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Address correspondence to Eyal D. Schejter, Dept. of Molecular Genetics, Weizmann Institute of Science, Rehovot 76100, Israel. Tel.: 972-8-934-2207. Fax: 972-8-934-4108. E-mail: eyal.schejter{at}weizmann.ac.il
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Abstract |
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Key Words: Scar; Arp2/3; Wasp; actin; Drosophila
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Introduction |
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Biochemical studies have provided detailed information about the molecules that influence actin dynamics (Pantaloni et al., 2001). Of particular significance is the Arp2/3 complex that stimulates microfilament nucleation, the rate-limiting step in actin polymerization (Mullins et al., 1998; Welch et al., 1998). The Arp2/3 complex consists of seven protein subunits, including the actin-related Arp2 and Arp3, and is conserved among eukaryotes (Machesky and Gould, 1999; May, 2001). Members of the evolutionarily conserved Wiskott-Aldrich Syndrome protein (WASp)* and Scar/WAVE family function as strong potentiators of Arp2/3 complex activity (Higgs and Pollard, 2001). Distinct WASp and Scar/WAVE branches of this family have been recognized in diverse organisms, including Dictyostelium, Caenorhabditis elegans, Drosophila, and mammals. WASp and Scar/WAVE proteins share a common domain structure that mediates activation of the Arp2/3 complex in response to multiple signaling pathways. All members of the WASp-Scar/WAVE family possess a common COOH-terminal (WA) domain that stimulates actin polymerization through association with monomeric actin and the Arp2/3 complex, whereas their NH2-terminal domains are structurally distinct and serve as signal-responsive regulatory regions (Fig. 1). The molecular mechanisms controlling WASp function are well characterized (Fawcett and Pawson, 2000), whereas regulatory aspects of Scar function are now beginning to emerge (Takenawa and Miki, 2001).
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Results |
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To examine SCAR protein expression and subcellular localization, we generated a polyclonal antibody to the unique SCAR NH2-terminal domain (see Materials and methods). We found that SCAR protein is present in early blastoderm embryos and in the embryonic CNS (Fig. 2) consistent with its mRNA expression (unpublished data). In the blastoderm, SCAR protein colocalizes with filamentous actin structures that are dynamically regulated during the cell cycle (Fig. 2, A and C; see below). In the CNS, SCAR protein is specifically localized to axons (Fig. 2, E and F). This pattern of SCAR protein expression in the embryo provided us with an initial indication of the potential sites of SCAR gene activity.
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To obtain deletions in the SCAR locus, we generated imprecise excision alleles of the SCARk13811 insertion, all of which were homozygous lethal and failed to complement the lethality of SCARk13811. The homozygous lethality of the 37 excision allele was rescued by ubiquitous expression of the full-length SCAR cDNA, and we refer to this allele as SCAR
37. SCAR
37 was molecularly characterized and removes all SCAR sequences downstream of the insertion site (Fig. 1 A). This excision event also removes portions of the neighboring piwi transcription unit. Since piwi function is restricted to maintenance and proliferation of germline stem cells (Cox et al., 1998, 2000), the SCAR
37 phenotypes described below, in distinct tissues, are likely to represent consequences of disrupting SCAR function alone. Developmental defects were considerably weaker in SCARk13811, indicating that the insertion allele retains partial SCAR activity.
In addition to SCAR and Wasp (Ben-Yaacov et al., 2001), the sequenced Drosophila genome contains predicted homologs of the seven members of the Arp2/3 complex (Fyrberg et al., 1994; Goldstein and Gunawardena, 2000). Mutations have been recovered in two Arp2/3 complex components, Arp3 (Rørth, 1996; Berkeley Drosophila Genome Project) and Arpc1 (Hudson and Cooley, 2002). This set of mutations provides an opportunity to analyze the role of Arp2/3-based signaling in different contexts within a multicellular organism and to ascertain the physiological contributions of the SCAR and Wasp activators.
SCAR and Arpc1 are required for cytoplasmic organization in the blastoderm embryo
Homozygous mutations in either SCAR allele result in zygotic lethality, which can occur during late embryogenesis, larval, and early pupal stages. However, maternally provided SCAR gene products may compensate for loss of zygotic gene function and mask an essential requirement during embryogenesis. To interfere with the maternal gene contribution, we used FLP-mediated recombination to produce homozygous mutant clones within the germline of heterozygous females (Chou and Perrimon, 1996). Strong disruption of maternal SCAR or Arpc1 in this manner results in developmental arrest during oogenesis (see below). However, germline clones homozygous for the weaker SCARk13811 or Arpc1R337st alleles give rise to fertilizable eggs, enabling study of functional requirements for SCAR and the Arp2/3 complex during embryogenesis. These embryos are designated SCARmat and Arpc1mat, respectively. To compare the roles of SCAR and Wasp, we examined embryos derived from germline clones for the strong loss of function Wsp3 allele (Wspmat embryos). The Wsp3 frameshift mutation truncates the protein before the highly conserved WA domain that is required for Arp2/3 activation and is a probable null allele (Ben-Yaacov et al., 2001).
The early blastoderm embryo undergoes 13 nuclear divisions without accompanying cytokinesis, producing a multinucleate syncytium. The majority of nuclei migrate to the surface by cycle 10, where they undergo four synchronous rounds of division before their compartmentalization into individual cells during interphase of cycle 14 (Zalokar and Erk, 1976). Surface nuclei maintain a uniform distribution throughout these final syncytial divisions (Fig. 3, A and B). Examination of the spatial distribution of nuclei revealed a requirement for SCAR and Arpc1, but not Wasp, during these cortical division cycles (Fig. 3, CH). SCARmat and Arpc1mat mutants exhibited defects in the uniform spacing of interphase nuclei beginning in cycle 11, whereas Wspmat mutants displayed wild-type nuclear organization. By cycles 12 and 13, increased defects in nuclear spacing in SCAR and Arpc1 were accompanied by the appearance of abnormal nuclear morphologies, including large or elongate DNA masses that are likely to represent the fusion of adjacent nuclei.
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SCAR and Arpc1 are required for actin polymerization and regulation of dynamic actin structures in the blastoderm embryo
The syncytial blastoderm contains well-defined filamentous actin structures that exhibit dynamic cell cycle regulation (Karr and Alberts, 1986). Actin is organized into caps overlying individual nuclei during interphase of cortical cycles 1014 (Fig. 4, A and C). During mitosis, actin is redistributed into a network of metaphase furrows that separate adjacent spindles (Fig. 5, A and C). Genetic and drug interference studies demonstrate that organization of the actin cytoskeleton is crucial for the uniform arrangement of blastoderm nuclei (Foe et al., 1993; Schejter and Wieschaus, 1993). The nuclear defects in SCAR mutants, and SCAR protein colocalization with filamentous actin (Fig. 2), raise the possibility that SCAR may function in the regulation of actin structures in the blastoderm embryo.
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Major defects in cortical actin structures were also observed in Arpc1mat embryos, where interphase actin caps were abnormal and actin appeared to be depleted from the regions above individual nuclei (Fig. 4, I and K, 37/37 embryos). This depletion is apparent most readily in cross-section (Fig. 5 Q). As in SCAR, metaphase furrows failed to form in Arpc1mat embryos (Fig. 5 M, 12/12 embryos). However, unlike SCAR, metaphase actin exhibited a diffuse localization to the broad region between spindles (Fig. 5 R). These results demonstrate that the Arp2/3 complex component Arpc1 is required for the formation of both interphase actin caps and metaphase actin furrows. The greater severity of the Arpc1 phenotype compared with SCAR could reflect a difference in residual gene activity of these partial loss of function alleles.
SCAR and Arpc1, therefore, provide functions that are critical for proper formation of cortical actin structures. In contrast, actin caps and furrows formed normally in Wspmat embryos (27 interphase and 5 metaphase embryos) (PFig. 5, O and P). It is worth emphasizing in this context that both the SCAR and Arpc1 phenotypes result from a partial loss of gene function, whereas early embryogenesis can proceed normally despite complete lack of Wsp gene activity.
In addition to defects in organization of microfilament structures, overall actin levels in Arpc1mat and SCARmat embryos appeared consistently lower than in wild type. To rigorously assess differences in actin levels, we quantitated surface filamentous actin in syncytial embryos using a phalloidin fluorescence assay. We found that both Arpc1mat and SCARmat mutants exhibited significantly reduced levels of surface actin compared with control embryos (Fig. 6). The more severe loss of actin in Arpc1mat correlates with the greater disruption of actin structures in this mutant. These results indicate that Arpc1 and SCAR are both required for the generation of bulk filamentous actin in the blastoderm and suggest a common basis for the defects in cortical actin structures of Arpc1 and SCAR mutant embryos.
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Since SCAR is essential for normal CNS axon morphology, we also examined the zygotic effect of mutations in two members of the Arp2/3 complex, Arp3 and Arpc1. Arp3 zygotic mutant embryos exhibited a partially penetrant defect in CNS axon morphology with a range of phenotypes that strongly resemble SCARmat/zyg mutants (Fig. 7, E and J). In particular, the majority of Arp3 mutants displayed breaks in the longitudinal and commissural axon bundles (Fig. 7 E) (48% of segments in mutant embryos, n = 115 segments). A subset of Arp3 mutants exhibited defects such as commissure defasciculation or fusion (28% of segments) and medially or laterally displaced axons (3% of segments). Arp3 heterozygotes also exhibited a low penetrance of axon defects (Fig. 7 J). The CNS morphology of zygotic Arpc1 single mutants appeared normal, perhaps due to the continued presence of maternal gene products. However, combining zygotic Arpc1 mutations with an Arp3 heterozygous background produced defects that were significantly more severe than in Arp3 heterozygotes alone (Fig. 7, F and J). These phenotypes demonstrate a similar functional requirement for SCAR and Arp2/3 complex components during CNS development.
The contribution of Wasp function to CNS axon morphology is more difficult to assess, since complete removal of maternal and zygotic Wsp using the strong Wsp3 allele (Wspmat/zyg embryos) produces cell fate defects in CNS lineages (Ben-Yaacov et al., 2001). An apparent thickening of commissural bundles suggestive of an increase in neuronal number was observed in a majority of Wspmat/zyg embryos (Fig. 7 D). In addition, most Wspmat/zyg embryos contained one to two segments with axon bundles collapsed at the midline (Fig. 7 D) (73% of embryos, n = 41). Despite these phenotypes, Wspmat/zyg embryos did not exhibit the severe defects in axon morphology present in SCAR and Arp3 mutants. Although removal of zygotic SCAR or Wsp function alone did not disrupt CNS morphology (Fig. 7, B and I), zygotic reduction of SCAR and Wsp together produced significant defects (Fig. 7, G, H, and I) that resemble the strong SCARmat/zyg phenotype. Therefore, although loss of Wasp function alone does not cause the significant axon defects produced by loss of SCAR, Wasp can influence axon morphology in situations where SCAR function is compromised.
SCAR, and not Wasp, is required with the Arp2/3 complex for egg chamber morphology during oogenesis
Although the partial reduction of SCAR function associated with the SCARk13811 insertion allele is sufficient for normal egg production, the more severe SCAR37 excision allele produces small and abnormally shaped eggs indicative of a defect in oogenesis. Drosophila ovaries house a series of egg chambers that each contain 16 cells (the oocyte and a 15-cell nurse cell complex) interconnected by cytoplasmic bridges (ring canals) that arise from incomplete cytokinesis during mitosis (Spradling, 1993). Morphological defects are apparent in SCAR
37 germline clones during the final phases of oogenesis (Fig. 8). In particular, nurse cells become multinucleate, as many of the actin-lined nurse cell membranes are absent (Fig. 8 B). The morphological abnormalities extend to additional structures, including the actin-rich ring canals, which are significantly smaller than in wild-type and aberrantly shaped (Fig. 8, F and H).
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SCAR and Wasp are required for distinct aspects of Arpc1 function in the adult eye
The above phenotypic analyses identify several Arp2/3-dependent morphological processes that rely on SCAR but are largely independent of Wasp. We therefore asked whether the reciprocal situation exists, namely, are there Arp2/3-mediated events that rely on Wasp but are independent of SCAR? Wasp provides an essential contribution to cell fate decisions in several neural lineages in the Drosophila embryo and adult (Ben-Yaacov et al., 2001). Furthermore, the Arp2/3 complex component Arpc1 is required for Wasp-dependent cell fate changes during sensory organ development, and association with Arp2/3 is essential for Wasp function in this context (Tal et al., 2002). This requirement provides an opportunity to examine whether developmental events dependent on Wasp also require SCAR function.
In the adult peripheral nervous system, a primary consequence of mutations in Wsp is the excessive differentiation of sensory organ neurons at the expense of nonneuronal cell types, resulting in a marked absence of mechanosensory bristles (Fig. 9, A and B). We used the ey-FLP-FRT system (Newsome et al., 2000) to generate mosaic SCAR and Arpc1 heterozygous flies in which head capsule structures and cuticle are derived from homozygous mutant clones induced in the eye imaginal disc. Arpc1 mosaic heads like, Wsp, display a pronounced bristle loss phenotype (Fig. 9 C), which results from cell fate defects similar to those present in Wsp mutants (Tal et al., 2002). In contrast, the sensory organ pattern in mosaic heads of strong SCAR alleles appears wild type (Fig. 9 D), suggesting that SCAR does not play an essential role in lineage decisions mediated by Wasp and the Arp2/3 complex.
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Discussion |
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SCAR is a key regulator of CNS axon morphology
Here we demonstrate a requirement for SCAR in the regulation of axon morphology in the Drosophila CNS. The striking enrichment of SCAR protein in axons is consistent with a direct role for SCAR in axon development. In particular, the breaks in longitudinal and commissural axon bundles in SCAR mutant embryos may indicate a defect in axon growth. However, these phenotypes could also reflect defects in other aspects of nervous system formation, such as axon guidance, axon initiation, or neuronal differentiation. Morphological characterization of SCAR mutants at single neuron resolution will provide greater insight into the processes that require SCAR function.
The CNS axon defects in SCAR mutant embryos resemble defects caused by simultaneous zygotic disruption of the Abl tyrosine kinase and a diverse set of elements including the Fasciclin I transmembrane protein, Armadillo/ß-catenin, Chickadee/profilin, and the Trio Rac/Rho guanine nucleotide exchange factor (Lanier and Gertler, 2000). Interestingly, Scar/WAVE-1 has been shown to associate with the SH3 domain of the Abl tyrosine kinase, suggesting that they may directly interact in vivo (Westphal et al., 2000). The observation that multiple zygotic mutations are required to replicate the SCAR phenotype is consistent with a model where SCAR functions downstream of multiple signaling pathways that converge on regulation of the actin cytoskeleton.
The defects in axon morphology caused by reduction of maternal and zygotic SCAR are similar to those produced by zygotic disruption of Arp3 or simultaneous zygotic disruption of Arp3 and Arpc1 or SCAR and Wasp. These results suggest that SCAR, Wasp, and the Arp2/3 complex may affect a common process in neuronal development involving actin regulation. The contribution of both SCAR and Wasp to axon morphology could be explained by several possible mechanisms. In one model, SCAR and Wasp might regulate a common activity of the Arp2/3 complex, such as in the context of a specific actin structure or in contribution to bulk actin levels. Their functional differences in vivo could be achieved through differences in expression, activation, or subcellular localization. Alternatively, SCAR and Wasp could regulate distinct activities of the Arp2/3 complex, producing different actin structures that participate in diverse cell biological processes such as cell morphology (SCAR) and asymmetric cell division (Wasp). It will be interesting to examine how SCAR and Wasp intersect with regulators and effectors to achieve the specific organization of actin structures in different contexts.
SCAR and the Arp2/3 complex regulate actin polymerization and organization in the blastoderm embryo
The dramatic reduction in actin levels of Drosophila Arpc1 mutants indicates that the Arp2/3 complex is an essential source of filamentous actin in the blastoderm embryo. This is consistent with experiments in other systems, where the Arp2/3 complex is required for actin polymerization in yeast actin patches (Pelham and Chang, 2001) and cell extracts in response to the Cdc42 GTPase (Ma et al., 1998; Mullins and Pollard, 1999) or the Listeria pathogen (Welch et al., 1997). Our results also suggest that the SCAR regulator mediates this Arp2/3-dependent actin polymerization in the blastoderm. A similar reduction in filamentous actin is observed in Dictyostelium Scar mutants (Bear et al., 1998), and budding yeast Bee1 is required for actin polymerization at actin patch structures in a permeabilized cell assay (Lechler and Li, 1997). Together, these results demonstrate a conserved role for the Arp2/3 complex and WASp/Scar proteins in promoting actin polymerization in vivo as well as in vitro.
In budding and fission yeast, inducible disruption of Arp2/3 complex function first leads to a cessation of actin patch movement followed by their eventual dissolution (Winter et al., 1997; Pelham and Chang, 2001). Therefore, the Arp2/3 complex is required for the motility of actin structures and their formation. Similarly, Dictyostelium Scar mutants exhibit a selective disruption of specific actin structures that cannot easily be explained by an overall reduction of actin. Actin correctly localizes to the cell cortex and extending pseudopods as in wild type; however, leading edge actin fails to coalesce in response to chemoattractant, often leading to the aberrant formation of multiple pseudopods (Bear et al., 1998). These results suggest that Scar is involved in the dynamic organization of actin structures as well as their generation.
An exciting possibility is that Scar and the Arp2/3 complex direct both the configuration and polymerization of actin in the Drosophila blastoderm. SCAR embryos in metaphase contain more than half the actin of wild-type embryos, yet this substantial amount of actin often fails to form even a discontinuous network of metaphase furrows. Instead, actin remains in aberrant surface structures that are not normally found at the surface of mitotic embryos. These observations suggest that SCAR plays a role in actin redistribution, perhaps through a local Arp2/3-dependent polymerization event that triggers a global cell cycledependent change in actin organization. This role of SCAR in the Drosophila embryo may be analogous to the reorganization of actin structures that occurs in other contexts, such as during cytokinesis or at the leading edge of migrating cells.
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Materials and methods |
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Germline clones were generated as described (Chou and Perrimon, 1996) by heat shock of hs-FLP; ovoD FRT40A/SCAR FRT40A larvae, hs-FLP; FRT82B ovoD/FRT82B Wsp larvae, or hs-FLP; ovoD FRT40A/Arpc1 FRT40A larvae. Adult germline clone females were mated to Oregon R males (blastoderm and oogenesis analysis) or to SCARk13811/CyO en-lacZ males (SCARmat/zyg) or Df(3R)3450/TM6B abdA-lacZ males (Wspmat/zyg) (CNS). Mosaic head clones were obtained in ey-FLP; Arpc1Q25sd FRT40A/l(2)cl-L31 FRT40A and ey-FLP; SCAR37 FRT40A/l(2)cl-L31 FRT40A flies. SCARk13811 and Wsp3 germline clones (blastoderm), Wsp3 germline clones (CNS), and all zygotic mutants were generated at 25°C. Arpc1R337st germline clones (blastoderm) and SCARk13811 germline clones (CNS) were generated at 2022°C.
We observed no contribution of zygotic gene activity to the blastoderm defects of embryos derived from SCARk13811 and Arpc1R337st germline clones (unpublished data). Wspmat embryos include embryos defective for both maternal and zygotic Wsp function and embryos defective only for maternal Wsp function.
cDNA expression
The full-length SCAR cDNA from the SD02991 EST was cloned as an EcoRV-XhoI (blunt) fragment into the pUASP vector (Rørth, 1998). Transgenic flies containing the UAS-SCAR transgene were generated by P-elementmediated transformation and two independent lines used for phenotypic rescue. Rescue to adult viability was obtained in SCAR/SCAR; UAS-SCAR/P{tubP-GAL4} for SCARk13811 and SCAR37 homozygotes.
SCAR antibody generation
The SCAR NH2-terminal region (amino acids 1237) was cloned into the pRSET vector (Invitrogen) to generate a 6xHis-tagged protein, which was purified and injected into guinea pigs (Cocalico Biologicals). SCAR polyclonal antibody was used at a 1:50 dilution to stain formaldehyde-fixed embryos as described (Theurkauf, 1994). SCAR polyclonal antibody was visualized with Alexa 546conjugated secondary antibody and costained with Alexa 488conjugated phalloidin (Molecular Probes).
P-element excision
The l(2)k13811 P-element insertion in the 5' UTR of SCAR was mobilized by introducing transposase on the chromosome CyO 2-3. Excisions were identified in F2 progeny derived from single males of the genotype w; SCARk13811/CyO
2-3. 15 homozygous viable alleles complemented the SCARk13811 lethality and are likely to represent precise excision events, verified by sequencing one such chromosome. 11 (presumably imprecise) excision alleles failed to complement the SCARk13811 lethality and were homozygous lethal. The
37 excision allele complements the lethality of l(2)06225, an insertion within the CG6105 transcription unit 400 bp upstream of SCAR (Fig. 1 A) and fails to complement the sterility of the downstream piwi gene.
Single larva PCR
SCAR37 homozygotes were chosen as nonfluorescing larvae from a SCAR
37/CyO, Act-GFP stock. Genomic DNA was prepared from five individual larvae, and PCR amplification for sequencing was performed using primers from the SCAR region.
CNS axon morphology
Embryos were fixed for 20 min in 3.7% formaldehyde/PBS:heptane and devitellinized in heptane:methanol. Embryos were stained with mouse mAb BP102 (1:10 dilution; Developmental Studies Hybridoma Bank) and rabbit antiß-galactosidase (1:1,500 dilution; Cappel), followed by fluorescence-conjugated Alexa 488 and Alexa 546 secondary antibodies (Molecular Probes), and mounted in Aquapolymount (Polysciences, Inc.). Images were z-series projections obtained on a ZEISS LSM 510 confocal microscope. CyO en-lacZ, TM6B abdA-lacZ, and TM3 Ubx-lacZ balancers were used to genotype embryos. For Arp3 heterozygotes, Arp3/balancer females were mated to WT males. When deficiencies were used, mutant/balancer females were mated to deficiency/balancer males. For double mutant analyses, double mutant females were crossed to deficiency/balancer or SCARk13811/balancer males. Statistics were computed using the Primer of Biostatistics program (Stanton Glantz) and Numerical Recipes in C (Press et al., 1992).
Actin quantitation
Embryos were collected at 2022°C (Arpc1mat and oskar control) or 25°C (SCARmat and oskar control), fixed for 45 min in 19% formaldehyde/PBS:heptane, and hand devitellinized. Control and mutant embryos were pooled and incubated in a single tube for 2 h with 6.6 nM Alexa 488conjugated phalloidin (Molecular Probes) and 0.1 µg/ml Hoechst (Sigma-Aldrich). Mean surface fluorescence intensity was measured for two areas of each embryo (the brightest surface 1.5 µm optical slice) and averaged. Images were obtained on a ZEISS LSM 510 confocal microscope using a C-Apochromatic 40x/1.2 NA water immersion objective. Images were generated using identical linear gain settings at zoom 3 and 1,024 pixel2 resolution to achieve Nyquist resolution, an optimal sampling rate. Gain settings were determined empirically to allow a range of intensities to be detected with minimal saturation of the higher control signal. Fluorescence intensity was quantitated in ImagePro Plus (Media Cybernetics). Background fluorescence was determined from 15 embryos processed identically but without phalloidin; these embryos displayed a nearly identical nonzero fluorescence, which was subtracted from the measured fluorescence in all images. Standard error of the mean was calculated in Kaleidagraph. In separate experiments to assess actin distribution (Fig. 5), embryos were costained with mouse antiß-tubulin antibodies (1:500 dilution; Sigma-Aldrich) for precise cell cycle staging. Antibody staining obscured the difference in phalloidin fluorescence; therefore, no antibodies were included in the quantitation experiment.
Egg chamber morphology
Ovaries were dissected from 35 d-old females and fixed for 15 min in 6% formaldehyde/PBS. Germline clones could first be identified after stage 8 of oogenesis (Spradling, 1993). Egg chamber microfilaments were visualized by staining with rhodamine-phalloidin (1 U/ml, 20 min; Molecular Probes), nuclei with OliGreen (1:5,000, 10 min, after a 1-h treatment with 5 µg/ml RNase; Molecular Probes), and ring canals with monoclonal Hts-RC antibody (clone 7C, 1:10 dilution).
SEM analysis of head cuticle and eye phenotypes
Adult heads underwent critical point drying and sputter coating with a gold film after dehydration in an ethanol series. Scanning EM was performed using a JEOL JSM-6400 microscope.
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Footnotes |
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Acknowledgments |
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J.A. Zallen was supported by a Koshland Postdoctoral Fellowship and is a Postdoctoral Fellow of the Damon Runyon-Walter Winchell Foundation Cancer Research Fund. E. Wieschaus is an Investigator of the Howard Hughes Medical Institute. E.D. Schejter is supported by grants from the Israel Science Foundation and the Minerva Foundation.
Submitted: 18 September 2001
Revised: 11 January 2002
Accepted: 14 January 2002
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