IntroductionWe often think of actin filaments as dynamic polymers thatassemble very rapidly to provide the cytoskeleton necessary forcell extension or movement and then depolymerize almost asabruptly. Examples of this type of activity include theextension of filopodia, pseudopodia and lamellipodia inneurons, neutrophils, keratocytes and fibroblasts and theformation of the actin tail of Listeria (reviewed in Borisy andSvitkina, 2000; Pantaloni et al., 2001; Pollard et al., 2000).There are other systems, however, in which the actin filamentsare much more stable. These include the thin filaments inskeletal muscle and filament bundles found in a variety ofsystems such as Drosophilabristles and nurse cells, microvilliof intestinal epithelial, stereocilia of the cochlea, Sertoliectocytoplasmic specializations and Limulus and Mytilussperm (reviewed in Bartles, 2000; DeRosier and Tilney, 2000).

We presume that the stability of the filaments is due to actinbinding and actin crosslinking proteins, as well as cappingproteins on both ends of the filaments. For example, thinfilament stability in muscle is due to the presence oftropomyosin, capZ and tropomodulin (reviewed in Littlefieldand Fowler, 1998). Filament stability in parallel filamentbundles is due to crosslinking proteins, for example, scruin inLimulus sperm (Sanders et al., 1996), fimbrin and espin instereocilia (Bartles, 2000; Tilney et al., 1989), fascin andforked in bristles (Tilney et al., 1995; Tilney et al., 1998), villinand fascin in nurse cells (Cant et al., 1994; Mahajan-Miklosand Cooley, 1994) and fimbrin, espin and villin in gut

microvilli (Bartles et al., 1998; Bretscher, 1981; Bretscher andWeber, 1980; Glenney et al., 1981; Matsudaira and Burgess,1982).

Although actin filament turnover occurs in skeletal muscle(Littlefield et al., 2001; Michele et al., 1999) and microvilli(Stidwill et al., 1984), the bundles in these systems remainintact for very long periods. An extreme case is thestereociliary bundles in hair cells that remain intact for the lifeof the organism, which in humans can exceed 75 years. In fact,with the exception of the Drosophila bristles, stable bundlesnever disappear but instead the cells die before they completelydisassemble their bundles. This is also true of the intestinalepithelial cells, although the microvilli can shorten withchanges in diet (Weinman et al., 1989). In bristle cells, the actinbundles break down completely (Tilney et al., 1996; Tilney etal., 2000b), but the cell remains viable.

We have been studying actin bundles in forming Drosophilabristle cells for some time now. Bristles first sprout from thesurface of the thorax in 32-hour pupae and gradually elongate,reaching a mature length in 48-hour pupae (Lees and Picken,1944; Overton, 1967; Petersen et al., 1994; Tilney et al., 1996;Wulfkuhle et al., 1998). Close examination of the actin bundlesin mature-length bristles (Appel et al., 1993; Tilney et al.,1995) shows that the bundles are composed of repeating unitsattached end to end, units we call modules (Tilney et al., 1996).On average each module is 3 µm long. Surprisingly, the gapsbetween the modules of one bundle are often in transverseregister with the gaps in adjacent bundles. Furthermore, the

641

Drosophilabristle cells form enormous extensions that aresupported by equally impressive scaffolds of modular,polarized and crosslinked actin filament bundles. As thecell matures and support is taken over by the secretedcuticle, the actin scaffold is completely removed. Thisremoval begins during cell elongation and proceeds via anorderly series of steps that operate on each module. Usingconfocal and electron microscopy, we found that the ~500-filament modules are fractured longitudinally into 25-50-filament subbundles, indicating that module breakdown isthe reverse of assembly. Time-lapse confocal analysis ofGFP-decorated bundles in live cells showed that moduleswere shortened by subunit removal from filament barbedends, again indicating that module breakdown is thereverse of assembly. Module shortening takes place at afairly slow rate of ~1µm/hour, implying that maximally

crosslinked modules are not rapidly depolymerized.Barbed-end depolymerization was prevented withjasplakinolide and accelerated with cycloheximide,indicating that barbed-end maintenance requirescontinuous protein synthesis. Subbundle adhesion was lostin the presence of cytochalasin, indicating that continuousactin polymerization is required. Thus, these polarizedactin filament bundles are dynamic structures that requirecontinuous maintenance owing to protein and actinfilament turnover. We propose that after cell elongation,maintenance falls behind turnover, resulting in the removalof this modular cytoskeleton.

Key words: Cytoskeleton, Crosslink, Development, Cycloheximide,Cytochalasin

Summary

Actin filament turnover removes bundles fromDrosophila bristle cellsGregory M. Guild, Patricia S. Connelly, Kelly A. Vranich, Michael K. Shaw and Lewis G. TilneyDepartment of Biology, University of Pennsylvania, Philadelphia, PA 19104-6018, USAAuthor for correspondence (e-mail: gguild@sas.upenn.edu)

Accepted 26 October 2001Journal of Cell Science 115, 641-653 (2002) © The Company of Biologists Ltd

Research Article

642

basal end of each module is flat, whereas the tip is pointed orirregular in shape. Finally, the barbed ends of all the filamentsin each module are located at the tip of the module (that is atthe irregularly shaped end), and the pointed ends of thefilaments are located on the base or flat end of each module(Tilney et al., 1996).

During the period of bristle growth, cuticular material isgradually deposited outside the plasma membrane so that by40-42 hours a thin cuticle surrounds the entire bristle except atits growing tip. In thin sections of 54-hour pupae, the cuticlenow supports the bristle, and the modules in the actin bundleshave large gaps between them (Tilney et al., 1996). By 58hours, the bundles and all of the actin filaments, in at least somebristles, have disappeared (Tilney et al., 1996). By 60 hours allthe bristles have disappeared.

We have two motivations for studying actin bundlebreakdown. First, we would like to know how a celldepolymerizes crosslinked actin bundles. Second,understanding the depolymerization process may reveal moreabout bundle formation during bristle elongation. What wefound in these studies is in many ways unexpected. Thedisappearance of these crosslinked modules is a slow processand takes, on average, more than three hours to complete. Incontrast, the process of module formation takes 30 minutes orless (Tilney et al., 1996). Breakdown of the bundles proceedsby two processes: the longitudinal splitting of bundles intosubbundles and the disassembly of individual modules.Interestingly, these processes begin well before bristleelongation is complete even though actin assembly is thedriving force for bristle elongation (Tilney et al., 2000a). Theseobservations suggest that bundle breakdown is driven by actin-filament turnover. Furthermore, breakdown begins at thebarbed ends of the modules. Thus, actin filament breakdownin crosslinked bundles is different from that in other moredynamic structures like the actin network at the edge oflamellipodia whose filament turnover takes place at filamentpointed ends.

Materials and MethodsDrosophila stocks and developmental stagingThe Oregon-R strain of Drosophila melanogasterwas used as thewild-type in these studies. The hs-GFP-moesinstrain (Edwards et al.,1997; Kiehart et al., 2000) was generously provided by D. Kiehart(Duke University) and used to visualize actin filament bundles inliving bristle cells. The hgm2– insert was maintained over a SM6abalancer chromosome. Flies were maintained on standard cornmeal-molasses-yeast food at 25°C, 60-70% relative humidity, with a 12hour/12 hour day/night cycle. Complete descriptions of genes andsymbols can be found in Lindsley and Zimm (Lindsley and Zimm,1992) and on FlyBase (Consortium, 1999).

All animals were staged from the point of puparium formation, aneasily recognizable and brief stage lasting 30 minutes at the beginningof metamorphosis (Bainbridge and Bownes, 1981). White prepupaewere collected and placed on double-sided scotch tape in a Petri dishand returned to the 25°C incubator. At the appropriate time afterpuparium formation the pupae were dissected.

Dissection of pupae and culturing the dorsal thoracicepitheliumAfter removing the pupal case, we filleted the pupae as outlined indetail elsewhere (Tilney et al., 2000a). The isolated thoracic fillets

were placed in 60×15 mm petri plates with 5 ml of Grace’s medium(GIBCO-BRL, Life Technologies, Inc., Rockville, MD) containinginhibitor. These compounds were diluted from concentrated stocksinto culture medium just before use. Control incubations usingDMSO, methanol or ethanol alone (typically 0.1-0.2%) wereperformed in parallel. After swirling the medium in the petri dishesto dilute the compounds, we added the isolated thoraces, and theplates were returned to the 25°C incubator for the indicated times.At the end of this period thoraces were removed and fixed for lightor electron microscopy. The volume of medium used was justsufficient to cover the tissue and allow for good oxygen exchange.Stock solutions of cytochalasin D (Sigma Chemical Co., St. Louis,MO; 2 mM in methanol), cycloheximide (Sigma ChemicalCompany; 10 mg/ml in ethanol) and jasplakinolide (MolecularProbes Inc., Eugene, OR; 1 mM in DMSO) were stored in aliquotsat –20° C.

Fixation and processing for light and electron microscopyThe thoraces were fixed by immersion in 2% formaldehyde in PBS for5 minutes, then washed three times in 0.1% Triton X-100 in PBS for5 minutes each, then placed in 0.1% Triton X-100 containing 10–6 Mphalloidin conjugated to rhodamine (Sigma Chemical Co., St. Louis,MO) or Texas Red (Molecular Probes) at 4°C overnight in the dark.The next morning, the sample was placed on a slide with the ventralside downward, the excess fluid removed with filter paper and thethoraces mounted in glycerol Citifluor (Ted Pella Inc. Redding, CA).A coverslip was applied, excess Citifluor glycerol removed and thepreparation sealed with nail polish. Slides were examined with eithera Leica model TCS 4D confocal microscope or an Olympus Fluoviewmodel BX50 confocal microscope. Bristles were visualized by thefluorescence of the phalloidin-stained actin bundles. For eachprocedure, at least 40 bristles were measured. The methods we usedfor electron microscopy are detailed in Tilney et al. (Tilney et al.,1998).

Time-lapse confocal microscopyDissected dorsal thoraces from GFP-moesinpupae were prepared asdescribed above and cultured in Grace’s medium on a microscopeslide under a coverslip at room temperature (22-25°C). Images werecollected every 30 minutes using an Olympus Fluoview model BX50confocal microscope equipped with a 60× oil immersion 1.4 NAobjective. GFP fluorescence was easily detected using the 488-nmline of the argon laser set at its minimum intensity (6%). Since asingle bristle cell was examined during each observation period(typically 2-4 hours), the stage was not moved in the X-Y planebetween scans although some adjustment in the Z plane was oftennecessary. Each time point resulted in 3-5 optical sections, whichwere projected together using the Image J application(http://rsb.info.nih.gov/nih-image/). The resulting confocal imageswere processed using Adobe Photoshop (Adobe Systems, Inc.). Insome cases, a small amount of red fluorescent, carboxylate-modified0.02 µm microspheres (Molecular Probes) was added to the thoracesprior to culture. A fraction of the microspheres adsorbed to thebristles and could be later used as fiduciary marks in the confocalimages to align the time-lapse sequence in register. Image registrationwas also possible by alignment of fluorescent irregularities found onsome of the bundles.

ResultsActin bundles breakdown in an orderly series of stepsAlthough bristles do not reach their mature length until 48 hoursafter pupariation, changes in the actin bundles occur beforematurity, even though new modules are still being assembled at

Journal of Cell Science 115 (3)

643Actin bundle breakdown in Drosophila bristles

the bristle tip. These changes are orderly, progressive andcumulative, and they begin in pupae of only 40 hours of age. Inorder to quantify these features, we examined 195 phalloidin-stained bristles from pupae 32-56 hours after pupariation anddetermined the fraction of bristles at each time point that haddiscernible characteristics involving the actin bundles.

When we examined bristles early in the process ofelongation (e.g., 32-34 hours after puparium formation), wefound fluorescent bundles that extend from the base of thebristle to a location near their tip. Modules were difficult tofind. Instead the bundles were smooth and uninterrupted bygaps (Fig. 1a,b).

Before we found discrete gaps between modules, we foundbristles in which the tip of one module overlaps with the baseof the next module (Fig. 1c,d). Overlapping modules wereparticularly easy to detect when the bristle was artificially bentbefore fixation. Quantitatively, smooth bundles were the mainfeature of the youngest bundles (Fig. 1a,b), whereasoverlapping bundles were featured in significantly older butstill elongating bristles (Fig. 1c,d).

As bristles elongated further, discrete gaps appearedbetween adjacent modules, and the gaps in adjacent bundleswere often in transverse register (Fig. 1e,f). The time whengaps first appear (43 hours) is important because the size of thegap increased with developmental age (see below). We notethat there was weak rhodamine-phalloidin fluorescence in thegaps between modules, indicating the presence of a smallpopulation of actin filaments connecting adjacent modules. Inaddition, the lengths of the gaps in adjacent bundles oftenappear dissimilar.

As the bristle cell matured further, the size of the gapsincreased. A detailed examination of the gap itself revealedanother important feature. The basal end of each moduletended to end abruptly and to be rounded, if not flat. In contrast,the apical end of each module tapered to varying degrees orwas irregular (Fig. 1g,h). This tapering was prominent in 50-hour bristles (Fig. 1i,j). Since the gap between modulesincreased with pupal age as a result of the increasing taper ofthe apical end of each module, removal of subunits fromfilaments would logically occur from the tapered end nearestto the bristle tip from filament barbed ends.

Fig. 1.Extensive actin filament loss from bundle modules is seen inmaturing Drosophilabristles. Wild-type bristles of increasing agewere stained with rhodamine-labeled phalloidin and imaged byconfocal microscopy. Images of bristles at the indicated age (32-56hours after puparium formation) were examined for smooth bundleswith no gaps (a), bundles with overlapping modules (c), bundles withgaps (e), apically tapering modules (g) and for bundles withextensive filament ghosts remaining (i). The percentage of bristleswith bundles of each type was plotted as a function of developmentaltime. Note that individual bristles could contain more than one typeof bundle. For example, 43-hour bristles often contain bundles withoverlapping modules and bundles with gaps (see text for details). Atotal of 195 thoracic bristles were examined. Portions of bristlesillustrating smooth bundles (b), bundles with overlapping modules(indicated with arrowheads) (d), and bundles with gaps (indicatedwith arrowheads) (f) are shown to the right. A portion of a bristleillustrating tapering modules is shown in (h). Arrowheads indicatealignment of modules. A portion of a bristle illustrating extensivefilament ghosts is shown in (j). Arrowheads indicate two bundleswith ghost connections. Bars, 2 µm.

644

This conclusion was strengthened by data from whatappears to be the next step in module disassembly in whichthe apical ends of the modules lose most of their actin content,becoming tiny fluorescent threads that extend from the end ofthe taper toward the basal end of the module directly above it.These thin fluorescent ‘ghosts’ become prominent in 53-hourbristles (Fig. 1i,j).

Analysis of the bristle populations that exhibit moduleshortening allows us to estimate the rate of module shortening.First, over 50% of the 43-55-hour bristles exhibit full-lengthmodules defined by very short gaps on both ends (Fig. 1e,f).Thus, initiation of module breakdown occurs at an averagebristle age of 49 hours. Second, over 50% of 53-56-hourbristles contain module ghosts that are one module or more inlength (Fig. 1i,j). Thus, module breakdown is complete at anaverage bristle age of 54.5 hours. Consequently, an averagemodule of 3 µm in length (Tilney et al., 1996) can be removed

in 5.5 hours (54.5−49 hours) by barbed-end shortening at a rateof ~0.5 µm/hour (3 µm/5.5 hour). This rate is in reasonableagreement with the value determined more directly by time-lapse imaging (see below).

Electron microscopy provides a high-resolution view ofbundle disassemblyWe examined thin longitudinal sections of bristle cells byelectron microscopy to determine the details of bundledisassembly. The overlapping modules were most easily seenwhen the bristle was bent before fixation (Fig. 2a). At what weperceive to be the next stage in module shortening, we see gapsof increased length (Fig. 2b,c). Filament distribution appearedless uniform in the regions that would later become the gap asif there was depolymerization at one or both ends of themodules that previously overlapped. The gap then became

Journal of Cell Science 115 (3)

Fig. 2.Longitudinal thin sections through actin bundles in bristles of 54-hour pupae. These sections were selected because they show junctionsbetween adjacent modules. If this plate is rotated 90° so the viewer is looking across the bundles, the 12-nm period owing to the fascin crosslinkscan be easily recognized. This period is indicated by the bars in (b) and (c), but it is especially visible near the base of each micrograph. If thebristle is bent before fixation, the overlap between adjacent modules separates at the point of overlap (a). As the module shortens, the number offilaments found in the overlap region decreases (b). Shortly thereafter a distinct gap is seen between adjacent modules (c).

645Actin bundle breakdown in Drosophila bristles

apparent. Interestingly, the filaments on both ends of themodules showed the 12-nm transverse pattern (Fig. 2a-c)attributable to the fascin crosslink (Tilney et al., 1995),demonstrating that filaments in the bundles were maximallycrosslinked right up to the gap. These images reinforce ourcontention, on the basis of light microscopy, that subunit lossoccurred at the ragged end of each module, which is the endoriented toward the bristle tip.

Real-time visualization of barbed end depolymerizationIn order to determine independently the polarity of modulebreakdown, we used time-lapse confocal microscopy. Wevisualized the actin bundles in living cells by decoratingthem with GFP fused to the C-terminal end of Drosophila

moesin, a segment that contains an actin-binding domain(Edwards et al., 1997). Although this transgene is driven bythe hsp70 heat shock promoter, we found that sufficientamounts of GFP-moesin accumulated in bristle cells at 25°Cto allow for easy bundle detection by confocal microscopy.This transgene has no deleterious effects on the fly stock(Kiehart et al., 2000).

Thoraces from animals allowed to develop for 48-54 hoursafter white puparium formation were put into culture andvisualized by time-lapse confocal microscopy. These cultureconditions allow normal bristle development for a 6-7 hourperiod (Tilney et al., 2000a). We typically followed one bristlein each thorax for 2-4 hours, imaging it every 30 minutes. Theresulting sequential images were aligned using either non-moving irregularities in the bundles or by including fluorescent

Fig. 3. Time-lapse confocal microscopy of living bristle cells shows barbed-end loss from actin filaments. (a) Actin bundles decorated withGFP-moesin from a single bristle-cell extension were visualized over a 90-minute period. Images were taken at 30-minute intervals (0, 30,60 and 90 minutes) and are displayed from left to right. These images were aligned using a constellation of fluorescent irregularities on thebundles. The position of one of these bright spots is adjacent to the asterisk (*). Bristle orientation is indicated in the rightmost panel. Thetop region of these images is characterized by gaps in ~eight bundles that do not appear to widen during this time course. The upper andlower boundaries of one of these gaps are indicated by the closely spaced arrows for each time point. The bottom region of these images –near the bristle base – is characterized by widening gaps in ~eight bundles. The upper and lower boundaries of two of these gaps areindicated by the pair of arrows (left side of image) and the pair of double-headed arrows (right side of image). Note that the lowerboundaries of these gaps, defined by the barbed ends of the modules below, are moving away from the fixed upper boundaries. (b and c) Therate of barbed-end module shortening was determined as described in the text by measuring the length of five well defined gaps in the upperregion of the bristle (b) and in the lower region of the bristle (c). A least-squares fit for each lengthening gap generates a line with a sloperepresenting the rate of module shortening. All modules in the five non-widening gaps (b) showed negligible shortening: average rate of0.08±0.05 (SEM) µm/hour. All modules in the five widening gaps (c) showed similar shortening rates: average rate of 1.10±0.04 (SEM)µm/hour.

646

microspheres in the preparation that adsorbed to the bristlecuticle. An example of one aligned time-lapse series is shownin Figure 3. What is obvious from this and other sequences isthat there is differential breakdown of modules along the bristlelength. For example, some regions showed little if any gapwidening (Fig. 3, top arrows) whereas other regions exhibiteddistinct gap-widening activity (Fig. 3, bottom arrows) withinthis time window. Thus, there seems to be local control overmodule breakdown. This is apparent at two levels. First,laterally associated modules in local regions seem to bebreaking down together as seen in the two gapped regions seenin Figure 3a. And second, those modules that constitute the~20 µm of bundled actin between these two gapped regions arenot breaking down at all.

Careful examination of the aligned image sequence of thewidening gap activity (Fig. 3, bottom arrows) shows that thiswidening results from barbed-end module shortening. Theupper boundary of each gap corresponds to the basal end ofthe tip-ward module. These boundaries did not move duringthe 90-minute sequence, indicating that the filament pointedends of the module were stable. The lower boundary of eachbundle gap corresponds to the apical end of a module. Theseboundaries moved toward the base of the bristle during thetime-lapse sequence, indicating that shortening of the modulesoccurred from the barbed ends of the actin filaments. Thus,using two independent approaches, length analysis of modulepopulations from static images taken during the course ofbundle breakdown (Fig. 1) and real-time analysis of individualmodules (Fig. 3) lead to the same conclusion – modularbundles of actin filaments breakdown by subunit loss from theirbarbed ends.

We used these time-lapse images to directly measure the

rate of module shortening. We found that the module base(filament pointed ends) that defined the upper boundary ofeach gap remained stationary relative to internal fiduciarymarks (Fig. 3, asterisk) during the time-lapse series.Accordingly, we measured the distance from this boundaryto the apical end (filament barbed ends) of the retreatingmodule in the gap at each time point. For each gapped bundleshown in Figure 3, we plotted increasing gap length (as ameasure of module shortening) as a function of time andused the least squares method to calculate the linearregression equation to fit the data. The slope of the resultingline represents the rate of module shortening. The averagerate of shortening exhibited by the widening gaps shown inFigure 3 (lower portion) was 1.10±0.04 µm/hour (Fig. 3c).Module shortening was almost undetectable in non-wideninggaps (Fig. 3b). Thus, the rate of barbed-end shorteningdetermined by time-lapse microscopy of individualmodules (1.1 µm/hour) is in reasonable agreement with thevalue (0.5 µm/hour) determined by analyzing static images

Journal of Cell Science 115 (3)

Fig. 4.Jasplakinolide prevents module shortening. Thoraces from48-hour animals were cultured for 5 hours in the presence or in theabsence of jasplakinolide (3 µM), stained with rhodamine-labeledphalloidin and imaged by confocal microscopy. (a) Bundles incontrol bristles develop the expected gaps owing to actindepolymerization. (b) In contrast, bundles in jasplakinolide-treatedbristles failed to undergo normal barbed-end depolymerization andremained smooth.

0

25

50

75

100

Control Cycloheximide

% B

ristle

s w

ith t

aper

ing

m

odul

es o

r gh

osts b

0

10

20

30

40

Control Cycloheximide

Leng

th

(µm

)

a

Fig. 5. Cycloheximide does not affect bristle growth but doespromote premature bundle breakdown. (a) A group of same-agethoraces (dissected from a single cohort of pupae 36 hours afterpuparium formation) were divided into two groups and cultured inthe presence or in the absence (control) of 1 µM cycloheximide for 5hours. Bristle lengths were measured from confocal images ofphalloidin-stained microchaetes. Bristle length was unaffected whencultured in cycloheximide. (b) A group of same-age thoraces(dissected from a single cohort of pupae 42 hours after pupariumformation) were divided into two groups and cultured in the presenceor in the absence (control) of 1 µM cycloheximide for two hours.Actin bundles were imaged by confocal microscopy and evaluated bythe criteria illustrated in Fig. 1. A total of 56 bristles were examinedand the percentages of bristles exhibiting tapering modules or bundleghosts characteristic of depolymerizing bundles are presented. Ofinterest here is that cycloheximide promotes the breakdown ofmodules usually seen in older bristles (e.g., Fig. 1).

647Actin bundle breakdown in Drosophila bristles

of module populations during the course of bundlebreakdown.

Inhibition of actin depolymerization prevents moduleshorteningJasplakinolide is a compound capable of stabilizing actinfilaments by both lowering the off-rate and increasing the on-rate of actin subunits at the barbed ends (Bubb et al., 1994;Bubb et al., 2000). In fact, growing bristles cultured in thepresence of jasplakinolide exhibit accelerated elongation rates,indicating that actin polymerization drives bristle-cellelongation (Tilney et al., 2000b). We reasoned that if actindepolymerization was responsible for module shortening thenjasplakinolide could prevent this. To test this idea, we culturedbristles from 48 hour pupae for 5 hours in the presenceor absence of jasplakinolide and evaluated actin bundlemorphology by confocal microscopy. As expected, bristlesincubated in the absence of jasplakinolide developed bundlesthat contained gaps and tapers typical of bristles of this age(Fig. 4a). In contrast, bristles cultured in the presence ofjasplakinolide failed to undergo module breakdown andexhibited smooth non-gapped bundles typical of much youngerbristles (Fig. 4b).

Inhibition of protein synthesis induces prematuredepolymerization of the actin bundlesIsolated thoraces from 36-hour pupae were cultured for 5hours in the presence of the protein-synthesis inhibitorcycloheximide. Bristle elongation was unaffected bycycloheximide (Fig. 5a). By confocal microscopy, we saw theexpected overlapping modules and, in one case, gaps in thebundles. However, the cross-sectional area of the bundles wasapproximately four-fold smaller than the controls. Even so, thefilaments in these small bundles were hexagonally packed (Fig.6) and maximally crosslinked.

In contrast, pupae cultured with cycloheximide from 42-44hours showed many examples of gap formation andpremature bundle breakdown. Almost 75% of the bundlesshowed tapering modules and even ghosts (Fig. 5b), whichwere not normally seen in the controls until later indevelopment, typically 50-56 hours (e.g., Fig. 1g-j). In thinsections, we saw bundles breaking apart into subbundles (notshown), again a stage normally seen in 48-53-hour bristlesbut not at this stage.

Modules separate into subbundles as breakdownproceedsIn longitudinal EM sections through modules of pupae of 54hours or more, we found that the once solid modules began toseparate longitudinally into a series of thinner subbundles (Fig.7d,e). The size of the subbundles varied, but in many casesthere were six or seven filaments in longitudinal sections,indicating that if these subbundles were round they would becomposed of 25-50 filaments. The filaments in each of thesubbundles displayed the 12-nm period attributable to thefascin crosslink (Fig. 7c), and in transverse section thefilaments in the subbundles were hexagonally packed (Fig.7a,b). This means that the filaments in the subbundles are

maximally crosslinked together. As the pupae aged, many ofthe subbundles separated from those still attached to the plasmamembrane (Fig. 7a,b). These were often found deep within thecytoplasm of the cell.

We also studied the stages of subbundle formation. Thesplits in the bundle appeared first at the frayed apical end of amodule (Fig. 7c) not the flat basal end. Later these splits ranthe length of the bundle proper (Fig. 7b). This invariably cutthe bundles into two pieces. The smaller piece remainedconnected to the plasma membrane while the other piece,containing about two-thirds of the filaments, lay deeper in thecytoplasm (Fig. 7b,d). Over time, these large subbundlessplit further into smaller subbundles composed of 25-50hexagonally packed crosslinked filaments.

The formation of the subbundles could also be followed intransverse sections through modules. The first step insubbundle formation seemed to be the appearance of holes orareas in the large bundles that lack filaments (Fig. 8). Thesebundles almost look like the end of a pallet of lumber fromwhich several internal planks were removed. Most often, theholes first appeared five to six rows inwards from the plasmamembrane.

Cytochalasin induces precocious bundle disassemblyIsolated thoraces cultured in Grace’s insect medium elongatebristles at normal rates over a 6-7-hour period (Tilney et al.,2000a). Bristle elongation stops abruptly when thoracesisolated from 36-hour or 41-hour old pupae are cultured in the

Fig. 6. Transverse thin section through a macrochaete from a thoraxof a 36-hour-old pupa that was cultured in the presence ofcycloheximide for 5 hours before fixation. Of interest is the actinbundles indicated by the arrows as they are small yet the filaments inthe bundles are hexagonally packed (see insert) and maximallycrosslinked by fascin.

648 Journal of Cell Science 115 (3)

Fig. 7.Solid modules separate longitudinally into a series of subbundles. All of these figures come from 54- and 55-hour old pupae. Panels(a and b) show thin transverse sections through a bristle. The chitinous exoskeleton of the bristle is being deposited outside the plasmamembrane surrounding the bristle. Located at the top of panel (a) are three bundles (arrows) illustrated at higher magnification in (b). Eachbundle has separated into two subbundles. The dots in the bundles are the actin filaments cut in cross section. Panel (c) shows a section througha module with a flat basal end and a pointed apical end that is splitting into two subbundles. The 12-nm period attributable to the cross-bridgefascin is indicated on this micrograph by the three bars. This period is best seen by looking across the bundle from its side. Panel (d) shows adeeper split of a bundle into two subbundles. Panel (e) shows two tandem modules splitting into a series of subbundles. At least one subbundleremains associated with the plasma membrane.

649Actin bundle breakdown in Drosophila bristles

presence of cytochalasin D. However, thebundles that were formed prior totreatment remain (Tilney et al., 2000a).These bundles fragment into subbundlesbecause there are many more bundlesthan in the controls (Fig. 9a,b). Thesesubbundles are correspondingly thinnerthan the bundles observed in controls. Wealso found a population of randomlyoriented small fluorescent units at thebase of the bristle shaft and in thecytoplasm around the nucleus and Golgiof the bristle cell (Fig. 9b,c). On average,each of these units was 3 µm, the averagelength of the bristle modules (Tilney etal., 1996). It looks as if the bundles fromthe basal end of the bristle have not onlyseparated into the subbundles but thesesubbundles have also moved into the cellbody.

In thin sections through the bristleshaft we can view this process at higherresolution. After only two hours incytochalasin, we found, instead of 7-11large bundles in a microchaete, 27 smallbundles located near the cell surface aswell as a few internally located bundles(data not shown). Each bundle is thinnerthan the 7-11 seen in the wild-type at thesame stage, and some of these were in theprocess of splitting into even smallersubbundles.

In our thin sections through the base of the bristle cell inthe thorax proper, in the plane that includes the nucleus andthe Golgi apparatus, we could resolve the randomly oriented~3 µm long bundles (Fig. 10) seen previously by confocalmicroscopy. Each of these bundles was composed ofapproximately 50 filaments rather than the 600-1000

filaments that typically make up each of the 7-11 bundlespresent in the shaft of fully elongated bristles. In essence,cytochalasin promotes the appearance of the subbundles thatnormally associate laterally to form each module. Of interestto our subsequent discussion is the observation that the 12-nm transverse period is readily apparent in longitudinal

Fig. 8. Transverse sections through a microchaete from a 54-hour pupa. This section illustratesthe first step in subbundle formation where holes or areas lacking in filaments appear withinthe actin bundle (e.g., arrow in bundle at 7 o’clock).

Fig. 9.Cytochalasin promotes the premature breakdown of modules into subbundles. Macrochaetes from wild-type thoraces cultured in thepresence of cytochalasin (35-42 hours) were fixed, stained with rhodamine-labeled phalloidin and imaged by confocal microscopy. Panel (b)shows an increased number of bundles in the bristle shaft (arrowhead). The modules seem to dissolve into subbundles at the base of the shaft(arrow) and accumulate in the cell body. Panel (a) shows a higher magnification of the extra bundles present in a different bristle. Panel (c)shows another bristle and what appear to be subbundles pouring off the end of the bristle shaft (arrow) and into the cell body. Bars, 5 µm.

650

sections of these subbundles (Fig. 10b), indicating that thefilaments in these subbundles remain maximally crosslinked.It is also interesting that bristle cells incubated incytochalasin do not form gaps prematurely. Thus,cytochalasin does not effect the rate of depolymerization ofthe bundles from their barbed ends. However, cytochalasininduces a premature splitting of the modules into subbundles,reminiscent of the process that occurs in bristles from pupae50 hours old or older. Also, at the base of the bristle, thesubbundles from the modules become randomly oriented inthe cytoplasm of the cell body.

DiscussionThe actin bundle in developing Drosophila bristle cells is auseful model for studying the in vivo biology of a functionalcytoskeletal unit composed of many proteins acting inconcert. In earlier studies, we showed that although actin

assembly is essential for bristle elongation (Tilney et al.,2000a), once the bristle elongates to its mature length theactin filament bundles disappear and the cell shape ismaintained by the chitinous exoskeleton (Tilney et al., 1996).We show here that modules breakdown in two ways. Thefilaments shorten from their barbed ends, and the modulessplit longitudinally into smaller subbundles (Fig. 11). Thisprocess literally represents bundle assembly in reverse, in thatactin is now lost from the barbed end of each filament andthe large maximally crosslinked bundles now splitlongitudinally into smaller subbundles. Interestingly, we findthat inhibitors of both actin filament assembly and proteinsynthesis cause bundles to breakdown prematurely. Further,breakdown is prevented if the filaments are stabilizedby jasplakinolide, a membrane-permeant phalloidin-likecompound. These results indicate that bundle breakdown isdriven by actin filament turnover.

Actin modules breakdown from their barbed endsWe find that even as actin polymerization and bundleformation at the bristle tip drives bristle elongation (Tilney etal., 2000a; Tilney et al., 2000b), transverse gaps develop in theolder portions of these same bundles near the base of thebristle. Once the bristle reaches its mature length (48-hourpupae), these gaps increase in size until all the F-actindisappears (60-hour pupae). Thus, actin filament loss occursover many hours.

We conclude that the subunits are being selectively removedfrom the apical ends of the modules for three reasons. First, the

Journal of Cell Science 115 (3)

Fig. 10. A thin section through the base of the bristle cell from anisolated thorax that had been treated for two hours with cytochalasinD before fixation. The thorax was dissected from a 41-hour pupa. By43 hours, bundle disassembly in the controls had not yet occurred. Inthe cytoplasm surrounding the nucleus (N) are numerous smallbundles (a) each containing about 50 filaments. One of the bundles isdepicted at high magnification in (b). The lines indicate the 12-nmperiod attributable to the crosslink fascin.

a b c d e bristle base

>>>>>>>>>>>>>>> >>>>>>>

bristle tip

>>>>>>>>>>>>>>>>>>>>>>>>> >>>>>>>>>>>>

barbed

pointed

Fig. 11.Model for the breakdown of the modular actin bundles thatsupport Drosophilabristles. (a) Two adjacent modules (gray boxes)are shown attached to the plasma membrane (vertical black line) ofthe cell. The orientation of polarized actin filaments in the modulesand the orientation of the modules relative to the bristle tip are shownto the right. In these panels, only the lower module undergoesbreakdown, whereas the upper module remains intact forcomparison. (b) Actin filaments shorten by the loss of subunits fromtheir barbed ends. This process can be accelerated by inhibitingprotein synthesis with cycloheximide. (c) The module also begins tobreakdown into submodules by longitudinal cleavages. This processcan be accelerated by inhibiting actin polymerization withcytochalasin. It is not clear whether depolymerization or subbundleformation is initiated sequentially or simultaneously. (d) The moduleis completely split from the plasma membrane, leaving behind amembrane-bound actin ‘ghost’. Some submodules can becomecompletely split off from the module proper and can be found deeperin the cytoplasm. (e) In the end, only the ‘ghost’ of the moduleremains attached to the membrane.

651Actin bundle breakdown in Drosophila bristles

basal end of each module is nearly flat and maintains its shapeat all stages. In contrast, by confocal microscopy, the apical endis ragged with tiny threads of F actin extending distally. Second,the basal ends seem to hold their relative positions in the bristleduring development, as evidenced by the tendency of these endsto remain in transverse register. In contrast, the apical ends ofadjacent modules can be highly variable in length. These apicalends shrink toward the flat end of the module, leaving behindtheir subbundle ghost still attached to the plasma membrane.Since the apical end corresponds to the barbed end of the actinfilaments (Tilney et al., 1996), subunits must be removed fromthis end and not from filament pointed ends at the module base.Third, we have directly monitored module breakdown in livingcells by time-lapse microscopy. It is clear from these images(Fig. 3) that actin subunits are lost from the filament barbedends. This conclusion agrees with our interpretation of ourconfocal ‘snapshots’ of modules in increasingly older bristles(Fig. 1). Furthermore, when jasplakinolide is used to preventdepolymerization, module shrinkage does not take place(Fig. 4).

Module breakdown is much slower than module synthesis.Although we know that the rate of bristle elongation increasesas bristles grow longer (Tilney et al., 2000a), we can estimatethe average rate of elongation of 70 µm microchaetes duringtheir 16 hour growth period to be at least 4.4 µm/hour and of250 µm macrochaetes to be at least 15.6 µm/hour. Since bristlegrowth is driven by actin polymerization (Tilney et al., 2000a),these are reasonable estimates for module growth as well.Careful analysis of module breakdown using time-lapsemicroscopy (Fig. 3c) indicates that module barbed-end shortening occurs at a rate of ~1 µm/hour. Thisdepolymerization rate is two orders of magnitude slower thanpointed-end depolymerization and actin turnover at the leadingedge of lamellipodia, which must match the rate of membraneprotrusion - up to 5-10 µm/min (Cassimeris et al., 1990;Svitkina and Borisy, 1999). This relatively slow rate of filamentbreakdown in bristle bundles could result from the actinfilaments being maximally crosslinked, or it could be becausethe ends of the filaments are capped or both. Interestingly, theactin bundles in microvilli of intestinal epithelial cells alsoshorten slowly in vivo, at a rate of ~0.2 µm/hour, when treatedwith lectins (Weinman et al., 1989), a rate comparable to whatwe observe in bristle modules.

Our conclusion that subunit loss from the barbed end of thefilaments in the bundles is particularly interesting as it conflictswith prevailing views of subunit loss in other systems, wheretreadmilling occurs by pointed-end loss dependent on‘depolymerizers’ like ADF/cofilin. Two things should be keptin mind. First, in bundles found in bristles, microvilli andstereocilia, the actin filaments are maximally crosslinkedtogether by two or more protein crosslinkers. These crosslinkscould presumably inhibit actin turnover. Second, the rate ofsubunit loss from these crosslinked bundles is orders ofmagnitude slower than the rate of subunit loss from actinmeshworks found in lamellipodia or in Listeria tails (e.g.,Cassimeris et al., 1990; Loisel et al., 1999; Svitkina and Borisy,1999; Theriot and Mitchison, 1991). Thus, how crosslinkedactin filaments depolymerize when they exist in bundles maybe very different from how dendritic arrays of actin filamentsand/or free actin filaments disassemble. In short, there is noconflict, and instead there are real differences.

Modules divide into subbundles during depolymerization– a process that reverses bundle formationWhen gaps appear between modules, individual modules beginto split longitudinally into subbundles containing 25-50filaments each. This splitting begins at the apical (barbed) endof each module where actin depolymerization is occurring.After splitting, some of the subbundles move to the center ofthe bristle. The filaments in the resulting subbundles are stillhexagonally packed and display the 12-nm period inlongitudinal section attributable to fascin, indicating that theyremain maximally crosslinked together.

The normal process of module breakdown into subbundles,a process that is accelerated by cytochalasin (an inhibitor ofactin polymerization) and cycloheximide (an inhibitor ofprotein synthesis), seems to represent a reversal of moduleassembly. For example, from studying the developmentalformation of modules, we know that they are formed fromsubbundles (Tilney et al., 1996) and that the forked crosslinkerplays a key role in assembling the subbundles into modules(Tilney et al., 1998). Thus, these subbundles are present in bothpupae, whose bristles are almost the mature length, and inyounger, still elongating bristles. Therefore, the subbundles arenot only a product of bundle breakdown but also represent astage in the formation of large bundles.

Bundle breakdown is really driven by actin filamentturnover There seems to be two different populations of filaments inthese actin bundles: the hexagonally packed filaments thatconstitute the subbundles plus the cytochalasin-sensitivefilaments that ‘glue’ the subbundles together. The subbundlefilaments seem to be very stable and only shorten late indevelopment, whereas the glue filaments are more dynamicbecause they are lost soon after cytochalasin treatment. It isinteresting to note that even the glue filaments contributesubstantially to bundle cross-sectional area. Modelingthe construction of a 1000-filament bundle requiresapproximately 750 subbundle filaments (30×25-filamentsubbundles) and approximately 250 glue filaments to holdeverything together.

Our inhibitor studies on growing bristles show that actinbundles are not static structures but require constantmaintenance to maintain their structure. For example, treatmentwith cytochalasin D blocked bristle elongation and led to thesplitting of the already-assembled bundles into smallersubbundles. However, the bundles still extended continuouslyfrom the base to the tip of each bristle, and no gaps were seen.Thus, the aggregation of the many smaller subbundles into the7-11 larger bundles requires continual actin filamentpolymerization and presumably filament crosslinking. Perhapsthe lateral assembly of subbundles into the larger bundles isimprecise, leading to subbundle interfaces that are notmaximally crosslinked with fascin and are subject tobreakdown. The resulting splits may be repaired by new actinfilament synthesis within the split and subsequent crosslinking.In fact, such imperfections in the hexagonal lattice can be seennormally during module development. This is presumably dueto the failure to zipper subbundles together perfectly during theearly phases of module formation. In contrast, inhibiting proteinsynthesis with cycloheximide does not block bristle elongation

652

but does cause the appearance of gaps within the bundles andthe breakdown of the modules from their apical barbed ends.

Although the precise details and molecular mechanisms ofsubunit removal remain to be determined, bundle breakdownseems to be the inevitable consequence of a failure to balancefilament breakdown with filament replacement that normallymaintains module and bundle integrity. A relevant analogy forthis is the ‘classic’ math problem of filling a bath without thedrain plug in place. As long as water enters the bath faster thanit can escape down the drain, the bath will fill. In the case ofactin polymerization, if filament bundling and the maintenanceprocesses exceed any breakdown processes, the bristle willelongate by actin polymerization. Once the volume of waterfilling the bath falls below the volume escaping, the bath willinevitably empty, the rate (and time) of emptying beingdetermined by the difference between the two processes. Weimagine that once the bristle becomes fully grown, theprocesses of actin polymerization, filament bundling andbundle maintenance are downregulated or stopped whilefilament breakdown continues or speeds up, resulting in thedisassembly of the crosslinked actin bundles.

We would like to express our thanks to Dan Kiehart for generouslymaking available the hs-GFP-moesinstock. This work was supportedby grants from the National Science Foundation (MCB-0077839) andthe University of Pennsylvania Research Foundation to G.M.G. andthe National Institute of Health (GM-52857) to L.G.T.

ReferencesAppel, L. F., Prout, M., Abu-Shumays, R., Hammonds, A., Garbe, J. C.,

Fristrom, D. and Fristrom, J. (1993). The DrosophilaStubble-stubbloidgene encodes an apparent transmembrane serine protease required forepithelial morphogenesis. Proc. Natl. Acad. Sci. USA90, 4937-4941.

Bainbridge, S. P. and Bownes, M.(1981). Staging the metamorphosis ofDrosophila melanogaster. J. Embryol. Exp. Morphol. 66, 57-80.

Bartles, J. R.(2000). Parallel actin bundles and their multiple actin-bundlingproteins. Curr. Opin. Cell Biol. 12, 72-78.

Bartles, J. R., Zheng, L., Li, A., Wierda, A. and Chen, B.(1998). Smallespin: a third actin-bundling protein and potential forked protein orthologin brush border microvilli. J. Cell Biol. 143, 107-119.

Borisy, G. G. and Svitkina, T. M. (2000). Actin machinery: pushing theenvelope. Curr. Opin. Cell Biol. 12, 104-112.

Bretscher, A. (1981). Fimbrin is a cytoskeletal protein that crosslinks F-actinin vitro. Proc. Natl. Acad. Sci. USA78, 6849-6853.

Bretscher, A. and Weber, K.(1980). Fimbrin, a new microfilament-associatedprotein present in microvilli and other cell surface structures. J. Cell Biol.86, 335-340.

Bubb, M. R., Senderowicz, A. M., Sausville, E. A., Duncan, K. L. andKorn, E. D. (1994). Jasplakinolide, a cytotoxic natural product, inducesactin polymerization and competitively inhibits the binding of phalloidin toF-actin. J. Biol. Chem. 269, 14869-14871.

Bubb, M. R., Spector, I., Beyer, B. B. and Fosen, K. M.(2000). Effects ofjasplakinolide on the kinetics of actin polymerization. An explanation forcertain in vivo observations. J. Biol. Chem. 275, 5163-5170.

Cant, K., Knowles, B. A., Mooseker, M. S. and Cooley, L.(1994).Drosophila singed, a fascin homolog, is required for actin bundle formationduring oogenesis and bristle extension. J. Cell Biol. 125, 369-380.

Cassimeris, L., McNeill, H. and Zigmond, S. H.(1990). Chemoattractant-stimulated polymorphonuclear leukocytes contain two populations of actinfilaments that differ in their spatial distributions and relative stabilities. J.Cell Biol.110, 1067-1075.

Consortium, F. (1999). The FlyBase Database of the Drosophila GenomeProjects and community literature. Nucleic Acids Res. 27, 85-88.

DeRosier, D. J. and Tilney, L. G.(2000). F-actin bundles are derivatives ofmicrovilli: What does this tell us about how bundles might form? J. CellBiol. 148, 1-6.

Edwards, K. A., Demsky, M., Montague, R. A., Weymouth, N. and

Kiehart, D. P. (1997). GFP-moesin illuminates actin cytoskeleton dynamicsin living tissue and demonstrates cell shape changes during morphogenesisin Drosophila. Dev. Biol. 191, 103-117.

Glenney, J. R. Jr., Kaulfus, P., Matsudaira, P. and Weber, K.(1981). F-actin binding and bundling properties of fimbrin, a major cytoskeletalprotein of microvillus core filaments. J. Biol. Chem. 256, 9283-9288.

Kiehart, D. P., Galbraith, C. G., Edwards, K. A., Rickoll, W. L. andMontague, R. A. (2000). Multiple forces contribute to cell sheetmorphogenesis for dorsal closure inDrosophila. J. Cell Biol. 149, 471-490.

Lees, A. D. and Picken, L. E. R.(1944). Shape in relation to fine structurein the bristles of Drosophila melanogaster. Proc. Roy. Soc. London. Ser. BBiol. Sci. 132, 396-423.

Lindsley, D. L. and Zimm, G. G. (1992). The Genome of Drosophilamelanogaster. San Diego: Academic Press, Inc.

Littlefield, R., Almenar-Queralt, A. and Fowler, V. M. (2001). Actindynamics at pointed ends regulates thin filament length in striated muscle.Nat. Cell Biol. 3, 544-551.

Littlefield, R. and Fowler, V. M. (1998). Defining actin filament length instriated muscle: rulers and caps or dynamic stability? Annu. Rev. Cell Dev.Biol. 14, 487-525.

Loisel, T. P., Boujemaa, R., Pantaloni, D. and Carlier, M. F.(1999).Reconstitution of actin-based motility of Listeria and Shigellausing pureproteins. Nature401, 613-616.

Mahajan-Miklos, S. and Cooley, L.(1994). The villin-like protein encodedby the Drosophila quailgene is required for actin bundle assembly duringoogenesis. Cell 78, 291-301.

Matsudaira, P. T. and Burgess, D. R.(1982). Partial reconstruction of themicrovillus core bundle: characterization of villin as a Ca++-dependent,actin-bundling/depolymerizing protein. J. Cell Biol. 92, 648-656.

Michele, D. E., Albayya, F. P. and Metzger, J. M.(1999). Thin filamentprotein dynamics in fully differentiated adult cardiac myocytes: toward amodel of sarcomere maintenance. J. Cell Biol. 145, 1483-1495.

Overton, J. (1967). The fine structure of developing bristles in wild type andmutant Drosophila melanogaster. J. Morphol. 122, 367-379.

Pantaloni, D., Le Clainche, C. and Carlier, M. F.(2001). Mechanism ofactin-based motility. Science292, 1502-1506.

Petersen, N. S., Lankenau, D. H., Mitchell, H. K., Young, P. and Corces,V. G. (1994). Forked proteins are components of fiber bundles present indeveloping bristles of Drosophila melanogaster. Genetics136, 173-182.

Pollard, T. D., Blanchoin, L. and Mullins, R. D. (2000). Molecularmechanisms controlling actin filament dynamics in nonmuscle cells. Annu.Rev. Biophys. Biomol. Struct. 29, 545-576.

Sanders, M. C., Way, M., Sakai, J. and Matsudaira, P.(1996).Characterization of the actin cross-linking properties of the scruin-calmodulin complex from the acrosomal process of Limulussperm. J. Biol.Chem. 271, 2651-2657.

Stidwill, R. P., Wysolmerski, T. and Burgess, D. R.(1984). The brush bordercytoskeleton is not static: in vivo turnover of proteins. J. Cell Biol. 98, 641-645.

Svitkina, T. M. and Borisy, G. G. (1999). Arp2/3 complex and actindepolymerizing factor/cofilin in dendritic organization and treadmilling ofactin filament array in lamellipodia. J. Cell Biol. 145, 1009-1026.

Theriot, J. A. and Mitchison, T. J. (1991). Actin microfilament dynamics inlocomoting cells. Nature352, 126-131.

Tilney, L. G., Tilney, M. S. and Guild, G. M. (1995). F actin bundles inDrosophila bristles. I. Two filament cross-links are involved in bundling. J.Cell Biol. 130, 629-638.

Tilney, L. G., Connelly, P., Smith, S. and Guild, G. M.(1996). F-actinbundles in Drosophila bristles are assembled from modules composed ofshort filaments. J. Cell Biol. 135, 1291-1308.

Tilney, L. G., Connelly, P. S., Vranich, K. A., Shaw, M. K. and Guild, G.M. (1998). Why are two different cross-linkers necessary for actin bundleformation in vivo and what does each cross-link contribute? J. Cell Biol.143, 121-133.

Tilney, L. G., Connelly, P. S., Vranich, K. A., Shaw, M. K. and Guild, G.M. (2000a). Actin filaments and microtubules play different roles duringbristle elongation in Drosophila. J. Cell Sci. 113, 1255-1265.

Tilney, L. G., Connelly, P. S., Vranich, K. A., Shaw, M. K. and Guild, G.M. (2000b). Regulation of actin filament cross-linking and bundle shape inDrosophilabristles. J. Cell Biol. 148, 87-100.

Tilney, M. S., Tilney, L. G., Stephens, R. E., Merte, C., Drenckhahn, D.,Cotanche, D. A. and Bretscher, A.(1989). Preliminary biochemicalcharacterization of the stereocilia and cuticular plate of hair cells of the chickcochlea. J. Cell Biol. 109, 1711-1723.

Journal of Cell Science 115 (3)

653Actin bundle breakdown in Drosophila bristles

Weinman, M. D., Allan, C. H., Trier, J. S. and Hagen, S. J.(1989). Repairof microvilli in the rat small intestine after damage with lectins containedin the red kidney bean. Gastroenterology97, 1193-1204.

Wulfkuhle, J. D., Petersen, N. S. and Otto, J. J.(1998). Changes in the F-actin cytoskeleton during neurosensory bristle development in Drosophila:the role of singed and forked proteins. Cell Motil. Cytoskeleton40, 119-132.