

STEADY-STATE MEASUREMENTS OF ACTIN ASSEMBLY: CRITICALCONCENTRATION PLOTS

The 25-fold increase in fluorescence of pyrenyl-labeled actin linked to the G-to-F tran-sition is the most accurate and convenient tool for measuring the critical concentration(Css) for actin polymerization at the barbed and pointed ends.

Materials

50 µM pyrenyl-labeled G-actin (see Support Protocol 2), 10% labeledG-buffer (see recipe)Gelsolin solution: 100 µM gelsolin (Sigma) in 10 mM Tris·Cl (pH 7.5)/1 mM

dithiothreitol (DTT), for assaying actin with capped barbed ends onlyProtein of interestKM solution: 500 µl of 4 M KCl/20 µl of 1 M MgCl2 (APPENDIX 2A)Spectrofluorometer (time base mode): λex = 360 nm, λem = 407 nm,

slits = 5- to 10-nm band width

1a. To assay actin with free barbed ends: Dilute 200 µl of 50 µM pyrenyl-labeled G-actinto 1 ml with G-buffer (continue with step 2).

1b. To assay actin with capped barbed ends: To pretreat a 50 µM pyrenyl-labeled G-actinsolution, mix together the following:

200 µl of 50 µM pyrenyl-labeled G-actin (10 µM final)800 µl G-buffer0.4 µl gelsolin solution (0.4 µM final).

Gelsolin needs calcium; EGTA must not be present in any solutions used for this assay.

2. To polymerize actin, add 50 µl KM solution (5%, v/v) to 1 ml untreated or gelsolin-pretreated G-actin (step 1a or 1b).

Two complementary types of assays will be carried out. In the first (step 3), the total actinconcentration is varied in the presence of a constant amount of the protein of interest. In thesecond (step 4), actin is maintained at a given concentration, whereas the concentrationof the protein of interest is varied.

3. Prepare fifteen dilutions of F-actin from 0 to 2 µM in F-buffer along with a singleconcentration of the protein of interest. The final volume should be 100 µl. Be sureto include control samples that do not contain the protein.

4. Prepare twelve dilutions of the protein of interest (e.g., from 0 to 10 or 20 µM) inF-buffer along with a single concentration of F-actin. The final volume should be100 µl.

5. Incubate all samples overnight in the dark either at room temperature or at 4◦C,depending on the stability of the protein of interest.

6. Add 10 µl of each protein dilution to a sample tube (step 2). Add 5 µl KM solutionto all 20 tubes and incubate overnight at room temperature.

7. Read fluorescence of pyrenyl-actin from control and sample tubes in a spectrofluo-rometer.

8. Plot fluorescence intensity versus either total actin concentration (samples fromstep 3) or concentration of the protein of interest (samples from step 4) and de-termine Css.


Assembly

13.6.8


The increase in pyrenyl-actin fluorescence is proportional to the amount of F-actin as-sembled. Fluorescence increases linearly with actin concentration. The slope is very lowwhen actin is monomeric and 25-fold higher when actin is polymerized. Css is definedas the total actin concentration at which a large change in slope takes place (i.e., abovewhich actin polymerizes).

Under physiological ionic conditions, a value of 0.1 µM is measured for Css when barbedends are free, and 0.6 µM when barbed ends are capped. The value of 0.6 µM is the crit-ical concentration (Cc) at the pointed ends. The value of 0.1 µM is slightly above the Css

at the barbed ends, because pointed ends also contribute to assembly. Because the con-tribution of barbed ends is largely predominant, however, it is routinely considered that0.1 µM is very close (within 15%) to the Css at the barbed ends.

BASICPROTOCOL 5

INITIAL RATE OF FILAMENT GROWTH AT BARBED OR POINTED ENDS

Initial rates of filament growth at barbed ends are measured spectrofluorometrically us-ing spectrin-actin seeds or F-actin filaments. Growth at pointed ends is measured usinggelsolin-actin seeds [i.e., the gelsolin(actin)2 (GA2) tight complex] or gelsolin-cappedfilaments.

Materials

Actin seeds (choose one):100 nM gelsolin-actin (for pointed-end growth): add 2.5 molar equiv G-actin to

gelsolin in G-buffer5 nM spectrin-actin isolated from human blood (Casella et al., 1986; for

barbed-end growth) in G-buffer5 µM F-actin (with or without 20 nM gelsolin)

KME solution (see recipe)5 µM pyrenyl-labeled (10%) G-actin (see Support Protocol 2)G-buffer (see recipe)100 µM protein of interest in 10 mM Tris·Cl (pH 7.5)/1 mM dithiothreitol (DTT)

Microcuvettes suitable for spectrofluorometerSpectrofluorometer (time base mode): λex = 366 nm, λem = 407 nm, slits = 5- to

10-nm band width

Additional reagents and equipment to generate a calibration curve (see BasicProtocol 4)

1. Mix the following in a microcuvette:

51 µl G-buffer4 µl actin seeds (final 4 nM for gelsolin-actin, 0.2 nM for spectrin-actin, or 0.2

µM for F-actin)5 µl KME solution40 µl 5 µM pyrenyl-labeled G-actin.

2. Immediately place the cuvette in a spectrofluorometer (time zero) and read the in-crease in fluorescence at 407 nm versus time for a few minutes.

3. Repeat the assay (steps 1 and 2) in the presence of the protein of interest at variousconcentrations (about twelve to fifteen different concentrations in a final range of 0 to100 nM for a Kd in the nanomolar range and 0 to 100 µM for a Kd in the micromolarrange), diluted from a 100 µM solution. Adjust the amount of G-buffer added to theassay to keep a final volume of 100 µl.

For proteins with an unknown Kd, the entire range should be tested at first, with a subse-quent finer resolution of solution concentrations tested once an approximate Kd is known.

OrganelleMotility

13.6.9


4. Determine the initial rate of fluorescence increase in absorbance units (arbitrary unitof fluorescence intensity) per second (AU/sec) and convert to molar amount of F-actinassembled per second using a calibration curve (see Basic Protocol 4).

The initial rate of actin assembly or velocity (V) measured when filaments or seeds at aconcentration [F] are placed in a solution of G-actin at concentration C is

Equation 13.6.4

Equation 13.6.5

Where k+ and k− are the rate constants for G-actin association to and dissociation from,respectively, the filament end, and Cc is the critical concentration (k−/k+). The effectof the protein on V provides insight into its function. Barbed-end capping and G-actinsequestering functions give different results (see Anticipated Results).

BASICPROTOCOL 6

MEASUREMENT OF DILUTION-INDUCED DEPOLYMERIZATION OFFILAMENTS

The depolymerization rate of actin filaments is monitored by the decrease in pyrenyl fluo-rescence upon dilution of the solution of pyrenyl-labeled F-actin in F-buffer to a final actinconcentration that is lower than the critical concentration (Css). When depolymerizationfrom the pointed end is to be measured, gelsolin-capped filaments are used.

Materials

G-buffer (see recipe)4 M KCl100 mM MgCl2 (APPENDIX 2A)50 µM G-actin in G-buffer (see Support Protocol 1)50 µM G-actin, 50% to 100% pyrenyl labeled in G-buffer (see Support Protocol 2)Gelsolin solution: 100 µM gelsolin in 10 mM Tris·Cl (pH 7.5)/1 mM dithiothreitol

(DTT), for assaying depolymerization from pointed ends only100 µM protein of interest in 10 mM Tris·Cl (pH 7.5)/1 mM DTTF-buffer (see recipe)

Microcuvettes, suitable for spectrofluorometerSpectrofluorometer (time base mode): λex = 366 nm, λem = 407 nm,

slits = 5- to 10-nm band width

1a. To measure total depolymerization from the actin filament: Mix together the followingin a 1.5-ml microcentrifuge tube:

865 µl G-buffer25 µl 4 M KCl (0.1 M final)10 µl 100 mM MgCl2 (1 mM final)50 µl 50 µM G-actin (5 µM final)50 µl 50 µM pyrenyl-labeled G-actin (5 µM final).

Incubate 30 min at room temperature.

1b. To measure depolymerization from the pointed end of the actin filament: Mix togetherthe same reagents as in step 1a but add 0.2 µl gelsolin solution (20 nM final gelsolinconcentration). Incubate 1 hr at room temperature.

Gelsolin needs calcium; EGTA must not be present in any solutions used for this assay. Alonger incubation allows a more complete exchange of calcium and magnesium.

Steps 1a and 1b yield 50% pyrenyl-labeled F-actin at 5 µM.


Assembly

13.6.10


2. Dilute a 100 µM solution of the protein of interest in F-buffer as needed to generatestock solutions to add to the assay. Generate stocks that will give twelve to fifteendifferent concentrations in a final range of 1 to 100 nM for a Kd in the nanomolarrange and 0 to 100 µM for a Kd in the micromolar range.

For proteins with an unknown Kd, the entire range should be tested at first, with a subse-quent finer resolution of solution concentrations tested once an approximate Kd is known.

3. At time zero, dilute 2 µl of the 5 µM actin solution (step 1a or 1b) 100-fold inF-buffer in a spectrofluorometer microcuvette and measure the decrease of pyrenylfluorescence in a spectrofluorometer in the absence or in the presence of the proteinof interest at various concentrations. Record the change in fluorescence for a fewminutes.

Total depolymerization occurs because the final actin concentration (50 nM) is lower thanthe critical concentration (0.1 µM).

4. Plot the change in fluorescence versus time. Calculate the initial rate of depolymer-ization from the slope of the time-dependent decrease in pyrenyl fluorescence.

5. Plot a calibration curve of the fluorescence of pyrenyl-labeled F-actin versus totalactin concentration. Convert fluorescence units into micromolar F-actin and thenexpress the initial rate of depolymerization in micromolar per second.

BASICPROTOCOL 7

MEASUREMENTS OF THE TREADMILLING OF ACTIN FILAMENTS

The turnover of filaments is evaluated by measuring the rate at which fluorescently labeledetheno (ε)-ADP-F-actin filaments assemble from ε-ATP-G-actin subunits and becomenonfluorescent ADP-F-actin filaments after addition of ATP to the medium. Turnoveroccurs via the consecutive steps of dissociation of ε-ADP-actin from the pointed end,exchange of ATP for bound ε-ADP on G-actin in the medium, and association of ATP-G-actin to barbed ends. The fluorescence of ε-ADP is 6-fold lower in the free state thanit is in the actin-bound state.

Materials

50% (w/v) suspension of Dowex-1-X8 (Sigma) or AG-1-X8 (Bio-Rad) stronganion exchange resin in G0-buffer

50 µM G-actin in G- buffer (see Support Protocol 1), store on ice5 mM etheno-ATP (ε-ATP), pH adjusted to 7.2 with NaOH, stock solution (Sigma)KME solution (see recipe)G-buffer (see recipe)100 µM protein of interest in 10 mM Tris·Cl (pH 7.5)/1 mM dithiothreitol (DTT)200 mM ATP, pH adjusted to 7.2 with NaOH, stock solution (Sigma)

Microcentrifuge, 4◦CSpectrophotometer microcuvette, suitable for spectrofluorometerSpectrofluorometer (time base mode; PM voltage adequate): λex = 350 nm,

λem = 410 nm, slits = 5- to 10-nm band width

1. Add 150 µl of a 50% Dowex-1-X8 suspension in G0-buffer to 1 ml of 50 µM G-actinin a 1.5-ml microcentrifuge tube. Incubate 30 sec at 0◦C. Microcentrifuge 1 min at10,000 × g, 4◦C, and transfer supernatant to a clean tube.

2. Repeat step 1 with 150 µl fresh Dowex-1-X8.

The supernatant should contain ATP-G-actin in a 1:1 complex.

3. Add 50 µl of 5 mM ε-ATP (0.25 mM final). Let the solution sit overnight on ice toallow exchange of ε-ATP for bound ATP.

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13.6.11


4. Repeat steps 1 and 2 to obtainε-ATP-G-actin in a 1:1 complex. Measure concentrationof G-actin spectrophotometrically (see Basic Protocol 3, steps 3 and 4).

5. Add 5 mM ε-ATP to make a final concentration of 50 µM free ε-ATP.

6. In a 1.5-ml microcentrifuge tube, mix together ε-ATP-G-actin (5 µM final) and 5%(v/v) KME buffer along with G-buffer as needed.

Adding 5% (v/v) KME buffer will lead to polymerization, yielding ε-ATP-F-actin.

7. Divide the sample in 100-µl aliquots among several tubes. Add 100 µM protein ofinterest to individual tubes to cover a final range of 0 to 50 µM. Adjust volumes asneeded with G-buffer.

This step is used to test the effects of actin depolymerizing factor (ADF), profilin, or otherproteins.

8. Transfer each sample to a spectrofluorometer microcuvette and measure its ε-ATPfluorescence intensity in a spectrofluorometer at 410 nm. At time zero, add 0.25 µlof 200 mM ATP (500 µM final).

9. Record the decrease in ε-ATP fluorescence using a time-based scan over at least2 hr. Determine the rate of fluorescence decrease, which provides an indication offilament turnover.

SUPPORTPROTOCOL 1

ACTIN PURIFICATION FROM RABBIT MUSCLE

Actin for these protocols is purified from an acetone powder of rabbit muscle. Actin isalso available commercially (Cytoskeleton). To prepare commercially purchased actinfor the other protocols in this unit, follow the manufacturer’s instructions.

Materials

Acetone powder of rabbit muscle (Cytoskeleton)Extraction buffer (see recipe), 4◦CSolid KCl and 4 M KClBuffer D1 (see recipe)1 M MgCl2 (APPENDIX 2A)G-buffer (see recipe)

500-ml beakerSorvall centrifuge and A641 rotor (or equivalent), 4◦CGlass wool20◦C water bathLarge (3-cm wide) dialysis bags (MWCO 14,000)Dounce homogenizer (glass-Teflon; 20-ml working volume)Sonicator with microtip (e.g., Vibra-cell; Sonics & Materials)10-ml ultracentrifuge tubesBeckman ultracentrifuge and 70.1 Ti rotor (or equivalent), 4◦C2.5 × 100–cm gel filtration column (Superdex-200 prep grade; Amersham

Biosciences)Spectrophotometer (290 nm) and quartz cuvettes

Extract actin1. Place 9 g acetone powder in a 500 ml beaker on ice and slowly add 270 ml cold

extraction buffer while gently stirring with a glass rod to thoroughly wet powder. Letsit on ice for 30 min, stirring gently once every 10 min.

Gentle stirring minimizes α-actinin extraction.


Assembly

13.6.12


2. Centrifuge 45 min at 25,000 × g, 4◦C, in a Sorvall A641 rotor. Filter supernatantthrough glass wool into a graduated cylinder and measure volume.

3. Transfer filtrate to a beaker. Add solid KCl to 3.3 M while stirring (add KCl all atonce).

The KCl adds ∼10% to the volume, which must be figured in when deciding how muchKCl to add (usually 55 to 57 g for ∼220 ml).

4. Place in a 20◦C water bath and stir until temperature reaches 15◦C. Place on icewithout stirring until temperature returns to 5◦C. Centrifuge 30 min at 25,000 × g,4◦C.

A tight white pellet indicates the presence of actin-α-actinin networks.

5. Filter supernatant through glass wool, transfer to a large dialysis bag, and dialyzeovernight at 4◦C against 32 vol (usually 6 to 8 liters) buffer D1.

Release tropomyosin from F-actin6. Add 4 M KCl to a final concentration of 0.8 M (0.22 vol of 4 M KCl) and stir for 1.5

hr at 4◦C.

7. Centrifuge 3.5 hr at 100,000 × g, 4◦C, in the Sorvall A641 rotor. Decant and discardsupernatant. Dislodge pellet and transfer to a Dounce homogenizer, using 25 ml(total) extraction buffer to transfer pellet and clean centrifuge tube.

8. Homogenize pellet with 20 strokes while keeping homogenizer on ice.

9. Add 75 µl of 1 M MgCl2 and 375 µl of 4 M KCl and adjust the volume to 38.6 mlwith extraction buffer. Leave at 4◦C overnight without stirring.

10. Add 9 ml of 4 M KCl and 2.37 ml extraction buffer and stir for 1.5 hr at 4◦C.Centrifuge 3.5 hr at 100,000 × g, 4◦C.

11. Resuspend pellets with 20 to 30 ml extraction buffer and homogenize as described(step 8).

12. Dialyze against 2 liters G-buffer for 2 days at 4◦C with one change of buffer.

13. Insert microtip of a sonicator into dialysis bag, sonicate twice for 30 sec each onlow-power setting. Reclose bag, change dialysis buffer, and continue dialysis oneadditional day.

Purify actin14. To clarify actin, transfer to 10-ml ultracentrifuge tubes and centrifuge 2 hr at 400,000

× g, 4◦C, in a Beckman 70.1 Ti rotor.

15. Load supernatant onto a 2.5 × 100–cm gel filtration column. Run 2 liters G-bufferthrough column at 1 ml/min 4◦C. Collect 5-ml fractions.

16. Read A290 of each fraction in a spectrophotometer and pool the fractions showing anabsorbance at 290 nm.

17. Determine the concentration of G-actin in the pooled fractions using A290 as follows:

Equation 13.6.6

where C is the actin concentration in milligrams per milliliter.

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18. Store actin on ice for up to 3 weeks

The typical yield of this preparation is 40 ml of 50 µM (2.1 mg/ml) G-actin (mol. wt. =42 kDa).

SUPPORTPROTOCOL 2

PREPARATION OF PYRENYL-LABELED ACTIN

N-pyrenyliodoacetamide (NPI) is a fluorescent compound that can be used to label actinspecifically on Cys374. Its fluorescence changes in response to the local environment;in particular, it increases 25-fold upon polymerization of G-actin into F-actin. Pyrenyl-labeled actin can also be used to monitor binding of proteins to actin.

Additional Materials (also see Support Protocol 1)

4 mg/ml (∼95 µM) G-actin in G-buffer (see Support Protocol 1)Buffer A (see recipe)N-pyrenyliodoacetamide (NPI; Sigma)Dimethylformamide0.2 M dithiothreitol (DTT; APPENDIX 2A)Rotating wheel, 4◦C

1. Mix 10 ml of 4 mg/ml G-actin and 10 ml buffer A. Incubate 1 hr at 25◦C (roomtemperature) to polymerize actin.

2. Transfer polymerized actin (F-actin) to a large dialysis bag and dialyze overnightagainst 2 liters buffer A at 4◦C.

As NPI is destroyed by the light, all labeling steps must be carried out in the dark.

3. Prepare 1 ml solution of 14 mg/ml NPI in dimethylformamide in a 1.5-ml microcen-trifuge tube. Add 0.2 ml NPI solution to 20 ml F-actin (step 2) in a 50-ml conicalcentrifuge tube wrapped with aluminum foil. Mix by inverting the tube several times.Incubate overnight on a rotating wheel in the dark at room temperature.

The solution becomes white after mixing because of insoluble NPI.

4. Add 50 µl of 0.2 M DTT to stop the reaction. Centrifuge 15 min at 1,000 × g, 4◦C,to remove excess insoluble NPI.

5. Centrifuge supernatant 3.5 hr at 100,000 × g, 4◦C in a Sorvall A641 rotor.

6. Dislodge the actin pellet and resuspend with 5 ml G-buffer. Homogenize with 20strokes of a Dounce homogenizer on ice.

7. Continue with actin purification as described (see Support Protocol 1, steps 12 to18), except dialyze against 1 liter G-buffer in steps 12 and 13 and perform all stepsin the dark.

G-actin thus prepared is 100% labeled. For a 10% labeled G-actin solution, 0.9× molesof regular unlabeled actin (see Support Protocol 1) is mixed with 0.1× moles of 100%pyrenyl-labeled G-actin in G-buffer.

SUPPORTPROTOCOL 3

PREPARATION OF7-CHLORO-4-NITROBENZENO-2-OXA-1,3-DIAZOLE-LABELED ACTIN

7-Chloro-4-nitrobenzeno-2-oxa-1,3-diazole (NBD-Cl) is a fluorescent compound thatcan be used to label actin specifically on Lys373. Its fluorescence changes in responseto the local environment. Thus, NBD-labeled actin can be used to monitor binding ofproteins to actin. The fluorescence of NBD-labeled actin also increases 2.6-fold uponpolymerization of G-actin into F-actin.


Assembly

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Additional Materials (also see Support Protocol 1)

0.1 M KCl (APPENDIX 2A)50 µM G-actin in G-buffer (see Support Protocol 1)100 mM N-ethylmaleimide (NEM), prepare fresh20 mM dithiothreitol (DTT; APPENDIX 2A)G-buffer (see recipe) with and without DTT and NaN3

F-buffer (see recipe) without DTT and NaN3

10 mM 7-chloro-4-nitrobenzeno-2-oxa-1,3-diazole (NBD-Cl) indimethylformamide

15◦C water bath

Label actin with NEM1. Add 750 µl of 0.1 M KCl to 15 ml of 50 µM G-actin in G-buffer in a tube. Add 375

µl of 100 mM NEM (2.5 mM final), mix, and incubate as follows:

15 min at room temperature15 min at 15◦C3 hr at 0◦C (on ice).

2. Stop the reaction by adding 375 µl of 20 mM DTT (5 mM final). Centrifuge 2 hr at200,000 × g, 4◦C a Beckman ultracentrifuge.

3. Discard supernatant and use 20 ml G-buffer with DTT and NaN3 to resuspend pelletand transfer to a Dounce homogenizer. Homogenize pellet with 20 strokes on ice.

The pellet contains NEM-F-actin.

Label NEM-actin with NBD4. Transfer homogenized actin to a large dialysis bag and dialyze 2 days against 1 liter

G-buffer without DTT and NaN3 at 4◦C.

5. Centrifuge 2 hr at 400,000 × g, 4◦C, in the Beckman 70.1 Ti rotor.

NEM-G-actin is in the supernatant.

6. Dilute supernatant 2-fold to a total volume of 30 ml with F-buffer without DTT andNaN3. Let polymerize overnight at 4◦C in the dark.

The actin concentration should be at 1 to 2 mg/ml to optimize NBD labeling.

7. Add 1.2 ml of 10 mM NBD-Cl (0.4 mM final). Incubate 5 hr at 15◦C.

Purify labeled actin8. Centrifuge NBD-labeled F-actin 2 hr at 400,000 × g, 4◦C. Homogenize pellet and

dialyze as described (steps 3 and 4).

9. Centrifuge 2 hr at 20,000 × g, 4◦C, in the Sorvall A641 rotor to eliminate aggregates.

NBD-labeled G-actin is in the supernatant.

10. Purify actin as described (see Support Protocol 1, steps 14 to 18) except perform allsteps in the dark.

G-actin thus prepared is 100% labeled. For a 10% labeled G-actin solution, 0.9× molesof regular unlabeled actin (see Support Protocol 1) is mixed with 0.1× moles of 100%NBD-labeled G-actin in G-buffer.

OrganelleMotility

13.6.15


REAGENTS AND SOLUTIONSUse deionized or distilled water in all recipes and protocol steps. Prepare all buffers 1 dayin advance. Store at 4◦C for up to 1 week. For common stock solutions, see APPENDIX 2A; forsuppliers, see SUPPLIERS APPENDIX.

Buffer A20 ml 0.5 M Tris·Cl, pH 8 (5 mM final; APPENDIX 2A)50 ml 4 M KCl (0.1 M final)4 ml 1 M MgCl2 (2 mM final; APPENDIX 2A)1 ml 0.2 M CaCl2 (0.1 mM final; APPENDIX 2A)5 ml 0.2 M ATP (0.5 mM final)1 ml 0.2 M dithiothreitol (DTT; 0.1 mM final; APPENDIX 2A)H2O to 2 liters

Buffer D12 mM Tris·Cl, pH 7.8 (APPENDIX 2A)1 mM MgCl2 (APPENDIX 2A)1 mM dithiothreitol (DTT; APPENDIX 2A)

Extraction buffer2 mM Tris·Cl, pH 7.8 (APPENDIX 2A)0.5 mM ATP0.1 mM CaCl2 (APPENDIX 2A)0.01% (w/v) NaN3

1 mM dithiothreitol (DTT; APPENDIX 2A)

CAUTION: Sodium azide is poisonous. Follow appropriate precautions for handling,storage, and disposal.

F-bufferG-buffer (see recipe) supplemented with 0.1 M KCl (APPENDIX 2A) and 1 mM

MgCl2 (APPENDIX 2A).

G-buffer5 mM Tris·Cl, pH 7.8 (APPENDIX 2A)0.2 mM ATP0.1 mM CaCl2 (APPENDIX 2A)1 mM dithiothreitol (DTT; APPENDIX 2A)0.01% (w/v) NaN3

CAUTION: Sodium azide is poisonous. Follow appropriate precautions for handling,storage, and disposal.

G0-buffer is G-buffer without ATP.

KME solution500 µl 4 M KCl (2 M final)20 µl 1 M MgCl2 (20 mM final; APPENDIX 2A)16 µl 250 mM EGTA (4 mM final)464 µl H2O

To polymerize actin, 0.5% (v/v) KME solution is added to G-actin (final 0.1 M KCl, 1mM MgCl2, 0.2 mM EGTA).


Assembly

13.6.16


COMMENTARY

Background Information

G-actin-sequestering proteinsG-actin sequesterers bind G-actin in a non-

polymerizable complex. β-Thymosins are themost abundant typical G-actin-sequesteringproteins (Cassimeris et al., 1992; Safer andNachmias, 1994). The β-thymosin-actin com-plex associates with neither the barbed northe pointed end of the filament, and thus β-thymosins do not affect filament assembly dy-namics or motility. It is often thought that theseproteins buffer the free ATP-G-actin concen-tration in cells. On the contrary, both in vitroand in vivo evidence shows that β-thymosinsreact passively to changes in the steady-stateconcentration of free G-actin (Carlier et al.,1993; Safer and Nachmias, 1994).

Profilin and profilin-like proteinsProfilin also binds G-actin but acts differ-

ently from sequestering proteins. The profilin-actin complex associates productively with thebarbed end (the dynamic end of the actinfilament) but not with the pointed ends ofthe filaments (Pantaloni and Carlier, 1993).Thus, profilin merely sequesters actin when allbarbed ends are capped but enhances barbed-end growth as soon as a few of them are notcapped. Recently, other proteins like the β-thymosin repeat proteins and, more generally,the actin-binding WH2 (WASP-homology 2)

Figure 13.6.2 (at right) Functional assays of a G-actin-sequestering protein. (A) Steady statemeasurements of F-actin in the presence of increasing amounts of the sequestering protein (seeBasic Protocol 4). F-actin (1.5 µM, 10% pyrenyl labeled) with free barbed ends (solid line) orgelsolin-capped barbed ends (dashed line) was supplemented by the protein as indicated. Lineardepolymerization was consistent with a Kd of 2 µM. (B) Steady-state measurements of F-actinin the presence of the sequestering protein. Experiments were conducted as in A at 2, 1.5, and1 µM F-actin with free barbed ends. The parallel straight lines are consistent with a Kd of 2 µM. (C)Critical-concentration (Css) measurements for actin alone or in the presence of 2 µM sequesteringprotein. The sequestering protein shifts Css for both free and capped barbed ends, in agreementwith the Kd obtained in B. Solid line, free barbed ends alone; short-dashed line, capped barbed endsalone; dotted line, free barbed ends with sequestering protein; long-dashed line, capped barbedends with sequestering protein. (D) Depolymerization rate measurements (see Basic Protocol 6).The solution of 2.5 µM F-actin (50% pyrenyl labeled) with free barbed ends was diluted 20-fold inF-buffer at time zero in the absence (solid line) and presence (dotted line) of 10 µM sequesteringprotein. The sequestering protein has no effect on actin depolymerization. (E) The sequesteringprotein inhibits pointed-end growth of actin filaments (see Basic Protocol 5). The rate of pointed-end growth was measured in the presence of 2.5 µM Mg-ATP-G-actin, 2 nM gelsolin-actin seeds,and the sequestering protein at the indicated concentration. The inhibition observed is consistentwith a Kd of 2 µM. (F) The sequestering protein inhibits barbed-end growth of actin filaments.The rate of barbed-end growth was measured in the presence of 2.5 µM Mg-ATP-G-actin and0.16 nM spectrin-actin seeds. The inhibition observed is consistent with a Kd of 2 µM, as inE. (G) Co-sedimentation assay (see Basic Protocol 1). F-actin (10 µM) was sedimented aloneor in the presence of increasing amounts of the sequestering protein. The sequestering proteindepolymerizes F-actin, leading to an increasing amount of G-actin in the supernatant (S) and lessG-actin in the pellet (P). Lines shown in A-F represent typical experimental curves, derived fromknown proteins (e.g., thymosin β4).

domains, which are inserted in a number ofproteins involved in motility, have been foundto behave functionally like profilin (Egile et al.,1999; Boquet et al., 2000). As a result, pro-filin and its functional homologs improve theprocessivity of treadmilling in synergy withactin depolymerizing factor (ADF) and so en-hance actin-based motility (Loisel et al., 1999;Paunola et al., 2002).

Capping proteinsCapping proteins bind tightly to the barbed

end and block actin association and dissocia-tion (Hug et al., 1995; Sun et al., 1995). Cap-ping proteins are required for efficient motil-ity of many cells. By blocking a large frac-tion of the barbed ends at the steady state ofactin assembly, capping proteins funnel thetreadmilling process: pointed-end depolymer-ization of many capped filaments feeds thegrowth of a few noncapped filaments, whichindividually grow faster than if the other fila-ments were not capped. Capping proteins co-operate with ADF to accelerate actin-basedmotility.

ADF/cofilin proteinsADF/cofilin proteins are essential proteins

in morphogenetic and motile processes. ADF/cofilin increases the turnover of actin fil-aments, which powers actin-based motility(Didry et al., 1998; Carlier et al., 1999). The

OrganelleMotility

13.6.17


function of ADF/cofilin proteins is complexbecause under physiological ionic conditions,ADF/cofilin recognizes the ADP-bound formof both F- and G-actin with a 100-fold higheraffinity than ATP-bound actin. ADF/cofilinmodifies both the structure and the assemblydynamics of filaments. (1) The rate of dissocia-tion of ADF-F-actin from the pointed ends (therate-limiting step in treadmilling) is 30-fold

higher than the rate of dissociation of F-actin.(2) The dissociation of ADP from ADF-G-actin is 10- to 20-fold slower than the dissoci-ation of ADP from G-actin. The combinationof these two properties affects actin dynam-ics at steady state. The physiologically rele-vant effect of ADF in actin-based motility isto enhance treadmilling by increasing the rate-limiting step in the ATPase cycle of actin.

Figure 13.6.2 Legend at left.


Assembly

13.6.18


TroubleshootingBecause in many protocols pyrenyl-labeled

or NBD-labeled actin is used, it is importantto check that measurements of F-actin con-centrations using these probes are not biasedby possible changes in fluorescence of labeledactin upon binding by the protein of interest.For instance, ADF quenches the fluorescenceof NBD- and pyrenyl-F-actin, hence alterna-tive measurements (e.g., using light scatter-ing or sedimentation) are required to elucidatethe function of this protein. Some components,such as glycerol in a buffer, could also modifythe fluorescence of labeled actin.

Labeling of actin may also affect its affinityfor the protein to be tested (Uyemura et al.,1978). It is recommended to carry out testsat different percentages of labeled actin andto verify that the result is independent of thepercentage of labeled actin in the mix. It isalso appropriate to use competition assays todetermine the affinity of labeled and unlabeledactin for the protein.

The solution of F-actin should be manipu-lated carefully to avoid breaking of filaments.

Anticipated Results

G-actin-sequestering proteinsIn the presence of β-thymosin, both β-

thymosin (T) and actin filaments (A) are atequilibrium with free G-actin at the steadystate (critical) concentration, Css. The dou-ble equilibrium implies that the amount of TAcomplex formed at equilibrium in the presenceof F-actin is determined by the value of Css

and the total β-thymosin concentration, [T]0,as follows (Fig. 13.6.2A):

Equation 13.6.7

where KT is the equilibrium dissociation con-stant of the TA complex.

According to this equation, sequesteringproteins shift the critical concentration plotsto a higher apparent value. The apparent criti-cal concentration is the sum of Css and [TA]given by Equation 13.6.7. Parallel Css plotsare obtained at different values of [T]0 (Fig.13.6.2C).

Changes in Css are elicited by proteins thataffect the dynamics of assembly; these changesare amplified by sequestering proteins. For in-stance, capping proteins increase the value ofCss up to the critical concentration at pointedends (Cc). It is easy to see (Equation 13.6.7)that when more filaments are capped, moreactin is sequestered at a given concentration[T]0 (i.e., F-actin depolymerizes in the cell).In contrast, when the proportion of capped fil-aments is lowered by creation of new barbedends, Css becomes lower; hence the pool of se-questered actin decreases and F-actin increases(e.g., upon platelet stimulation).

The ability of sequestering proteins to de-polymerize F-actin in a manner that dependson the value of Css is used to determine thevalue of KT. Addition of increasing amountsof β-thymosin to a given amount of F-actincauses depolymerization because of formationof TA with a linear dependence on [T]0. The(negative) slope of the plot of F-actin versus[T]0 is Css/(Css + KT), which is independentof the amount of actin. Hence at different con-centrations of actin, the decreases in F-actinare parallel straight lines (Fig. 13.6.2B). The

Figure 13.6.3 (at right) Functional assays of profilin-like proteins (barbed-end assembly-promoting factors). (A) Steady-state measurements of F-actin in the presence of increasingamounts of profilin-like protein. The experiment was conducted as in Figure 13.6.1A. When barbedends are capped (dashed line), the profilin-like protein depolymerizes F-actin. Linear depolymer-ization was consistent with a Kd of 2 µM. When the barbed ends are free (solid line), the profilin-likeprotein fails to depolymerize F-actin. (B) Critical-concentration (Css) measurements for actin aloneand in the presence of 2 µM profilin-like protein. The experiment was conducted as in Figure13.6.2C. Solid line, free barbed ends alone; short-dashed line, capped barbed ends alone; dottedline, free barbed ends with sequestering protein; long-dashed line, capped barbed ends with se-questering protein. (C) Profilin-like protein inhibits pointed growth of actin filaments. The inhibitionis consistent with a Kd of 2 µM. The experiment was conducted as in Figure 13.6.2E. (D) Theprofilin-like protein fails to inhibit barbed-end growth of actin filaments. The experiment was con-ducted as in Figure 13.6.2F. (E) Co-sedimentation assay in the absence or presence of profilin-likeprotein. The experiment was conducted as in Figure 13.6.2G. (F) Depolymerization rate measure-ments in the absence (solid line) and presence (dotted line) of 10 µM profilin-like protein. Theexperiment was conducted as in Figure 13.6.2D. The profilin-like proteins have no effect on actindepolymerization. Lines shown in A-D and F represent typical experimental curves derived fromknown proteins (e.g., profilin).

OrganelleMotility

13.6.19


slope is steeper when filaments are capped be-cause the value of Css is higher. Analysis ofthe slopes of F-actin versus [T]0 yields thesame value of KT, however, when barbed endsare free or capped. This property identifies asequestering function.

In a co-sedimentation assay, sequesteringproteins cause an accumulation of actin in thesupernatant, which is dependent on the valueof Css (i.e., it is greater when barbed ends are

capped; Fig. 13.6.2G). The sequestering pro-teins are always present in the supernatants.

Sequestering proteins, in forming a non-polymerizable complex with actin, cause adecrease in the rate of filament growthwhen tested at a given G-actin concentration(Fig. 13.6.2D,F). The equilibrium dissociationconstant for the TA complex is derived from thedependence of the elongation rate at barbedends on [T]0 using the following equation,



Assembly

13.6.20


which describes saturation of G-actin in thegrowth assay by the sequestering protein:

Equation 13.6.8

where V is the initial rate of elongation mea-sured at concentration [T]0 of the sequesteringprotein, V0 is the initial rate of elongation inthe absence of the sequestering protein, [A]0

is the total G-actin concentration (assumed forsimplicity to be much higher than the criti-cal concentration at the barbed end), and KT

is the equilibrium dissociation constant for thesequestering protein-actin complex. G-actin-sequestering proteins display no effect on therate of depolymerization at either end of thefilament (Fig. 13.6.2D).

Profilin and profilin-like proteinsAt the steady state of assembly, when

the barbed ends of filaments are capped,profilin-like proteins cause depolymerizationof F-actin (Fig. 13.6.3). The amount ofdepolymerized actin increases linearly withthe concentration of the protein, which is con-sistent with sequestration of Mg-ATP-G-actinby protein in a 1:1 complex and with an equi-librium constant as calculated with Equation13.6.7; this behavior is typical of a seques-tering protein. In contrast with sequesteringproteins, however, profilin-like proteins fail todepolymerize F-actin when barbed ends arefree, because the protein-actin (PA) complexparticipates in monomer-polymer exchangesat the barbed ends. Because both actin (A)

and the PA complex contribute to stabilizationof filament barbed ends, the presence of PAlowers the contribution of A (i.e., the partialcritical concentration of A is lowered byprofilin-like proteins; Fig. 13.6.3A,B).

Co-sedimentation assays confirm theseconclusions. When barbed ends are free,profilin-like proteins fail to depolymerizeF-actin, but depolymerization is readily ob-served if barbed ends are capped. Profilin-like proteins are present in the supernatantfraction (Fig. 13.6.3E). Finally, these proteinsdisplay no effect on the initial rate of de-polymerization at either end of the filament(Fig. 13.6.3F).

In seeded growth assays, profilin-like pro-teins prevent association of actin to the pointedend of the filament (Fig. 13.6.3C). The equi-librium dissociation constant (0.2 µM for pro-filin) for the profilin-like PA complex was de-rived from the dependence of the elongationrate inhibition on the protein concentrationwith the same equation used for the seques-tering protein (Equation 13.6.8).

Profilin-like proteins fail to prevent fullyelongation at the barbed end of the filament be-cause the PA complex productively associateswith the barbed end. When the G-actin that ispresent in the growth assay is saturated by theprotein, the rate constant for association of PAto the barbed end is derived from the growthrate measurements. For profilin, the value ofk+ is 30% lower than that for G-actin (Fig.13.6.3D).

Capping proteinsIn measurements of F-actin assembly at

steady state, when capping protein is added

Figure 13.6.4 (at right) Functional assays of capping proteins. (A) Steady state measurements ofF-actin in the presence of increasing amounts of capping protein. The experiment was conductedas in Figure 13.6.2A. When barbed ends are free (solid line), capping proteins depolymerize F-actin, partially blocking all barbed ends; the maximum amount of unpolymerized actin equals thecritical concentration of the pointed end (Cc = 0.6 µM, under physiological conditions). Whenbarbed ends are capped (dashed line), addition of another capping protein has no effect on thesteady-state amount of F-actin; all barbed ends remain capped and Css remains equal to the Cc

at the pointed ends. (B) Css measurements for actin alone or in the presence of capping proteins.The experiment was conducted as in Figure 13.6.2C. When barbed ends are capped, cappingproteins have no effect on Cc. When barbed ends are free, capping proteins shift Cc to 0.6 µM,in physiological conditions. Solid line, free barbed ends alone; dotted line, free barbed ends withcapping protein; dashed line, capped barbed ends alone or with capping protein. (C) Cappingproteins do not inhibit pointed-end growth of actin filaments. The experiment was conducted as inFigure 13.6.2E. (D) Capping proteins inhibit barbed-end growth of actin filaments. The experimentwas conducted as in Figure 13.6.2F. (E) Depolymerization rate measurements. The experimentwas conducted as in Figure 13.6.2D. Capping proteins block depolymerization at the barbed end.Dotted line, no capping protein; solid lines, capping protein present at indicated concentrations.(F) Co-sedimentation assay. The experiment was conducted as in Figure 13.6.2G. At a saturatingamount of capping protein, the amount of G-actin in the supernatant corresponds to the Cc at thepointed end. Lines shown in A-E represent theoretical lines in each case.

OrganelleMotility

13.6.21


to a solution of F-actin with free barbed ends,partial depolymerization occurs until the crit-ical concentration of the pointed ends (CP

c ) isreached. The amount of F-actin that disassem-bles therefore equals the difference CP

c − CBc ,

at any F-actin concentration. Under physiolog-ical ionic conditions, this amount is 0.6 µM− 0.1 µM = 0.5 µM (Fig. 13.6.4A,B). Whenbarbed ends are capped by a standard cappingprotein like gelsolin, addition of another cap-ping protein does not cause further depolymer-

ization of F-actin. The same conclusions arederived from Cc plots (Fig. 13.6.4B).

In co-sedimentation assays, capping pro-teins cause limited depolymerization of0.5 µM F-actin because of the establishmentof CP

c . Capping proteins co-sediment with F-actin in a largely substoichiometric molar ratiowith F-actin, but they are detectable by westernblotting (Fig. 13.6.4F).

In seeded growth assays, capping proteinsinhibit elongation at the barbed end, generally



Assembly

13.6.22


with high affinity (nanomolar range) but notat the pointed end (Fig. 13.6.4C,D). Barbed-end growth inhibition can be described by thefollowing equation:

Equation 13.6.9

where V0 is the rate of elongation in the ab-sence of the capping protein, V∞ is the rateof elongation in the presence of a saturatingamount of capping protein (corresponding tothe rate of elongation of the pointed end), Kc isthe equilibrium constant between the cappingprotein and actin, and C is the concentration ofcapping protein.

These proteins block dilution-induced de-polymerization at the barbed end (Fig.13.6.4E). The depolymerization rates areinhibited when the barbed ends are free. Theequilibrium dissociation constant (20 nM forgelsolin) for the capping protein–actin com-plex can be derived from the dependence ofthe rate of dilution-induced depolymerizationon the protein concentration using Equation13.6.9.

Particular class of leaky cappersThe above properties characterize strong

cappers that completely block reactions of bothactin association to and dissociation from thebarbed ends. Proteins that bind to barbed endsmay still allow either growth only or depoly-merization only, or may allow both reactionswhile remaining bound to the end, with alteredkinetic parameters. Recent evidence for theseactivities has been reported for a new class ofactin-binding proteins called formins (Pruyneet al., 2002; Wallar and Alberts, 2003). Theseproteins nucleate barbed-end actin assemblyand can remain bound to the barbed end as itgrows, thus mediating insertional polymeriza-tion under certain conditions. The steady statemeasurements and kinetic measurements de-scribed above allow the quantification of ef-fects of formins on actin dynamics.

ADF/cofilin proteinsIn steady-state measurements of actin

assembly, ADF causes partial (limited)depolymerization; however, its behavioris different from a capper because partialdepolymerization is observed in the presenceas well as in the absence of capping proteins.Co-sedimentation assays show evidence forbinding of ADF to F-actin, but the partial

depolymerization implies that ADF alsobinds to G-actin. Binding studies show thatADF binds preferentially to ADP-actin andkinetic data indicate that ADF slows downnucleotide exchange, implying that at steadystate a large fraction of G-actin consists ofADF-ADP-G-actin. Kinetic measurements ofdilution-induced depolymerization provideevidence for faster depolymerization fromthe pointed ends, which is confirmed bymeasurements of filament turnover. Takentogether, these observations help to explainthe function of ADF/cofilin proteins. (for areview, Carlier et al., 1999)

Time ConsiderationsPurification of G-actin requires 6 days

(Support Protocol 1). Labeling actin witheither pyrenyl or 7-chloro-4-nitrobenzeno-2-oxa-1,3-diazole (NBD-Cl) requires 3 days(Support Protocols 2 and 3). Basic Protocols1, 2, 3, 5, and 6 each take half of a day. BasicProtocols 4 and 7 take 2 days each.

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Carlier, M.F., Ressad, F., and Pantaloni, D. 1999.Control of actin dynamics in cell motility.Role of ADF/cofilin. J. Biol. Chem. 274:33827-33830.

Casella, J.F., Maack, D.J., and Lin, S. 1986. Purifi-cation and initial characterization of a proteinfrom skeletal muscle that caps the barbed endsof actin filaments. J. Biol. Chem. 261:10915-10921.

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Combeau, C. and Carlier, M.F. 1989. Characteri-zation of the aluminum and beryllium fluoridespecies bound to F-actin and microtubules at thesite of the gamma-phosphate of the nucleotide.J. Biol. Chem. 264:19017-19021.

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Didry, D., Carlier, M.F., and Pantaloni, D. 1998.Synergy between actin depolymerizing fac-tor/cofilin and profilin in increasing actin fila-ment turnover. J. Biol. Chem. 273:25602-25611.

Egile, C., Loisel, T.P., Laurent, V., Li, R., Pantaloni,D., Sansonetti, P.J., and Carlier, M.F. 1999. Ac-tivation of the CDC42 effector N-WASP by theShigella flexneri IcsA protein promotes actin nu-cleation by Arp2/3 complex and bacterial actin-based motility. J. Cell Biol. 146:1319-1332.

Hug, C., Jay, P.Y., Reddy, I., McNally, J.G., Bridg-man, P.C., Elson, E.L., and Cooper, J.A. 1995.Capping protein levels influence actin assemblyand cell motility in dictyostelium. Cell 81:591-600.

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Valentin-Ranc, C. and Carlier, M.F. 1989. Evidencefor the direct interaction between tightly bounddivalent metal ion and ATP on actin. Bindingof the lambda isomers of beta gamma-bidentateCrATP to actin. J. Biol. Chem. 264:20871-20880.

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Contributed by Maud HertzogIstituto FIRC di Oncologia MolecolareFondazione Italiana per la Ricerca sulCancroMilano, Italy

Marie-France CarlierLaboratoire d’Enzymologie et BiochimieStructurale Centre National de laRecherche ScientifiqueGif-sur-Yvette, France

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