Article |
Address correspondence to Lynn Cooley, Department of Genetics, Yale University School of Medicine, P.O. Box 208005, New Haven, CT 06520-8005. Tel.: (203) 785-5067. Fax: (203) 785-6333. E-mail: lynn.cooley{at}yale.edu
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Abstract |
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Key Words: oogenesis; kelch; Src; actin binding; ring canal
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Introduction |
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In Drosophila melanogaster, 15 syncytial nurse cells and 1 oocyte are enveloped by a monolayer of somatic follicle cells and constitutes an egg chamber, the structural and functional unit of the Drosophila ovary (for review see Spradling, 1993). A ring canal is a gateway through which mRNAs, proteins, and nutrients flow from nurse cells into the oocyte during the entire course of oogenesis. Ring canals are derived from arrested mitotic cleavage furrows that are modified by the addition of several proteins. These include abundant F-actin (Koch and King, 1969), at least one protein that is recognized by antiphosphotyrosine antibodies (PY protein), a mucin-like glycoprotein (Kramerova and Kramerov, 1999), the Hts ring canal protein (HtsRC)* (Yue and Spradling, 1992; Robinson et al., 1994), ABP280/filamin (Li et al., 1999; Sokol and Cooley, 1999), Tec29 and Src64 tyrosine kinases (Dodson et al., 1998; Roulier et al., 1998), and Kelch (Xue and Cooley, 1993; Robinson and Cooley, 1997a).
As nurse cell cytoplasm transport proceeds, the diameter of ring canals grows from <1 µm to 1012 µm. This represents the addition of over one inch of filamentous actin during a period in which the filament density remains constant (Tilney et al., 1996). Near the end of oogenesis, the ring canal actin transforms from a single continuous bundle into several interwoven actin cables (Tilney et al., 1996). Ring canal expansion probably involves the nucleation of new actin filaments and an increase in actin filament length, coupled with filament reorganization that requires the establishment of reversible actin cross-links.
Previous work has shown that the Kelch protein is required for ring canal morphogenesis (Xue and Cooley, 1993; Tilney et al., 1996; Robinson and Cooley, 1997a). Ring canal actin in kelch mutant egg chambers is severely disorganized and partially occludes the lumen. This leads to a defect in cytoplasm transport and the production of small, sterile eggs (Xue and Cooley, 1993). Kelch is a multidomain protein (Fig. 1 A) and a member of a superfamily of proteins defined, in part, by the presence of six 50amino acid kelch repeats (KREPs). Based on sequence similarity to galactose oxidase, the KREP domain is predicted to fold into a six-bladed ß-propeller (Bork and Doolittle, 1994; Adams et al., 2000). In Limulus the KREP domain is present in at least three scruin proteins, each of which contains two KREP domains (Way et al., 1995). The KREP domains of -scruin each form an F-actin binding domain that allows
-scruin to act as an actin filamentcross-linking protein (Tilney, 1975; Bullitt et al., 1988; Sanders et al., 1996; Sun et al., 1997). Another KREP protein, Mayven, is found in human brain extracts and tightly colocalizes with F-actin in cultured human U373-MG astrocytoma/glioblastoma cells (Soltysik-Espanola et al., 1999). The second conserved domain in Kelch is the BTB/POZ (broad complex, tramtrack, and bric-á-brac; also known as the poxvirus and zinc finger domain) dimerization domain (Ahmad et al., 1998). The molecular makeup of the Kelch protein and the morphology of the kelch mutant ring canals suggest that Kelch could organize actin filaments by acting as a dimeric cross-linking protein (Robinson and Cooley, 1997a).
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Using a series of two-dimensional (2D) gel electrophoresis experiments, we determined that Kelch is phosphorylated in an SFK-dependent manner. We used site-directed mutagenesis to map the phosphorylated tyrosine residue. Thin section electron microscopy revealed striking differences in actin organization and filament number in lines expressing wild-type Kelch when compared with src6417 and the nonphosphorylatable form of Kelch. This showed that phosphorylation of Kelch is necessary for normal filament organization. Binding studies showed that the phosphorylated form of Kelch does not interact with actin. Therefore, Src64-mediated phosphorylation probably dissociates Kelch cross-links in ring canals. The nonphosphorylatable mutant also caused a reduction in actin monomer turnover kinetics. This suggests that reversible cross-links are required to allow dynamic actin monomer turnover and maintain overall ring canal morphology. These observations suggest that a major cytoskeletal target of Src64 signaling at the ring canal is the actincross-linking protein Kelch.
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Results |
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To examine Kelch phosphorylation in the Drosophila ovary, we performed 2D electrophoresis of ovary lysates from several genetic backgrounds. When wild-type ovary lysates were treated with phosphatase inhibitors, a tyrosine-phosphorylated protein that comigrated with one of two Kelch isoforms was detected (Fig. 2, A, C, and H). Antibodies to phosphoserine and phosphothreonine showed no immunoreactivity comigrating with Kelch (unpublished data). Comparison to pH standards showed that the shift observed between the two Kelch isoforms was equivalent to the addition of a single phosphate. In the absence of phosphatase inhibitors, Kelch migrated as a single spot with no corresponding phosphotyrosine staining (Fig. 2, B and D). Egg chambers dissected from src6417 homozygous flies did not contain the phosphorylated form of Kelch (Fig. 2 E). In the presence of phosphatase inhibitors, Kelch protein from kelDE1;P[kelY132A]/+ ovaries continued to be tyrosine phosphorylated (Fig. 2 F). However, Kelch tyrosine phosphorylation in kelDE1;P[kelY627A]/+ ovaries was absent (Fig. 2 G). To characterize the effects of phosphorylation on the ability of Kelch to bind actin, we performed an actin overlay experiment. Total ovary lysates from wild-type ovaries were separated using 2D electrophoresis, and the blots were incubated with F-actin and then actin antibodies to detect bound actin. Only nonphosphorylated Kelch (KelY627A) bound actin (Fig. 2 I). To verify that actin binding was due to Kelch, we tested ovary lysates from a kelch mutant and determined that there was no longer actin binding present in the area where Kelch protein would have focused (unpublished data). These results indicated that the tyrosine residue at position 627 was phosphorylated in a Src64-dependent manner, and phosphorylation of that residue disrupted Kelch binding to actin.
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We mapped the F-actin binding domain to a single KREP by expressing and purifying each individual repeat and then performing high speed centrifugation in the presence of F-actin. KREPs 14 and 6 failed to cosediment with F-actin (Fig. 3 D, repeat 4, lanes 1 and 2). KREP five was capable of binding F-actin in a saturable manner (Fig. 3 D, lanes 312). To address the effect of phosphorylation, we introduced aspartate or glutamate residues at position 627 to mimic phosphorylation (Jordan and Karess, 1997; Waites et al., 2001). Both substitutions disrupted the ability of KREP five to bind F-actin, resulting in >50% of each mutant repeat remaining in the supernatant after high speed centrifugation (Fig. 3 E, lanes 7 and 11). To demonstrate that the substitution made in the KelY627A protein would not affect actin binding, we tested the substitution in vitro. Neither alanine nor phenylalanine substitutions disrupted actin binding (Fig. 3 E, lanes 1320). These data suggest that F-actin binding by KREP five is likely to be reduced by phosphorylation of the tyrosine residue at position 627.
Characterization of KelY627A ring canal morphology
To study the role of Kelch phosphorylation in vivo, we examined kelch mutant flies expressing KelY627A protein in the germline. Wild-type stage 10A egg chambers stained with the Kel1B monoclonal antibody had a ring canal staining pattern that colocalized with F-actin (Fig. 4, A and A'). Egg chambers from kelch mutants showed a complete absence of Kelch protein staining (Fig. 4 B), and the well-characterized phenotype of ring canal actin disorganization with partial occlusion of the lumen (Tilney et al., 1996; Robinson and Cooley, 1997a) (Fig. 4 B'). The expression of one copy of P[kelY627A] in kelch mutants resulted in restoration of Kelch localization to ring canals and a rescue of F-actin organization (Fig. 4, C and C'), showing that the KelY627A protein had F-actin binding and cross-linking activity in vivo. The phenotype of KelY627A was not changed in a src6417 background (Fig. 4, D and D'). At a higher magnification, horizontal sections of wild-type ring canals were characterized by the appearance of two parallel rims of actin (Fig. 4 E). In kelch mutants, there was the typical collapse of actin into the lumen, almost completely obstructing the ring canal (Fig. 4 F). Comparison of wild type, kelDE1;P[kelY627A]/+, and src64
17 revealed an increase in the concavity of the actin rim in both mutants, causing the appearance of a bicycle rim shape as well as a decrease in the diameter of the ring canal (Fig. 4, compare E, G, and H). These observations suggested that the phenotype in kelDE1;P[kelY627A] closely resembled src64
17. Additionally, the introduction of two copies of P[kelY627A] into a wild-type background caused a decrease in ring canal diameter (Fig. 4 I). This dominant-negative phenotype indicated that the introduction of "irreversible" cross-links formed by KelY627A perturbed the function of endogenous Kelch.
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Actin monomer exchange decreases in KelY627A
To further characterize the phenotype caused by KelY627A protein, we performed the fluorescence recovery after photobleaching (FRAP) assay using actin tagged with green fluorescent protein (GFPactin). The expression of GFPactin was driven by a germline-specific promoter to label ring canals throughout all stages of development. FRAP was measured in stage 10A egg chambers from transgenic flies expressing wild-type Kelch (ORF1) and KelY627A. 18 P[UAS-GFPActin]/P[ORF1], 21 P[UAS-GFPActin]/P[kelY627A], and 15 P[UAS-GFPActin];src6417 ring canals were analyzed. Three representative experiments are shown (Fig. 8, AC). After bleaching, ring canals in P[UAS-GFPActin]/P[ORF1] recovered fluorescence with a t1/2 of 65 s (Fig. 8 D). In contrast, fluorescence in P[UAS-GFPActin]/P[kelY627A] ring canals recovered with a t1/2 of 227 s (Fig. 8 D). P[UAS-GFPActin];src64
17 had a t1/2 of 244 s. These data indicate that in ring canals with nonphosphorylatable Kelch or reduced Src64, actin monomers took >3.5 times as long to exchange as in control ring canals.
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Discussion |
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Regulation of ring canal expansion
The dynamics of actin filaments in ring canals have been elegantly described at the ultrastructural level (Riparbelli and Callaini, 1995; Tilney et al., 1996). Ring canals are built at the positions of arrested cleavage furrows that form during the mitotic divisions of germline cells. The mechanism of cleavage furrow arrest is likely to be conserved among animal species because incomplete cytokinesis occurs during the proliferation of germline cells in many animals (for review see Robinson and Cooley, 1996). In Drosophila, once egg chambers are fully assembled, ring canal growth happens in two phases. First, the thickness of the actin rim increases to 0.3 µm as the diameter of the ring grows slowly to 2 µm. Subsequently, the thickness of the actin rim and the density of actin filaments remain constant while the rate of ring canal expansion increases. The net increase of actin within ring canals overall is 134-fold (Tilney et al., 1996). During the rapid phase of ring canal growth, actin filaments must be polymerized, probably at the plasma membrane, to expand the ring canal rim, and disassembled at the cytoplasmic face to maintain the lumen. Our analysis of the Kelch protein shows that precise regulation of actin filament cross-linking by phosphorylation is critical during rapid ring canal growth.
The behavior of ring canals that contain KelY627A provide significant insight into Kelch function. The absence of Kelch phosphorylation leads to ring canals that accumulate more actin filaments than normal, possibly due to a slowing in the rate of actin depolymerization relative to the rate of polymerization. After about stage 8 of oogenesis, the failure to resolve the continuous sheet of actin filaments into discreet cables may be another consequence of inhibiting depolymerization. The presence of more "permanent" Kelch cross-links may reduce the accessibility of the filament network to depolymerizing factors. In vitro experiments have demonstrated that actincross-linking proteins alone are capable of inhibiting the rate of pyrenyl F-actin depolymerization (Cano et al., 1992). Another possible explanation for these phenotypes is that because Kelch cross-links are no longer easily reversible, filament reorientation or sliding is restricted during ring canal growth.
The FRAP experiments provide additional insight into the actin dynamics at the ring canal. First, ring canal actin is highly dynamic. The rate of actin monomer turnover that we found in wild-type ring canals is comparable to the kinetics of actin turnover found in the leading edge of motile goldfish epithelial keratocytes (Theriot and Mitchison, 1991). This would be consistent with a population of actin that is constantly undergoing a rapid cycle of polymerization and depolymerization. Second, the presence of nonregulated Kelch clearly results in a dramatic reduction in the dynamics of actin. This supports the model that mutant Kelch protein reduces accessibility to other actin-binding proteins, in this case proteins involved with polymerization or depolymerization. We propose that this effect could be due to bound Kelch acting as a stabilizing protein much in the same way that tropomyosin protects F-actin from actin depolymerizing factor/cofilin (for review see Pollard et al., 2000).
Studies in our lab involving the actin polymerization factor Arp2/3 have demonstrated that ring canal stability and growth is dependent on the presence of a functional Arp2/3 complex (Hudson and Cooley, 2002). The effects of mutations in Arp2/3 complex subunits are progressively more severe as egg chambers develop, and by stage 6, ring canals begin to collapse. In kelch null mutants, the actin filaments are initially well organized, begin to show signs of disorganization around stage 4, and are completely disorganized starting at stage 6 (Tilney et al., 1996; Robinson and Cooley, 1997a). Interestingly, thin section electron micrographs of kelDE1;P[kelY627A]/+ show signs of actin filament disruption beginning at stage 6. The coincidence of kelch and Arp2/3 complex mutant phenotypes with the onset of rapid ring canal expansion and the presence of highly dynamic actin, suggest a model where ring canal growth is powered by de novo actin polymerization accompanied by regulated cross-links. Therefore, ring canal growth may be mechanistically similar to the movement of plasma membranes at the leading edge of motile cells. Future work on ring canal actin organization should include platinum replica electron microscopy to understand the overall organization of the ring canal actin filament network. This will allow direct comparison to the actin filament networks of lamellipodia in Xenopus laevis keratocytes and fibroblasts (Svitkina and Borisy, 1999).
Intriguingly, the accumulation of actin during earlier stages of oogenesis is apparently independent of both Kelch and the Arp2/3 complex. Characterization of other mutants affecting ring canals has revealed genes required for initial stages of ring canal assembly. These include the cheerio gene that encodes the actin filamentcross-linking protein ABP280/filamin (Li et al., 1999; Sokol and Cooley, 1999). In cheerio mutants, ring canal actin is absent. In addition, HtsRC is required for the early accumulation of actin filaments (Robinson et al., 1994); however, it has not been determined whether HtsRC interacts directly with F-actin or affects actin polymerization. Therefore, additional research is needed to elucidate the mechanism of early ring canal biogenesis.
The regulation of Kelchactin cross-links could be accomplished by Src64 directly phosphorylating Kelch. Alternatively, Src64 may activate another protein tyrosine kinase, such as Tec29 (Roulier et al., 1998), which in turn phosphorylates Kelch. However, the shared phenotype seen by electron microscopy of the src64 and P[kelY627A] ring canals is strongly suggestive of Kelch being the major downstream component of a Src64 cascade. Analysis of Kelch phosphorylation in tec29 mutants is difficult because available tec29 alleles are lethal. SFKs have been shown to signal rearrangements in the actin cytoskeleton in other contexts. In Drosophila, embryos mutant for src64 or tec29 fail to complete epidermal closure at the end of gastrulation. This is, in part, because the leading edge cells contain reduced quantities of F-actin, and the cells only partially elongate and fail to migrate completely (Tateno et al., 2000). SFKs are also known to interact directly with cytoskeletal proteins, as in the case of c-Src and cortactin. Phosphorylation of cortactin by c-Src tyrosine kinase decreases its ability to cross-link F-actin in vitro (Bourguignon et al., 2001). These examples suggest that there could be a critical role played by tyrosine phosphatases to ensure that F-actin does not become disorganized due to excessive phosphorylation of cross-linking proteins. There are several candidate phosphatases in Drosophila; however, their roles in ring canal development have not been studied.
Kelch family members and actin
It should be noted that not all Kelch family members are actin-binding proteins (for review see Adams et al., 2000). For example, nuclear restricted protein/brain (NRP/B) is a novel nuclear matrix protein that contains a highly conserved KREP domain. NRP/B is specifically expressed in primary neurons and participates in the regulation of neuronal process formation (Kim et al., 1999). A direct interaction with actin by the ectoderm neural cortex-1 protein has been demonstrated by coimmunoprecipitation; however, it does not exclusively colocalize with F-actin in Daoy cells, and it is perinuclear in neuronal cell lines (Hernandez et al., 1997).
A Kelch family member that interacts with actin is called Mayven. Mayven localizes to the leading edge of the lamellipodia in U373-MG astrocytoma/glioblastoma cells (Soltysik-Espanola et al., 1999). Mayven is also localized with the focal adhesion kinase (Soltysik-Espanola et al., 1999), suggesting it could play a role in actin reorganization at focal adhesion plaques. A role for phosphorylation in the regulation of Mayven has not been reported.
In Limulus, it has been postulated that the Kelch homologue -scruin acts as a protein that allows F-actin to rapidly twist and slide during acrosome extension or "true discharge" (Sherman et al., 1999). Biochemical studies performed on
-scruin (Sun et al., 1997) have shown that the cysteine corresponding to Drosophila Kelch residue 628 lies within the
-scruin actin binding domain (Fig. 2 B, boxed). Thus, both Kelch and
-scruin contain an actin binding site within KREP number 5. However,
-scruin does not have a tyrosine comparable to Kelch residue 627 in the primary sequence; therefore, regulation of
-scruin cross-linking is likely to be different than that for Kelch.
-Scruin regulation may target scruinscruin interactions rather than scruinactin interactions.
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Materials and methods |
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The GFPactin fusion construct was made by subcloning the EcoRI/NotI GFPactin fragment from pUAST-GFPactin (Verkhusha et al., 1999) into pBluescript II KS(-). The 1.9-kb KpnI/NotI fragment from pBluescript was subcloned into pUASp2, a modified pUASp (Rorth, 1998).
GFPactin was then expressed in the ovary using nanos-Gal4-VP16 (Van Doren et al., 1998). The three genotypes used for FRAP assays were (1) P[UAS-GFPActin]/P[ORF1]: w1118;kelDE1,P[w+ nos-Gal4-vp16]/kelDE1;P[w+ ORF1]/P[w+ UASp2-GFPactin]; (2) P[UAS-GFPActin]/P[kelY627A]: w1118;kelDE1,P[w+ nos-Gal4-vp16]/kelDE1;P[w+ kelY627A]/P[w+ UASp2-GFPactin]; and (3) P[UAS-GFPActin]/src6417: w1118;P[w+ nos-Gal4-vp16]/P[w+ UASp2-GFPactin];src64
17/src64
17.
Electron microscope procedures
Egg chambers were dissected into IMADS (100 mM sodium glutamate, 25 mM KCl, 15 mM MgCl2, 5 mM CaSO4, and 2 mM sodium phosphate buffer, pH 6.9) (Singleton and Woodruff, 1994). Egg chambers were then fixed, stained, and embedded as previously described (Tilney et al., 1996). Detergent extraction and actin quantitation of ring canals were performed as previously described (Tilney et al., 1996). Negative staining of actin and Kelch mixtures was performed as previously described (Harris, 1999). Electron microscopy was performed on a Phillips Tecnai electron microscope at an accelerating voltage of 80 kV. Wild-type and mutant ring canal dimensions were compared using the t test, and a P value of <0.01 was taken as significant. The t test was performed using Microsoft Excel 2000.
Protein purification
Full-length Kelch and individual KREP constructs were made using amino acid numbers corresponding to the sequence in Xue and Cooley (1993). ORF1, the full-length Kelch cDNA (Robinson and Cooley, 1997a), was modified using PCR to introduce HindIII and NotI restriction enzyme sites at the 5' and 3' ends. The following individual KREPs were generated: repeat 1 (L405-K450), repeat 2 (V451-C497), repeat 3 (I498-L544), repeat 4 (L545-I593), repeat 5 (L594-L640), repeat 5 (L594-[Y627D]-L640), repeat 5 (L594-[Y627E]-L640), repeat 5 (L594-[Y627A]-L640), repeat 5 (L594-[Y627F]-L640), and repeat 6 (L641-M689). The PCR products were cloned into pCR 2.1-TOPO according to the manufacturer's instructions (Invitrogen). The constructs were sequenced and subcloned into the PinPoint Xa-3 vector (Promega) using HindIII and NotI. Escherichia coli DH5 cells were transformed with the PinPoint Xa-3 Kelch constructs. Soluble Kelch and KREP fragments were obtained by following the manufacturer's instructions. Only full-length Kelch constructs were cleaved from the purification tag using Factor Xa and the manufacturer's instructions.
Image acquisition and FRAP
GFP-actinexpressing ovaries were dissected onto a 22-mm x 40-mm glass coverslip as previously described (Theurkauf and Hazelrigg, 1998). FRAP experiments were conducted on a Zeiss 510 scanning laser microscope (Center for Cell Imaging, Yale University, New Haven, CT). Images were captured with either a 63x 1.4 NA objective or a 40x 1.3 NA objective using a pinhole diameter equivalent to one to two times the Airy disk diameter. In each experiment, at least 20 consecutive baseline images were obtained before bleaching. A region of the ring canal was selected, and fluorescence was bleached by scanning the region with high intensity illumination (100% transmittance). After photobleaching, fluorescence of the entire field was collected by the ArKr laser at 1% power every second for at least 4 min after photobleaching. The fluorescence intensity in the photobleached region of the ring canal was normalized to the fluorescence measured in a nonbleached region of the same ring canal, and in the cytoplasm. Wild-type and mutant rate constants of three curves were compared, and a P value of <0.01 was taken as significant. Nonlinear regression and t tests were performed using Microsoft Excel 2000. Images were prepared for publication using Adobe Photoshop.
Protein electrophoresis and Western blot analysis
Ovaries were dissected in the presence or absence of 25 mM sodium vanadate, 10 mM sodium fluoride, and 0.05% hydrogen peroxide in IMADS buffer and incubated on ice for 5 min. 10 ovaries were then solubilized with 25 µl isoelectric focusing sample buffer (8 M urea, 4% [wt/vol] CHAPS, 40 mM Tris, 65 mM DTE, and a trace of bromophenol blue). Samples were focused using Immobiline Drystrips, pH 47 linear (Amersham Pharmacia Biotech). The second dimension was performed using standard SDS-PAGE techniques on 8% gels and transferred onto nitrocellulose membranes (Hybond ECL; Amersham Pharmacia Biotech). Membranes were blocked with PBS containing 5% nonfat dry milk and incubated with the following antibodies and dilutions: 1:10 anti-kelch 1B (Xue and Cooley, 1993), 1:10 anti-actin JLA20 (Developmental Studies Hybridoma Bank, University of Iowa, Iowa City, IA), and 1:2,000 anti-phosphotyrosine PY20 (ICN Biomedicals); and detection of biotin-labeled Kelch proteins was performed using a 1:5,000 dilution of avidin-HRP (Pierce Chemical Co.). Secondary antibodies conjugated with HRP were purchased from Pierce Chemical Co. Visualization of HRP was performed using an ECL detection kit (Amersham Pharmacia Biotech).
Egg chamber staining
Ovaries were dissected, fixed, and processed as previously described (Robinson and Cooley, 1997a). For filamentous actin staining, egg chambers were incubated in 5 U of rhodamine-conjugated phalloidin (Molecular Probes) per 200 µl of PBTO (1 x PBS, 0.3% Triton X-100, 0.5% BSA). For antibody staining, ovaries were immunostained with anti-kelch 1B at a 1:1 dilution (Xue and Cooley, 1993), or anti-phosphotyrosine PY20 (ICN Biomedicals) at a 1:500 dilution. Secondary antibodies conjugated with Alexa Fluor 488 were purchased from Molecular Probes. Fluorescence intensity measurements were collected using the ZEISS image analysis software.
F-actin binding assays
Purified rabbit skeletal muscle monomeric actin was purchased from Cytoskeleton Inc. Actin concentration was estimated as previously described (Pollard, 1976). The concentration of Kelch and KREPs was determined using a modified Bradford assay (Bio-Rad Laboratories). G-actin was polymerized in A' buffer (10 mM imidazole, pH 7.0, 75 mM KCl, 2.5 mM MgSO4, 1 mM EGTA, 1 mM ATP, 0.1% NaN3). Immediately before use, purified Kelch or KREPs and actin were centrifuged at 100,000 g at 4°C for 1 h to pellet any aggregated protein. Mixtures of F-actin with Kelch or KREPs were incubated for 10 min at 25°C. Low speed and high speed cosedimentation assays were performed as previously described (Matova et al., 1999).
Far Western blot analysis
For binding experiments, total ovary lysates were separated using 2D electrophoresis as described above. Nitrocellulose blots were incubated at 4°C overnight in in PBS containing 5% milk and 0.1% Tween 20, followed by incubation with 10 mg of F-actin (Sigma-Aldrich) for 2 h at 4°C. The blots were washed, and a 1:10 dilution of rabbit anti-actin antibody JLA20 was added for 1 h. After washing, HRP-conjugated antirabbit antibody (Pierce Chemical Co.) was added for 1 h. Bands were visualized using ECL reagents.
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Footnotes |
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Acknowledgments |
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This research was supported by National Institutes of Health grant GM52702 to L. Cooley.
Submitted: 11 October 2001
Revised: 13 December 2001
Accepted: 14 January 2002
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References |
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