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Vol. 17, Issue 9, 3930-3939, September 2006
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*Department of Biology, Washington University, St. Louis, MO 63130; and Laboratory of Developmental Biology, Institute of General and Molecular Biology, Nicolaus Copernicus University, 87-100 Torun, Poland
Submitted September 21, 2005;
Accepted June 27, 2006
Monitoring Editor: David Drubin
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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). On activation by a member of the Wiscott Aldrich Syndrome protein (WASp) family, this complex can nucleate new actin filament assembly as well as create branched networks of filaments through side binding to existing filaments (Egile et al., 1999; Machesky et al., 1999; Rohatgi et al., 1999; Winter et al., 1999a; Yarar et al., 1999). In addition to its actin nucleation activity, the Arp2/3 complex can cap the slow growing, or pointed, ends of actin filaments (Mullins et al., 1998). Capping protein, a heterodimer of alpha and beta subunits, binds the fast growing, or barbed, ends of actin filaments and prevents both depolymerization and the addition of new monomers (Isenberg et al., 1980). The importance of capping protein and the Arp2/3 complex is evidenced by the fact that they are two of the five components absolutely required for formation of actin comet tails that drive intracellular movement of the pathogenic bacteria Listeria monocytogenes (Loisel et al., 1999).
Although much has been learned in the last several years about the proteins and mechanisms regulating actin filament assembly during motility and in vitro (for review see Pollard and Borisy, 2003), much less is understood about the processes governing the assembly of more stable actin structures in cells that do not move. Many questions remain unanswered about such structures. For example, how are stable actin structures positioned within the cell? Do the same proteins control filamentous actin assembly in stationary cells as in the leading edge of moving cells? Do these proteins function in the same way in these very different contexts?
To investigate these questions, we are using the large bristles, or macrochaetes, of the fruit fly Drosophila melanogaster. The long bristle shaft (up to 400 µm in length) is generated from a single cell that undergoes extension during pupal development (Lees and Picken, 1944; Lees and Waddington, 1944). Bristle cell elongation is dependent on actin assembly (Tilney et al., 2000), and bundles of actin filaments lie against the membrane, parallel to the long axis of the bristle (Overton, 1967; Appel et al., 1993). These cortical bundles give the adult bristle its grooved appearance; the bundles lie at the positions of the valleys between the tall ridges of cuticle. Although it is apparent that the barbed ends of the actin filaments in bundles are oriented toward the bristle tips (Tilney et al., 1996), it is not clear that extension of actin filaments in the bundles provides the pushing force of membrane protrusion. Rather, the bundles might serve as structural girders that maintain the cellular extension. Between the bundles are very dynamic masses of actin filaments termed snarls (Tilney et al., 1996, 2003). Snarls were proposed to be transient structures that are not stabilized by cross-linking to other filaments or attachment to the membrane (Tilney et al., 2003), perhaps byproducts or aberrantly formed structures that are eliminated. Whether snarls have any function is unknown.
Mutations in genes encoding many actin-regulating proteins, such as the cross-linkers forked and singed (encoding fascin; Tilney et al., 1995), chickadee (encoding profilin, an actin monomer-binding protein that promotes filament assembly; Verheyen and Cooley, 1994), the actin monomer sequesterer twinfilin (Wahlstrom et al., 2001), and capping protein (Hopmann et al., 1996), result in deformed bristles. Close examination of bristle development reveals aberrant actin bundle morphologies in these mutants. For example, capping protein beta mutant pupal bristles have irregularly sized actin bundles that are displaced from the membrane. We hypothesized that this may be due to unregulated assembly of the filaments within the actin bundles, though the precise mechanism remained unclear (Hopmann et al., 1996). In previous work we have used the bristle system to elucidate the antagonistic interaction between capping protein and profilin: we found that a chickadee mutation was able to suppress the bristle phenotype of a transheterozygous combination of capping protein beta mutant alleles (Hopmann and Miller, 2003). Reduction of capping protein leads to an increase in actin filament assembly that is abrogated by loss of profilin. Thus these proteins appear to act in a very similar manner to their function at the leading edge of motile cells (for review see Pollard and Borisy, 2003).
In this article we report that, as predicted from studies in motile cells, capping protein and the Arp2/3 complex act antagonistically to one another during bristle development. However, we show that the role of capping protein in actin bundle organization is quite different from previously thought. Instead of primarily acting directly on actin filaments within bundles, our results indicate that capping protein and the Arp2/3 complex regulate actin snarls. Actin bundle positioning is affected by the quantity and persistence of these dynamic actin filaments since their overabundance displaces bundles from the membrane, whereas their absence results in excess actin bundles attached to the membrane.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Scanning Electron Microscopy
Adult flies were processed for scanning electron microscopy (SEM) as described previously (Hopmann et al., 1996; Hopmann and Miller, 2003) with the following exceptions: after removal of the wings and legs under CO2 anesthesia, flies were stored in 75% ethanol at 4°C for several days to weeks before processing. In some cases flies were rehydrated in 50% ethanol for 1 h and then in distilled water for 1 h and were fixed using the Osmium-Thiocarbohydrazide-Osmium method (Kelley et al., 1973). The samples were dehydrated in an ethanol series and were critical point-dried from CO2. Mounted samples were sputter-coated with 30 nm of gold and examined in a Hitachi S-450 SEM (Hitachi, Tokyo, Japan) operated at 20 kV accelerating voltage. For quantitative analysis, the four scutellar bristles were scored without knowledge of the genotype using the scale described in the legend for Figure 1. p values were determined using a chi square test.
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Developmental Staging of Pupae
In most cases animals were picked as white prepupae and allowed to develop in a humid chamber at 25°C for the indicated length of time after puparium formation (APF). We have previously noted that cpb mutants are developmentally delayed relative to wild type (Hopmann and Miller, 2003), but in this staging we found that wild-type and mutant bristles reached approximately the same length at the same time APF. Thus much of the developmental delay likely occurs during the five preceding days of embryogenesis and larval development. arp3/arp3 pupal development was delayed by several hours compared with wild type, so comparing these two genotypes at the same time APF was not reasonable. Instead, we examined bristles that were newly emerging (indicated "early" in Figure 3) or fully elongated (indicated "late" in Figure 3). In some cases pupae were maintained at 20°C instead of 25°C; at this temperature animals develop more slowly, but there was no temperature effect on bristle development.
Fluorescent Microscopy
Preparation, staining, and confocal imaging of pupal bristles was performed as described previously (Hopmann and Miller, 2003). Primary antibodies used were affinity purified rat anti-capping protein beta (Hopmann et al., 1996), rabbit anti-Arp 3 (Stevenson et al., 2002), and mouse anti-profilin (chickadee; Verheyen and Cooley, 1994). Secondary antibodies were Alexa-488 anti-rat and Alexa-568 anti-rabbit (both from Molecular Probes, Eugene, OR). Actin was visualized either with GFP-actin or Alexa-568 phalloidin (Molecular Probes).
Transmission Electron Microscopy
The scutellar region of the cuticle was dissected from pharate adults in 0.1 M sodium phosphate, pH 6.8, and fixed for 1 h in 1% glutaraldehyde in the above buffer. Specimens were rinsed extensively and washed overnight in this buffer and then transferred to 0.1 M sodium phosphate, pH 6.0. They were then treated with 1% osmium tetroxide in this buffer for 1 h followed by extensive washing. This was followed with a dehydration series from 10 to 100% ethanol with 10% increases at each 10-min step. Samples were then soaked overnight at 4°C in a 1:1 solution of 100% ethanol and LR White Resin (London Resin Company, Berkshire, England), followed by overnight in 3:1 resin:ethanol and then 3 d at 4°C in 100% resin with one change after the first day. Finally, samples were placed in gelatin capsules filled with 100% resin, left at room temperature for 1 d, and then incubated at 60°C for 48 h to polymerize the resin. All steps were done with mixing, and in some cases a vacuum was applied to the samples for 10 min at various steps in an attempt to increase the penetration of the resin into the bristles. Eighty- to 90-nm sections were cut using a Reichert-Jung Super Nova ultratome (Leica Microsystems, Nussloch, Germany) equipped with a diamond knife and analyzed on a Hitachi H-600 transmission electron microscope. Resin penetration was a constant problem throughout, and only samples that were well embedded were used for analysis. Cuticle areas were measured using ImageJ (NIH, Bethesda, MD).
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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(hereafter referred to as cpb) mutations result in excess F-actin (Hopmann and Miller, 2003) and loss of the Arp2/3 complex, a potent actin nucleator (Welch et al., 1997b; Mullins et al., 1998), is predicted to reduce actin assembly, we hypothesized that mutations in the genes encoding Arp2/3 complex subunits would suppress the phenotypes observed in cpb mutants. cpb mutants have defects in macrochaete morphology (Hopmann et al., 1996). Among other defects, bristles have abnormally organized surface grooves and regions that lack grooves entirely. Additionally, protruding spikes, split ends, and sharp bends are occasionally apparent. Figure 1A, a and b, shows two representative cpb mutant bristles. This phenotype was suppressed by reducing the amount of the Arp 2/3 complex subunit, arp3. Macrochaetes from flies of the genotype cpbF19/cpb6.15; arp3EP3640/+ were often nearly wild type in appearance, with normal groove patterns (Figure 1A, compare c–d with a–b). The extent of phenotypic suppression was quantitated in two ways. In the first assay we scored scutellar bristles for defects by SEM. Our second assay was to count the number of severely defective thoracic macrochaetes per fly visible at the dissecting microscope level. In both assays, scoring was performed without knowledge of genotype. Both assays showed that the suppression of the cpb phenotype by arp3 was strong and statistically significant (Figure 1, B and C), supporting the hypothesis that capping protein and the Arp2/3 complex act in opposition to one another in the developing bristle.
To confirm that the suppression by arp3 was due to its role in the Arp2/3 complex, we examined the effect of mutations in arpc1 and wasp (an activator of the Arp2/3 complex) and deficiencies that remove arpc4 and arpc5. As expected, a mutation in wasp and the deficiencies that remove arpc4 and arpc5 suppressed the cbp phenotype to a similar extent as did the mutation in arp3 (Figure 1 and Supplementary Figure 1). Surprisingly however, mutations in arpc1 enhanced the cpb phenotype (Figure 1 and Supplementary Figure 1). In flies of the genotype cpbF19arpc1Q25st/cpb6.15 +, normal-looking bristles were completely absent, and cuticle morphology defects appeared more severe. In addition to very irregular groove patterns, numerous sharp bends, spikes, splits, and fusions of neighboring bristles were observed (Figure 1A, g and h). We interpret the bristle fusions to indicate that during pupal development, the cellular protrusions that form the bristles lack rigidity and thus sometimes lie across one another. When chitin secretion occurs, they become "glued" together. This enhancement of the cpb bristle phenotype was observed in two different arpc1 alleles (Supplementary Figure 1) isolated in independent screens in different labs (Hudson and Cooley, 2002). The strength of the suppression seen (Supplementary Figure 1) correlates with allele strength (Hudson and Cooley, 2002). Because mutations in arp3 and wasp and deficiencies that remove arpc4 and arpc5 all suppress the cpb phenotype, we conclude that the Arp2/3 complex acts in opposition to capping protein in developing bristles as it does in motile cells. However, our observation that mutations in arpc1 enhance the cpb phenotype suggests that Arpc1 might have an additional role, independent of the rest of the Arp2/3 complex.
Capping Protein and the Arp2/3 Complex Act on Actin Snarls to Indirectly Affect Actin Bundle Positioning
To uncover the mechanisms by which capping protein and the Arp2/3 complex act in opposition to one another, we sought to determine the structure(s) on which these assembly regulators act by examining their localization. We used anti-capping protein serum (Hopmann et al., 1996) and an Arp3-specific antibody (Stevenson et al., 2002) to stain pelts from pupae in which bristles were just starting to emerge. We found that capping protein did not localize to actin bundles in early bristles, but instead localized to snarls, clumps of filamentous actin visible between the actin bundles at the membrane (Figure 2, a–e). In previous work, we found that capping protein was located in small puncta distributed along the length of the actin bundles late in bristle development (Hopmann et al., 1996). Staining of pupal bristles at various times through development revealed that capping protein first localized to the bundles at 40–42 h APF (unpublished data).
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Because both capping protein and Arp3 localized to snarls, we wondered if other actin regulators are on snarls as well. We stained newly emerging bristles with an antibody specific for profilin (Verheyen and Cooley, 1994) and found that it was distributed fairly uniformly throughout the bristles, both on and off of snarls. Like capping protein and Arp3, it was completely excluded from the actin bundles (Figure 2, p–t). The difference between profilin’s localization and that of capping protein and Arp3 was especially evident when we compared computed cross-sections through the bristles; profilin was abundant in the center of the bristles (Figure 2t), whereas capping protein (Figure 2e) and Arp3 (Figure 2j) were restricted to the membrane. This staining pattern appears consistent with profilin’s role as an actin monomer binding protein that functions to promote actin filament assembly.
If the Arp2/3 complex and capping protein do regulate snarls, our expectation is that loss of Arp3 would lead to a loss of snarls, whereas loss of capping protein would result in excess snarls. A further expectation is that reduction of Arp3 should abrogate the excess snarl phenotype of cpb mutants, because of the opposing effects of these regulators on actin assembly. To test these hypotheses, we first examined the effect of loss of Arp3 on developing bristles in comparison to wild-type bristles. Snarls were clearly evident between the bundles in wild-type bristles just after emergence (Figure 3, a and b). Once the bristles were fully elongated, actin snarls were no longer visible, leaving only 10–15 regularly spaced actin bundles against the membrane (Figure 3, j–l). A strikingly different pattern was observed when we examined developing bristles of pupae homozygous for a strong loss of function allele of arp3 (Hudson and Cooley, 2002). In early arp3EP3640 mutant bristles, we were unable to detect snarls and an excess number of actin bundles were juxtaposed to the membrane (Figure 3, d and e). This was especially clear in cross-sectional images of bristles. In wild type, space was visible between the actin bundles (Figure 3c), whereas in arp3 mutants bundles were present nearly continuously along the membrane (Figure 3f). These excess bundles persisted throughout bristle development, such that when the bristles were fully elongated, there was still a substantial excess of actin bundles on the membrane in arp3 mutants compared with wild type (Figure 3, compare m–o to j–l). These excess membrane-associated actin bundles likely are the reason that bristles in homozygous arp3 mutant adults have more, smaller grooves than wild-type bristles (Hudson and Cooley, 2002).
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If capping protein exerts its effect on bundle positioning by controlling snarls, the snarls should be affected in capping protein mutants. Because reduction of capping protein amount increases actin assembly, excess snarls should form when capping protein amount is reduced. To determine if this is the case, we compared developmental time courses in wild-type and cpb mutants. In wild-type pupae, actin snarls were clearly evident at 32–33 h (Figure 4Aa), began diminishing around 35–36 h (unpublished data), and were nearly undetectable at 38–39 h (Figure 4Ac). By contrast, in cpb mutants snarls persisted much longer; they were still clearly evident at 38–39 h (Figure 4Ag). Furthermore, examination of confocal images of planes through the middle of the bristles showed that although the center of wild-type bristles was devoid of actin bundles, this region was full of bundles in cpb mutants (Figure 4A; compare h with d). This suggests that excess persisting snarls interfere with the ability of bundles to localize adjacent to the membrane.
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). Reducing the amount of profilin balances the reduction of capping protein so that actin assembly is more normal in this mutant combination. If the effect of profilin is also via snarls, the amount and persistence of snarls should resemble wild type in capping protein/profilin mutant combinations. Indeed we found that snarls in cpbF19 chic221/cpb6.15 + bristles looked similar to wild type: they were obvious at 32–33 h (Figure 4A, i), reduced at 35–36 h (unpublished data), and were not visible at 38–39 h (Figure 4Ak). Additionally, in contrast to cpb mutants, the actin bundles in cpbF19 chic221/cpb6.15 + bristles were properly localized against the membrane (Figure 4A, compare l with h). Overexpression of profilin in the bristle resulted in excess snarls and actin bundles displaced into the center of the bristle (Figure 4A, m–p). Thus, like capping protein and the Arp2/3 complex, profilin regulates actin assembly in snarls. In addition, bundle positioning correlates with the amount and persistence of snarls in this genotype as well. Because capping protein and the Arp2/3 complex both localized to snarls and had opposing effects on snarl abundance, we predicted that loss of one copy of arp3 should suppress the excess snarl phenotype seen in cpb mutants. Although no suppression was observed at early time points—bristles on pupae of the genotype cpbF19/cpb6.15; arp3EP3640/+ looked nearly identical to cpb bristles from 32 to 39 h APF (Figure 4A, compare q–t to e–h)—we did see suppression beginning at 41–42 h APF. At this time point, we found bristles of two different types in both genotypes: those lacking snarls and having membrane-localized bundles and those with snarls and interior bundles. However, bristles with snarls and interior bundles were far more prevalent in cpbF19/cpb6.15; +/+ pupae than in cpbF19/cpb6.15; arp3EP3640/+ pupae (Figure 4B). At 46 h APF, no snarls were seen in either genotype, but a range of bundle position phenotypes were observed. We categorized them as all membrane associated, mostly membrane associated, mostly internal, or all internal. Representative examples of each are shown in Figure 5A. Although both cpbF19/cpb6.15; +/+ and cpbF19/cpb6.15; arp3EP3640/+ pupae had bristles in all of the categories, more of the cpbF19/cpb6.15; arp3EP3640/+ bristles had all or mostly membrane attached bundles (p < 0.01; Figure 5B). These results indicate that capping protein and the Arp2/3 complex act in opposition to one another and likely act primarily through effects on snarls.
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Proper Bundle Positioning Is Important For Organized Cuticle Secretion
We wondered why having the correct number of properly spaced bundles is important for bristle development. We postulated that this might be important for proper cuticle secretion at the end of bristle elongation. To address this idea, we analyzed cross-sections through the tips of wild-type and homozygous arp3 pharate adult macrochaetes by transmission electron microscopy. To determine if arp3 mutants were deficient in their ability to secrete the proper amount of cuticle material, we measured the cuticle area and compared this to the area that had previously been occupied by the bristle cell (the hole in the center of each bristle). We found no significant difference between wild-type and arp3 mutants (for wild type, the average ratio of cuticle area/cell area was 3.2 ± 0.7; for arp3 the ratio was 2.5 ± 1.0, p = 0.1). Although there was no difference in total amount of cuticle secretion, we did observe striking defects in the organization of the cuticle. Although wild-type bristles were quite uniform and had an average of 8.1 large ridges (range 7–10, n = 8) at the tips (Figure 6, a and b), arp3 bristles had an average of 11.2 ridges (range 7–16, n = 10) at the tips (Figure 6, c and d) and this difference was significant (p < 0.005). Additionally, 5 of the 10 arp3 bristles we analyzed had an irregular shape or a dramatic protrusion of the cuticle (Figure 6d). This was never seen in wild type. We conclude that loss of functional Arp2/3 complex leads to a reduction of actin snarls, resulting in an overabundance of actin bundles associated with the membrane that interfere with orderly cuticle secretion.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Nonbundle Actin Snarls Contribute to the Proper Size and Distribution of Cortical Actin Bundles
Here we present evidence that early in bristle development capping protein and the Arp2/3 complex control actin assembly not in actin bundles, but rather in the actin snarls that are present between these bundles. We hypothesize that snarls influence the proper positioning of the cortical actin bundles by competing with them for binding sites on the membrane. When the level of snarls is severely reduced (in homozygous arp3 mutant pupae), excess actin bundles associate with the membrane. This results in adult bristles that are shorter than wild type (unpublished data) and have more, smaller grooves in their cuticle (Hudson and Cooley, 2002). In the presence of too many snarls (in transheterozygous cpb mutant pupae), actin bundles are unable to form against the membrane and instead develop internally. Because they are not stabilized by attachment to the membrane, they become irregularly sized and highly variable in number. This results in a cell membrane that is inadequately supported by actin bundles. Adult bristles that have numerous defects, such as irregular groove patterns and smooth regions in the cuticle, as well as bends, splits, and protrusions in the shaft (Hopmann et al., 1996) are produced.
Tilney et al. (2003) have previously demonstrated that actin snarls are dynamic and have suggested that their 2-min half-life is due to their failure to be stabilized either by cross-linking to other filaments or by forming stable attachments to the membrane. These authors argued that those filaments that do not become stabilized are turned over, and thus the actin bundles develop in a Darwinian "survival of the fittest" manner. The snarls were proposed to be nonproductive actin structures. We propose here that the snarls have a function in forming properly shaped bristles: they are important for proper positioning of bundles. Snarls may have additional functions as well. They cause protrusions in the membrane between the bundles (Tilney et al., 1996, 2003) and lie beneath the regions that will become the ridges in the cuticle of the adult bristle (the actin bundles lie below the grooves; Overton, 1967). Thus the Arp2/3 complex–directed actin assembly in snarls may push the membrane out, much in the way that this complex causes membrane protrusion at the leading edge of motile cells (Welch et al., 1997a; Mullins et al., 1998; Svitkina and Borisy, 1999). This region of the bristle membrane is then kept free from actin bundles so that the proper number and size of bundles are formed. We have shown that when too many bundles form, cuticle is secreted in a disorganized manner. Thus it may be important that some regions of the membrane are kept clear of actin bundles to allow for proper secretion of chitin. Alternatively, actin filaments in the snarls could be directly involved in membrane deposition that occurs during cell extension and/or in the secretion of cuticle (patches of cuticulin are evident on developing bristles as early as 36 h APF; Overton, 1967). Snarls may be analogous to actin patches in Saccharomyces cerevisiae that are turned over very rapidly (Smith et al., 2001), contain the Arp2/3 complex (Winter et al., 1997) and capping protein (Amatruda and Cooper, 1992), and are involved in endocytosis and cell wall synthesis (Utsugi et al., 2002; Kaksonen et al., 2003).
The interplay we see between actin bundles and snarls in bristles is reminiscent of that seen between actin cables and patches in S. cerevisiae. Yeast cells lacking functional capping protein have diminished actin cables and excess patches (Amatruda et al., 1990, 1992). Conversely, long-term lack of Arp3 results in cells that have lost patches and accumulate actin bundles (Winter et al., 1997). The antagonistic relationship between capping protein and the Arp2/3 complex, as well as their opposing effects on actin patches and bundles, might thus be a common theme in the regulation of the actin cytoskeleton.
Why Do Mutations in arpc1 Enhance the cpb Bristle Phenotype?
We have shown that mutations in arp3 and wasp, as well as deficiencies that remove arpc4 and arpc5, all suppress the cpb bristle phenotype as would be expected from the opposing biochemical functions of the Arp2/3 complex (filament assembly nucleator) and capping protein (prevents further assembly of actin filaments). So it is surprising to find that mutations in arpc1 have the opposite phenotype. This is an especially unexpected result given that previous work has demonstrated that adult bristles completely lacking Arpc1 or Arp3 have an identical mild phenotype of more, smaller grooves (Hudson and Cooley, 2002). It is tempting to suggest that there is a genetic background issue at play here. We believe this is unlikely, however, because two different arpc1 alleles isolated in independent labs in different genetic backgrounds gave the same result.
One explanation for the enhancement seen by mutations in arpc1 is that in the absence of this subunit, a complex forms that in some way acts as a neomorph. Arpc1 contains the WASp interaction capability of the complex (Pan et al., 2004), so complexes lacking this subunit might be unactivatable, and thus might block the interaction between actin filaments and other important binding proteins. Although we cannot rule this possibility out, we think it is unlikely because of the following: Gournier et al. (2001) have shown that the Arpc1 (therein referred to as p41) and Arpc5 (p16) subunits form an interacting pair. They find that when complexes are formed in the absence of Arpc5, Arpc1 also fails to assemble into the complex. If this neomorph model were correct (and assuming fly Arp2/3 complex components behave in the same way as their human homologues), we would expect that reduction of Arpc5 would lead to loss of Arpc1 from the complex and cause enhancement of the cpb phenotype. However, this is not what we observed. A deficiency chromosome lacking the arpc5 region in fact suppressed the cpb bristle phenotype.
To explain arpc1 enhancement of bundle displacement observed in cpb mutants, we note that the difference in phenotype between cpb6.15/cpbF19 and cpb6.15 +/cpbF19 arpc1Q25st is only apparent at late developmental times. In both genotypes early in development, bundles are displaced from the membrane. However, in cpb6.15/cpbF19 at late times, bundles often are associated with the membrane, whereas in cpb6.15 +/cpbF19 arpc1Q25st they rarely are. Interestingly, in cpb null epithelial clones, bundles also remain displaced (Hopmann and Miller, 2003 and unpublished data). These similar phenotypes suggest that capping protein and Arpc1 both participate in a late membrane-attachment process. Thus, we suggest that in cases where bundles form not associated with the membrane initially, capping protein and Arpc1 promote bundle attachment late in development. The ability of some bundles to associate with the membrane in cpb6.15/cpbF19 bristles is likely due to the fact that in this genotype capping protein is not completely eliminated but rather is reduced to 48% of the wild-type level (Hopmann et al., 1996). When capping protein is completely absent or when both capping protein and Arpc1 are reduced in amount, then this late membrane attachment does not occur. Interestingly, capping protein localization on bundles starts at 
42 h APF (Hopmann et al., 1996 and unpublished data), around the time when the effect of capping protein and Arpc1 on bundle membrane association is manifest. Under normal circumstances, this late bundle attachment activity is not essential, because bundles are already associated with the membrane. Similarly, in cases where snarls are reduced and excess bundles form (arp3, and presumably, arpc1 homozygous) this late attachment activity is not needed because the bundles form initially against the membrane. Obviously, other as yet unidentified proteins must be important for membrane association at early times.
Capping protein has previously been observed to be required for attachment of actin filaments to the Z disks during myofibrillogenesis (Schafer et al., 1995); a role in membrane attachment in bristles would be consistent with this function. Welch et al. (1997a) have suggested that Arpc1 (therein referred to as p41-Arc) might serve as a link to the membrane through its WD repeats binding to pleckstrin homology domains, which could in turn bind to phosphatidyl inositol 4,5-bisphosphate. Additionally, Arpc1 binds to the activating domain of WASp (Pan et al., 2004), which is itself activated at the membrane.
We suggest here that Arpc1 has a function outside the Arp 2/3 complex. Although work in yeast (Winter et al., 1999b) demonstrating an unique function for Arpc1 was later shown to be an experimental artifact (Pan et al., 2004), recent studies in the moss Physcomitrella patens has revealed that removal of Arpc1 results in different phenotypes than removal of Arpc4 (Harries et al., 2005; Perroud and Quatrano, 2006). Thus, it will be important to explore this phenomenon of potential differential Arp2/3 complex subunit functions further in the future.
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ACKNOWLEDGMENTS |
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Footnotes |
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The online version of this contains supplemental material at MBC Online (http://www.molbiolcell.org). 
This article was published online ahead of print in MBC in Press (http://www.molbiolcell.org/cgi/doi/10.1091/mbc.E06-06-0500) on July 5, 2006.
Address correspondence to: Kathryn G. Miller (miller{at}biology.wustl.edu)
Abbreviations used: APF, after puparium formation; Arp2/3, actin-related protein 2/3; cpb, capping protein beta; SEM, scanning electron microscopy; WASp, Wiscott Aldrich Syndrome protein.
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REFERENCES |
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Amatruda, J. F. and Cooper, J. A. (1992). Purification, characterization, and immunofluorescence localization of Saccharomyces cerevisiae capping protein. J. Cell Biol 117, 1067–1076.
Amatruda, J. F., Gattermeir, D. J., Karpova, T. S., Cooper, J. A. (1992). Effects of null mutations and overexpression of capping protein on morphogenesis, actin distribution and polarized secretion in yeast. J. Cell Biol 119, 1151–1162.
Appel, L. F., Prout, M., Abu-Shumays, R., Hammonds, A., Garbe, J. C., Fristrom, D., Fristrom, J. (1993). The Drosophila Stubble-stubbloid gene encodes an apparent transmembrane serine protease required for epithelial morphogenesis. Proc. Natl. Acad. Sci. USA 90, 4937–4941.
Egile, C., Loisel, T. P., Laurent, V., Li, R., Pantaloni, D., Sansonetti, P. J., Carlier, M. F. (1999). Activation of the CDC42 effector N-WASP by the Shigella flexneri IcsA protein promotes actin nucleation by Arp2/3 complex and bacterial actin-based motility. J. Cell Biol 146, 1319–1332.
Gournier, H., Goley, E. D., Niederstrasser, H., Trinh, T., Welch, M. D. (2001). Reconstitution of human Arp2/3 complex reveals critical roles of individual subunits in complex structure and activity. Mol. Cell 8, 1041–1052.[CrossRef][Medline]
Harries, P. A., Pan, A., Quatrano, R. S. (2005). Actin-related protein2/3 complex component ARPC1 is required for proper cell morphogenesis and polarized cell growth in Physcomitrella patens. Plant Cell 17, 2327–2339.
Hopmann, R., Cooper, J. A., Miller, K. G. (1996). Actin organization, bristle morphology, and viability are affected by actin capping protein mutations in Drosophila. J. Cell Biol 133, 1293–1305.
Hopmann, R. and Miller, K. G. (2003). A balance of capping protein and profilin functions is required to regulate actin polymerization in Drosophila bristle. Mol. Biol. Cell 14, 118–128.
Hudson, A. M. and Cooley, L. (2002). A subset of dynamic actin rearrangements in Drosophila requires the Arp2/3 complex. J. Cell Biol 156, 677–687.
Isenberg, G., Aebi, U., Pollard, T. D. (1980). An actin-binding protein from Acanthamoeba regulates actin filament polymerization and interactions. Nature 288, 455–459.[CrossRef][Medline]
Kaksonen, M., Sun, Y., Drubin, D. G. (2003). A pathway for association of receptors, adaptors, and actin during endocytic internalization. Cell 115, 475–487.[CrossRef][Medline]
Kelley, R. O., Dekker, R. A., Bluemink, J. G. (1973). Ligand-mediated osmium binding: its application in coating biological specimens for scanning electron microscopy. J. Ultrastruct. Res 45, 254–258.[CrossRef][Medline]
Lees, A. D. and Picken, L.E.R. (1944). Shape in relation to fine structure in the bristles of Drosophila melanogaster. Proc. Roy. Soc. Lond. Ser. B. Biol. Sci. 132, 396–423.
Lees, A. D. and Waddington, C. H. (1944). The development of the bristles in normal and some mutant types of Drosophila melanogaster. Proc. Roy. Soc. Lond. Ser. B. Biol. Sci. 132, 396–423.
Loisel, T. P., Boujemaa, R., Pantaloni, D., Carlier, M. F. (1999). Reconstitution of actin-based motility of Listeria and Shigella using pure proteins. Nature 401, 613–616.[CrossRef][Medline]
Machesky, L. M., Atkinson, S. J., Ampe, C., Vandekerckhove, J., Pollard, T. D. (1994). Purification of a cortical complex containing two unconventional actins from Acanthamoeba by affinity chromatography on profilin-agarose. J. Cell Biol 127, 107–115.
Machesky, L. M., Mullins, R. D., Higgs, H. N., Kaiser, D. A., Blanchoin, L., May, R. C., Hall, M. E., Pollard, T. D. (1999). Scar, a WASp-related protein, activates nucleation of actin filaments by the Arp2/3 complex. Proc. Natl. Acad. Sci. USA 96, 3739–3744.
Mullins, R. D., Heuser, J. A., Pollard, T. D. (1998). The interaction of Arp2/3 complex with actin: nucleation, high affinity pointed end capping, and formation of branching networks of filaments. Proc. Natl. Acad. Sci. USA 95, 6181–6186.
Overton, J. (1967). The fine structure of developing bristles in wild type and mutant Drosophila melanogaster. J. Morphol 122, 367–379.[CrossRef][Medline]
Pan, F., Egile, C., Lipkin, T., Li, R. (2004). ARPC1/Arc40 mediates the interaction of the actin-related protein 2 and 3 complex with Wiskott-Aldrich syndrome protein family activators. J. Biol. Chem 279, 54629–54636.
Perroud, P. F. and Quatrano, R. S. (2006). The role of ARPC4 in tip growth and alignment of the polar axis in filaments of Physcomitrella patens. Cell Motil. Cytoskeleton 63, 162–171.[CrossRef][Medline]
Pollard, T. D. and Borisy, G. G. (2003). Cellular motility driven by assembly and disassembly of actin filaments. Cell 112, 453–465.[CrossRef][Medline]
Rohatgi, R., Ma, L., Miki, H., Lopez, M., Kirchhausen, T., Takenawa, T., Kirschner, M. W. (1999). The interaction between N-WASP and the Arp2/3 complex links Cdc42-dependent signals to actin assembly. Cell 97, 221–231.[CrossRef][Medline]
Schafer, D. A., Hug, C., Cooper, J. A. (1995). Inhibition of CapZ during myofibrillogenesis alters assembly of actin filaments. J. Cell Biol 128, 61–70.
Smith, M. G., Swamy, S. R., Pon, L. A. (2001). The life cycle of actin patches in mating yeast. J. Cell Sci 114, 1505–1513.[Abstract]
Stevenson, V., Hudson, A., Cooley, L., Theurkauf, W. E. (2002). Arp2/3-dependent pseudocleavage [correction of psuedocleavage] furrow assembly in syncytial Drosophila embryos. Curr. Biol 12, 705–711.[CrossRef][Medline]
Svitkina, T. M. and Borisy, G. G. (1999). Arp2/3 complex and actin depolymerizing factor/cofilin in dendritic organization and treadmilling of actin filament array in lamellipodia. J. Cell Biol 145, 1009–1026.
Tilney, L. G., Connelly, P., Smith, S., Guild, G. M. (1996). F-actin bundles in Drosophila bristles are assembled from modules composed of short filaments. J. Cell Biol 135, 1291–1308.
Tilney, L. G., Connelly, P. S., Ruggiero, L., Vranich, K. A., Guild, G. M. (2003). Actin filament turnover regulated by cross-linking accounts for the size, shape, location, and number of actin bundles in Drosophila bristles. Mol. Biol. Cell 14, 3953–3966.
Tilney, L. G., Connelly, P. S., Vranich, K. A., Shaw, M. K., Guild, G. M. (2000). Actin filaments and microtubules play different roles during bristle elongation in Drosophila. J. Cell Sci 113, 1255–1265.[Abstract]
Tilney, L. G., Tilney, M. S., Guild, G. M. (1995). F actin bundles in Drosophila bristles. I. Two filament cross-links are involved in bundling. J. Cell Biol 130, 629–638.
Utsugi, T., Minemura, M., Hirata, A., Abe, M., Watanabe, D., Ohya, Y. (2002). Movement of yeast 1,3-beta-glucan synthase is essential for uniform cell wall synthesis. Genes Cells 7, 1–9.[Abstract]
Verheyen, E. M. and Cooley, L. (1994). Profilin mutations disrupt multiple actin-dependent processes during Drosophila development. Development 120, 717–728.[Abstract]
Wahlstrom, G., Vartiainen, M., Yamamoto, L., Mattila, P. K., Lappalainen, P., Heino, T. I. (2001). Twinfilin is required for actin-dependent developmental processes in Drosophila. J. Cell Biol 155, 787–796.
Welch, M. D., DePace, A. H., Verma, S., Iwamatsu, A., Mitchison, T. J. (1997a). The human Arp2/3 complex is composed of evolutionarily conserved subunits and is localized to cellular regions of dynamic actin filament assembly. J. Cell Biol 138, 375–384.
Welch, M. D., Iwamatsu, A., Mitchison, T. J. (1997b). Actin polymerization is induced by Arp2/3 protein complex at the surface of Listeria monocytogenes. Nature 385, 265–269.[CrossRef][Medline]
Winter, D., Lechler, T., Li, R. (1999a). Activation of the yeast Arp2/3 complex by Bee1p, a WASP-family protein. Curr. Biol 9, 501–504.[CrossRef][Medline]
Winter, D., Podtelejnikov, A. V., Mann, M., Li, R. (1997). The complex containing actin-related proteins Arp2 and Arp3 is required for the motility and integrity of yeast actin patches. Curr. Biol 7, 519–529.[CrossRef][Medline]
Winter, D. C., Choe, E. Y., Li, R. (1999b). Genetic dissection of the budding yeast Arp2/3 complex: a comparison of the in vivo and structural roles of individual subunits. Proc. Natl. Acad. Sci. USA 96, 7288–7293.
Yarar, D., To, W., Abo, A., Welch, M. D. (1999). The Wiskott-Aldrich syndrome protein directs actin-based motility by stimulating actin nucleation with the Arp2/3 complex. Curr. Biol 9, 555–558.[CrossRef][Medline]
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