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Correspondence to Yosef Gruenbaum: gru{at}vms.huji.ac.il
Barrier to autointegration factor (BAF) binds double-stranded DNA, selected histones, transcription regulators, lamins, and LAP2–emerin–MAN1 (LEM) domain proteins. During early Caenorhabditis elegans embryogenesis, BAF-1 is required to organize chromatin, capture segregated chromosomes within the nascent nuclear envelope, and assemble lamin and LEM domain proteins in reforming nuclei. In this study, we used C. elegans with a homozygous deletion of the baf-1 gene, which survives embryogenesis and larval stages, to report that BAF-1 regulates maturation and survival of the germline, cell migration, vulva formation, and the timing of seam cell fusion. In the seam cells, BAF-1 represses the expression of the EFF-1 fusogen protein, but fusion still occurs in C. elegans lacking both baf-1 and eff-1. This suggests the existence of an eff-1–independent mechanism for cell fusion. BAF-1 is also required to maintain the integrity of specific body wall muscles in adult animals, directly implicating BAF in the mechanism of human muscular dystrophies (laminopathies) caused by mutations in the BAF-binding proteins emerin and lamin A.
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Structural roles were also seen in Xenopus laevis egg extracts, in which nuclei can assemble in vitro; adding recombinant BAF either inhibited or enhanced nuclear assembly, depending on the amount of BAF added (Segura-Totten et al., 2002), suggesting important roles for BAF in organizing chromatin and the nucleus. Indeed, BAF is essential in both C. elegans (Margalit et al., 2005) and Drosophila melanogaster (Furukawa et al., 2003). BAF-null Drosophila die at the larval-pupal transition, when they run out of maternally deposited BAF, with phenotypes that include arrest at various stages of the cell cycle, chromatin clumping, abnormal lamin distribution, aberrant nuclear morphology, small brains, and missing imaginal discs (Furukawa et al., 2003). RNAi-mediated down-regulation of baf-1 in C. elegans revealed that the loss of both maternal and zygotic BAF-1 caused anaphase chromatin bridges, abnormal chromatin morphology, and chromosome missegregation as early as the two-cell stage, and embryos died at or before the 
100-cell stage (Zheng et al., 2000; Margalit et al., 2005). The few embryos that escaped baf-1(RNAi) lethality grew into sterile adults with misplaced distal tip cells and gonads (Margalit et al., 2005).
To better understand the potential cellular and developmental roles of BAF, we studied a loss of function mutation in baf-1(gk324). In animals homozygous for the gk324 allele, the maternal supply of BAF-1 was sufficient to allow these animals to complete embryogenesis and larval stages, bypassing BAF-1's mitotic roles and allowing us to focus on later stages of C. elegans development. This genetic analysis reveals several novel tissue-specific roles for BAF-1 and sheds new light on the disease mechanisms of Emery-Dreifuss muscular dystrophy, which is caused by mutations in each of BAF's direct binding partners emerin and lamin A.
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baf-1–null animals have diverse tissue-specific phenotypes
The VC699 strain contains the gk324 allele in which the baf-1 promoter and ORF are deleted (Fig. S1 A, available at http://www.jcb.org/cgi/content/full/jcb.200704049/DC1). This allele was outcrossed three times and balanced with the hT2 balancer, which carries an integrated pharyngeal GFP element. PCR analysis was used to confirm the 766-bp deletion in the gk324 allele (Fig. S1 B). baf-1 mRNA was undetectable by RT-PCR analysis of RNA from 4-d-old gk324/gk324 homozygous animals (Fig. S1 C). Controls showed that an unrelated transcript encoded by cah-3 was detected at similar levels in homozygous, heterozygous, and wild-type animals (Fig. S1 C). BAF-1 protein was not detected in Western blots of extracts from 4-d-old gk324/gk324 animals, whereas their Ce-lamin signal was comparable with wild type (Fig. S1 D). Heterozygous gk324/hT2 animals were indistinguishable from wild type (N2), with similar body size and normal brood size (unpublished data) despite their 40% reduced levels of BAF-1. In contrast, homozygous gk324/gk324 animals, which apparently used maternally supplied BAF-1 to complete embryogenesis and larval stages, were 50% thinner and 35% shorter and arrested at late L4/early adult stage with several tissue-specific phenotypes, as detailed in the following sections. The short/thin phenotype could be the result of normal numbers of smaller cells, fewer cells, and/or abnormal gonads.
BAF-1 is required to prevent the premature fusion of seam cells to the epidermis
Homozygous gk324/gk324 animals frequently exploded when touched, suggesting cuticle defects. Worms contain two syncytial rows of seam cells that interrupt the hypodermis and form alae on the cuticle surface during certain developmental stages (Fig. 3 A).
During the L1–3 stages, the epithelial seam cells positioned along each side of the worm proliferate to create two cell types: epidermal daughter cells that fuse to the main body hyp7 hypodermis and ectoblastic daughter cells that remain unfused (Fig. 3 A; Sulston and Horvitz, 1977). During the L4 stage, these unfused ectoblastic seam cells, which extend from the tail to the head, fuse to each other laterally to form two syncytia on each side of the animal that eventually create cuticular structures (Fig. 3 A; Podbilewicz and White, 1994). To visualize the borders of the ectoblastic seam cells, we crossed the apical junction marker AJM-1–GFP into heterozygous gk324/hT2 worms (Shemer et al., 2000) and examined progeny that were homozygous or heterozygous for the baf-1 deletion. Until the late L3 stage, seam cells in both homozygous and heterozygous worms behaved like wild-type animals, forming one row of 12 cells on each side of the worm (unpublished data). In heterozygous gk324/hT2 animals, the ectoblastic seam cells remained unfused at the late L3/early L4 stage (Fig. 3 B, a; arrows), which is similar to wild-type animals (Podbilewicz and White, 1994). In contrast, the ectoblastic seam cells of homozygous gk324/gk324 animals fused prematurely with the epidermis, which was visualized by loss of the GFP signal at the apical borders of the seam cells (Fig. 3 B, b; arrows). Differential interference contrast (DIC) analysis revealed that at L3, after ectopic fusion, the number of seam cells remained at 12 (unpublished data). By late L4/early adult stage, all seam cells had fused to the epidermis instead of to each other in all tested worms (n = 20; Fig. 3 B, c; arrows). DIC analysis showed that after that stage, the seam cells disappeared. These cells did not stain for SYTO 11 (Invitrogen; Fridkin et al., 2004), indicating that their disappearance probably did not involve apoptosis (unpublished data). In addition, DIC analysis revealed no apoptotic bodies. This phenotype was specific for BAF-1 because in gk324/gk324 animals that expressed both GFP–BAF-1 and AJM-1–GFP transgenes, the seam cells remained unfused until the late L4 stage, as in heterozygous worms, and later fused correctly to each other to form lateral syncytia (Fig. 3 B, d; arrow). We conclude that BAF-1 is required to prevent the premature fusion of seam cells to the epidermis.
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BAF-1 is required for vulva formation
During the L3 stage, the vulva begins to form in the ventral epidermis when the anchor cell induces three (P5.p, P6.p, and P7.p) of the six vulval precursor cells (VPCs) to divide and to adopt vulval fates (Sharma-Kishore et al., 1999; Sommer, 2001). Strikingly, all gk324/gk324 animals were vulvaless (n = 50; Fig. 6 D; normal vulva in the control worm is indicated by an arrow in Fig. 6 C).
To determine whether the lack of vulva formation is caused by a missing anchor cell, we introduced the anchor cell–specific marker CDH-3–GFP (Pettitt et al., 1996) into gk324/hT2 worms. During the L3 and L4 stages, CDH-3–GFP expression was detected in the anchor cell in both baf-1 heterozygous and homozygous animals (Fig. 6, A–D). We next followed the VPCs in the offspring of AJM-1–GFP;gk324/hT2 animals and wild-type worms expressing AJM-1–GFP. In early L3 stage, the six VPCs, P3.p to P8.p, were present in both wild-type and baf-1 homozygous gk324/gk324 worms (Fig. 6, E and F; arrows). During the L3 and L4 stages, the VPCs continued to divide in wild-type worms and formed a normal vulva structure (Fig. 6 G; arrow indicates vulva at the late L3 stage). However, in 73% of gk324/gk324 worms, the second VPC division did not occur, and, in 27% of the worms, only one division occurred, usually that of P6.p (Fig. 6 H, arrowhead). During the L4 stage, all VPCs degenerated but did not stain for SYTO 11 (unpublished data). The vulvaless phenotype of gk324/gk324 animals was specific to the loss of BAF-1 expression because vulva formation was fully rescued in strains expressing the GFP–BAF-1 transgene, as seen in L4 larvae (Fig. 6, I and J; arrows) and adults (Fig. 6 K, arrow). We conclude that BAF-1 is required for VPC divisions and vulva formation either at the level of anchor cell signaling, defects in responding cells, or both.
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The disorganization of Ce-lamin in the germ cell nuclei of gk324/gk324 animals suggested potential gross defects in nuclear architecture. Therefore, we used the thin section transmission electron microscopy method to visualize the nuclear membranes and chromatin of germ cells derived from gk324/gk324 and control animals (Fig. 9). Gonads of control gk324/hT2 animals had normal-appearing nuclei (Fig. 9 A, arrows) and were indistinguishable from wild-type (N2) gonads (not depicted). In contrast, the gonads of gk324/gk324 animals had only a few large nuclei (Fig. 9 B, arrowhead), whereas most nuclei were small and lobulated (Fig. 9 C, arrows). About 65% of these small nuclei (n = 26) had gaps in their nuclear envelope (Fig. 9 C, arrowheads), and 27% had extra layers of nuclear membranes (Fig. 9 D, arrow). In some cases, the chromatin was condensed in electron-dense patches (Fig. 9 D, arrowheads). We speculate that the more drastic phenotype of gapped nuclear envelopes seen in germline cells might be caused by proliferation-linked defects in nuclear assembly. Thus, for germline cells, we conclude that BAF-1 is required not only to organize Ce-lamin but also for nuclear envelope and chromatin organization. Loss of nuclear envelope integrity could account for germline failure, but our results did not distinguish whether this integrity was lost during germ cell mitotic proliferation, meiosis, interphase, or a combination of these events.
Loss of BAF-1 causes rapid deterioration of body and tail muscles
An unexpected finding was that baf-1–null animals had an uncoordinated (unc) phenotype. When grown at 20°C, the movement of gk324/gk324 animals was similar to that of gk324/hT2 or wild-type animals up to day 4 (Video 1, available at http://www.jcb.org/cgi/content/full/jcb.200704049/DC1). From day 5 onwards, all homozygous gk324/gk324 animals (n = 146) developed an uncoordinated movement (Video 2 shows homozygous animals at day 5). In addition, a gradually increasing fraction of animals became paralyzed in the mid- and tail regions but not the head (Video 3 shows homozygous animals at day 12). The fraction of gk324/gk324 animals that could only move their head was 2.4% on day 7, increasing to 21.7% on day 11 and 88.5% on day 18 (n = 130, n = 104, and n = 76, respectively). Control gk324/hT2 animals did not develop uncoordinated movement, and the fraction of paralyzed animals was 0%, 0%, and 32.6% on days 7, 11, and 18, respectively (n = 144, n = 108, and n = 49, respectively). The paralysis phenotype of the gk324/hT2 controls differed from the gk324/gk324 animals because the control worms could still move their tail and were classified as class C aging animals, in which worms do not move even after prodding and can only move their head and/or tail or twitch in response to touch (Herndon et al., 2002). Both the uncoordinated and paralysis phenotypes of the gk324/gk324 animals were completely rescued by the GFP–BAF-1 transgene (unpublished data).
The Unc and paralyzed phenotypes could be caused either by nerve degeneration or muscle cell–intrinsic deterioration. To test the latter possibility, we used the thin section transmission electron microscopy method to examine the morphology of muscle cells in the head and tail regions of wild-type (N2), heterozygous gk324/hT2, and homozygous gk324/gk324 animals grown at 20°C on days 4, 8, and 12. In both control gk324/hT2 and homozygous gk324/gk324 animals, the head muscle tissues were similar to wild-type animals in each age group (Fig. 10, A–F). Muscle cells in the tail region of gk324/hT2 animals had normal morphology in all examined days, including day 12 (Fig. 10, G–I). In striking contrast, muscle cell morphology in gk324/gk324 animals was normal only at day 4 (Fig. 10 J) and deteriorated considerably by days 8 and 12 (Fig. 10, K and L). By day 12, the thin and thick filaments in the tail muscles became misorganized and contained dark aggregates (Fig. 10 L, arrows), which appeared already in day 8 (Fig. 10 K, arrow). Tail muscle cells in homozygous worms also had deteriorated nuclei (unpublished data). A previous study had shown that bodywall muscle cells are necessary for the normal distribution of myotactin, a protein that maintains the association between the muscle contractile apparatus and hypodermal fibrous organelles (Hresko et al., 1999). To further study the muscle deterioration, we immunostained gk324/hT2 and gk324/gk324 worms at day 12 with MH46, an antibody against myotactin. Myotactin distribution was normal at the tail and head of gk324/hT2 (Fig. S2, A and C; available at http://www.jcb.org/cgi/content/full/jcb.200704049/DC1) and at the head of gk324/gk324 worms (Fig. S2 B). However, myotactin distribution at the tail of gk324/gk324 worms was abnormal (Fig. S2 D). We conclude that BAF-1 is required to maintain the integrity of specific muscles in the body.
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60% the normal level of BAF-1 protein, were apparently normal with brood sizes similar to wild-type animals (unpublished data). Furthermore, two extra copies of the GFP–BAF-1 transgene (seen in the YG1001-3 strains) also had no apparent effects, although protein levels were not measured. Thus, in contrast to nuclear assembly extracts derived from Xenopus eggs, C. elegans may tolerate a wider range of BAF-1 protein levels. The mitotic and chromatin phenotypes seen in baf-1(RNAi) C. elegans embryos (Margalit et al., 2005) were seen to a limited extent at later stages of development, specifically in germ cells and in the VPCs, which might reflect a failure to assemble nuclei after mitosis. However, the successful embryonic development of homozygous gk324/gk324 embryos indicates that maternally contributed BAF-1 is sufficient for early development, when most somatic nuclear divisions occur. In this respect, our findings are consistent with findings in Drosophila, wherein maternal D-BAF was sufficient to complete all larval stages in flies homozygous for a baf deletion (Furukawa et al., 2003). At later stages of Drosophila development, the D-BAF deletion caused defects in chromatin organization, including clumped heterochromatin, which is similar to our findings for germ cell nuclei of baf-1–null C. elegans.
Perhaps there were defects in anaphase at postembryonic cells, but we could not see anaphase bridges in any cells examined either by DAPI staining, thin section EM, or DIC microscopy. The lack of anaphase chromatin bridges in somatic nuclei of post-L2 gk324/gk324 animals can be explained by the fact that most somatic cells are not dividing or by the activity of checkpoints that block entry into mitosis. Consistent with the first possibility, we saw severe nuclear morphology defects in proliferating germline cells after the L3 stage. Furthermore, the lack of mitotic figures in germline cells was consistent with the activity (in germline cells) of one or more checkpoints that are inactive in embryos (Encalada et al., 2005). We cannot rule out that many of the phenotypes of baf-1 mutants, including the lack of proliferation of germ cells and VPCs and the thin and uncoordinated phenotype, could be the results of defects in overall postembryonic cell divisions. On the other hand, it is likely that most phenotypes are probably caused by the aberrant regulation of BAF-1–regulated genes because these phenotypes were either absent or different in worms containing mutations in the cell cycle genes (Boxem et al., 1999; Fay and Han, 2000). In addition, the specific binding of BAF-1 to the eff-1 promoter confirms that BAF-1 is directly involved in gene regulation, as suggested previously in mammalian retinal cells (Wang et al., 2002). Given the differences in the timing of various developmental stages in C. elegans and Drosophila and the uncertain rate of loss of maternally provided BAF-1 protein, we are struck by the general similarities between the null phenotypes of these two organisms. In both organisms, BAF is required for efficient mitosis, chromatin organization, and nuclear envelope formation and also has partners involved in regulating tissue-specific roles during development. These results strongly support the hypothesis that many of BAF's roles are conserved in evolution. Our results suggest that BAF has additional partners involving the regulation of tissue-specific functions that remain to be discovered.
BAF-1 blocks the premature fusion of seam cells
In baf-1–null animals, the seam cells fused prematurely at the L3 stage. Most known genes involved in fusion and patterning of the epidermis are transcription factors (Podbilewicz, 2006). Ectopic and premature fusion of seam cells is also seen in worms deleted for ceh-16/engrailed (Cassata et al., 2005), which represses the transcription of a key gene, eff-1, encoding a cell surface protein that directly mediates most somatic cell fusion events in C. elegans (Shemer et al., 2004), including the ventral cell fusions required for vulva formation. The EFF-1 protein must be expressed in both cells for fusion to occur (Podbilewicz et al., 2006). Mutations in Ceh-16/engrailed derepress eff-1 and lead to abnormal fusion events during embryogenesis (Cassata et al., 2005). Ceh-16/engrailed normally blocks seam cell fusion to the syncytial hypodermis during embryogenesis. BAF is also involved in gene regulation. In mouse retinal cells, BAF inhibits gene expression by binding directly to Otx2, Crx, and other paired-like homeodomain transcription activators and blocks Crx-dependent gene expression in vivo (Wang et al., 2002). Our findings demonstrate that BAF-1 has more extensive roles in tissue-specific gene regulation because BAF-1 was required to prevent the premature fusion of seam cells. Our results suggest that BAF-1 normally represses eff-1 during embryogenesis and later stages of development by binding to the eff-1 promoter. It is also worth noting that BAF-1 is currently the only factor known to bind the eff-1 promoter directly.
Although eff-1 expression was altered in gk324/gk324 worms, premature seam cell fusion still occurred in worms homozygous for mutations in both eff-1 and baf-1. The seam cells in these worms are smaller and disappear later. Thus, the eff-1 mutant background may cause additional deleterious effects. We speculate that there are other proteins mediating cell fusion, which are repressed by BAF-1. One such protein is AFF-1, which has fusogenic activities that do not involve EFF-1 activity, especially in anchor cell fusion and fusion between the lateral seam cells (Sapir et al., 2007). Our ChIP analysis suggests that BAF-1 also binds the promoter of aff-1 and, therefore, prevents seam cell fusion by regulating both the aff-1 and eff-1 promoters (Fig. 4 I).
baf-1–null adults develop dystrophic muscles
One of the most intriguing phenotypes of animals homozygous for a baf-1 deletion was the accelerated deterioration of specific muscles in aging animals. This suggests a role for BAF-1 in adult muscle integrity. This finding is consistent with the idea that BAF is required to efficiently localize emerin at the nuclear envelope of mammalian cells (Bengtsson and Wilson, 2006) and Ce-lamin in C. elegans cells (this study). The observed deterioration of muscle cells in C. elegans differed from normal muscle aging (Herndon et al., 2002) in several ways. First, the baf-1–null muscular dystrophy–like phenotype appeared only in midbody and tail muscles (which are innervated by the ventral or dorsal nerve cords), whereas head muscles (which synapse to the nerve ring) remained functional. Consequently, the head muscles often behave differently. During normal aging, C. elegans suffers from loss of muscle mass and the function of muscles in the body wall and pharynx (Herndon et al., 2002). We cannot rule out the possibility that the midbody and tail muscle phenotypes result from problems with the ventral or dorsal nerve cords. However, we disfavor this model because the morphology of the affected muscles was visibly defective on day 8, when the gk324/gk324 animals could still move, and, therefore, nerve cells were still functional. Furthermore, similar phenotypes of the disorganization of muscle cell filaments and gradual paralysis were previously reported in worms with mutations in muscle-related genes such as unc-54 (myosin heavy chain; Tajsharghi et al., 2005), unc-27 (Troponin I; Burkeen et al., 2004), or unc-52 (perlecan; Mackenzie et al., 1978), which also influence DTC migration (Merz et al., 2003). These genes might be regulated directly or indirectly by BAF-1. In addition, muscle attachment to hypodermal cells was also aberrant in the tail of the gk324/gk324 animals. Therefore, we hypothesize that loss of baf-1 function directly disrupts the function of selected muscle cells in C. elegans.
Possible involvement of BAF in laminopathies
In humans, mutations in A-type lamins cause several forms of muscular dystrophy, including autosomal dominant Emery-Dreifuss muscular dystrophy, Limb-Girdle muscular dystrophy, and dilated cardiomyopathy with conduction system defects (Decostre et al., 2005). Mutations in emerin, an inner nuclear membrane LEM domain protein that directly binds to lamin A, cause X-linked recessive Emery-Dreifuss muscular dystrophy, which is clinically indistinguishable from autosomal dominant Emery-Dreifuss muscular dystrophy (Decostre et al., 2005). Both proteins (lamin A and emerin) directly interact with BAF (Holaska et al., 2003), and these interactions are conserved in C. elegans: Ce-emerin and Ce-lamin (B type) each bind BAF-1 directly (Lee et al., 2001; Liu et al., 2003; unpublished data). The selective muscular dystrophy–like phenotype of baf-1–null animals strongly suggests a novel role for BAF in muscle cell integrity, potentially at the level of muscle-specific gene regulation. Future work will aim to identify putative BAF-regulated genes in muscles to shed new light on the mechanisms of Emery-Dreifuss muscular dystrophy and other laminopathies.
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Microscopy and live cell imaging
Transmission electron microscopy analysis of C. elegans was performed as described previously (Haithcock et al., 2005). DIC and immunofluorescence images were taken either with a CCD camera (Axiocam; Carl Zeiss MicroImaging, Inc.) mounted on a microscope (Axioplan II; Carl Zeiss MicroImaging, Inc.) equipped for fluorescence and DIC or with a confocal scanhead (MRC-1024; Bio-Rad Laboratories) coupled to an inverted microscope (Axiovert 135M; Carl Zeiss MicroImaging, Inc.) equipped with a 63x NA 1.3 oil immersion objective (Carl Zeiss MicroImaging, Inc). For FRAP analysis, gk324/gk324 worms expressing GFP–BAF-1 were imaged using a confocal microscope (FV-1000; Olympus) equipped with an inverted microscope (IX81; Olympus) and a 60x NA 1.4 oil immersion objective (Olympus). GFP–BAF-1 fluorescence was photobleached by a 405-nm laser in a defined region of each cell and was imaged with a 488-nm laser line for excitation and a 505–525-nm filter for emission before, during, and after the photobleach. For FRAP analysis, fluorescence intensity in the bleached area, the backgroup area, and the total cell area were measured as a function of time after bleaching and were normalized essentially as described previously (Rabut and Ellenberg, 2005).
Antibodies, indirect immunofluorescence staining, and immunoblots
Adult C. elegans were fixed and stained by indirect immunofluorescence as described previously (Fridkin et al., 2004). MH46 (Francis and Waterston, 1991) was used at a 1:10 dilution. 135 4-d-old young adults were collected in 30 µl M9 buffer, mixed with 15 µl of 2x SDS sample buffer, boiled for 10 min, and subjected to protein blot analysis as described previously (Margalit et al., 2005).
ChIP
For extract preparations, N2, PS3729, YG1001, or YG1002 asynchronous population worms grown in six 9-cm plates were collected. Worms were washed twice with M9 and fixed with 2% formaldehyde for 30 min at room temperature, washed once with 100 mM Tris, pH 7.5, twice with M9 buffer, and once with homogenization buffer (50 mM Hepes/KOH, pH 7.5, 1 mM EDTA, 140 mM KCl, 0.5% NP-40, 10% glycerol, and 5 mM DTT with protease inhibitors), and frozen in liquid nitrogen. Worms were sonicated on ice 10 times for 30 s each with a sonicator (Sonic; Heat Systems Ultrasonic, Inc.) and centrifuged at 6,500 rpm for 20 min at 4°C. The supernatant was sonicated again to shear the DNA on ice five times for 30 s each and was centrifuged at 14,000 rpm for 20 min. The supernatant was collected and tested for the presence of GFP–BAF-1 or AJM-1–GFP by immunoblotting and frozen in liquid nitrogen. Lysates were incubated with 5 µg anti-GFP antibody (Roche) for 2 h, and cellular debris was removed by centrifugation at 6,500 rpm for 15 min at 4°C. Lysates were then centrifuged at 14,000 rpm for 10 min, and 50 µl of protein G–Sepharose (Roche) was added to the supernatant. Immunocomplexes were washed twice with each buffer: ChIP buffer (50 mM Hepes/KOH, pH 7.5, 1 mM EDTA, 0.5%, NP-40, and 5 mM DTT with protease inhibitors) with 100 mM KCl and ChIP buffer with 1 M KCl and Tris-EDTA. Complexes were eluted with elution buffer (1% SDS and 10 mM Tris-HCl, pH 8), and 16 µl of 5 M NaCl was added to the elution and heated at 65°C overnight. DNA was then isolated using a standard procedure (phenol-chloroform extraction) and was resuspended in 20 µl Tris-EDTA. The amount of eluted DNA was quantified using locus-specific primers. Quantitative PCR was used to monitor ChIP results. 20 µl of quantitative PCRs contained 1:2 SYBR green Mix (ABgene), 250 nM of each primer, and an appropriate amount of DNA. The quantitative PCR results were analyzed essentially as described previously (Wang et al., 2004).
PCR and RT-PCR analyses
Single-worm PCR analysis using primers 5'-AACCGAAATTCTCAGCCCTT-3' and 5'-GATCGCGGCCGCCTTAGAAACACTCTTCAGGATCG-3' to distinguish between wild-type, gk324/hT2, and gk324/gk324 worms (Fig. S1, A and B) was performed essentially as described previously (Williams et al., 1992). For RT-PCR, 100 wild-type (N2), gk324/hT2, or gk324/gk324 worms were collected from each strain in 700 µl of extraction mixture (0.1 M NaOAc, 50% phenol, 2 M guanidinium thiocyanate, 12 mM sodium citrate, pH 7.0, 0.25% Sarkosyl, and 50 mM ß-mercaptoethanol), immediately frozen in liquid nitrogen, and incubated at –80°C for at least 20 min. Samples were thawed at 60°C, vortexed, and incubated for 1 min on ice. Total RNA was isolated using a standard procedure (phenol extraction) and digested with RNase-free DNase I (Promega), and cDNA was synthesized from 400 ng RNA using a 15-nt oligodT primer using Moloney murine leukemia virus reverse transcriptase (Promega) according to the manufacturer's instructions. From each 20 µl cDNA, 2 µl was analyzed by PCR using the following primers: baf-1 forward (5'-GATCGAATTCATGTCGACTTCTGTTAAGCATCG-3'), baf-1 reverse (5'-GATCGCGGCCGCCATGAACTGATCTGCCCACTCG-3'), cah-3 forward (5'-CACTTCCATTGGGGAGAGAA-3'), and cah-3 reverse (5'-ACAACGCCTTTCCCTCTTTT-3').
Worm movement assays
N2, VC699, and gk324/gk324-expressing GFP–BAF-1 worms were collected, washed with M9 buffer, treated for 5 min with hypochlorite solution (1.1% hypochlorite and 0.62 M NaOH), and washed with M9, and the embryos were collected and grown on nematode growth medium plates at 16°C for 3 d until they reached the L4/young adult stage. The VC699 and gk324/gk324 worms that expressed GFP–BAF-1 were sorted into two classes representing the gk324/hT2 and gk324/gk324 genotypes based on the presence of GFP fluorescence in the pharynx. For each experiment, 120 worms from each line were transferred to new nematode growth medium plates at 20°C (40 worms per plate), and movement was classified essentially as described previously (Henderson et al., 1997). Worms were also processed for transmission electron microscopy at days 4, 8, and 12 from synchronized embryos.
Online supplemental material
Fig. S1 describes the gk324 allele and provides evidence that baf-1 is not expressed in L4 worms homozygous for the gk324 allele. Fig. S2 shows that muscle attachment to hypodermal cells is aberrant in the tail region of gk324/gk324 animals by staining gk324/gk324 animals with MH46 antibody. The supplemental text gives further details on primers that were used in this study. Videos 1–3 show movement of heterozygous and homozygous worms for the gk324 allele at days 5 and 12. Online supplemental material is available at http://www.jcb.org/cgi/content/full/jcb.200704049/DC1.
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Acknowledgments |
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This work was supported by the USA-Israel Binational Science Foundation, Israel Science Foundation (ISF), Israel Ministry of Health, European Union's FP6 Life Science, Genomics, and Biotechnology for Health (grant LSHM-CT-2005-018690 to Y. Gruenbaum), support from the ISF (grant to B. Podbilewicz), and support from NIH (grant RO1 GM48646 to K.L. Wilson).
Submitted: 10 April 2007
Accepted: 13 July 2007
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References |
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Bengtsson, L., and K.L. Wilson. 2006. BAF phosphorylation on Ser-4 regulates emerin binding to lamin A in vitro and emerin localization in vivo. Mol. Biol. Cell. 17:1154–1163.
Boxem, M., D.G. Srinivasan, and S. van den Heuvel. 1999. The Caenorhabditis elegans gene ncc-1 encodes a cdc2-related kinase required for M phase in meiotic and mitotic cell divisions, but not for S phase. Development. 126:2227–2239.[Abstract]
Brenner, S. 1974. The genetics of Caenorhabditis elegans. Genetics. 77:71–94.
Burkeen, A.K., S.L. Maday, K.K. Rybicka, J.A. Sulcove, J. Ward, M.M. Huang, R. Barstead, C. Franzini-Armstrong, and T.S. Allen. 2004. Disruption of Caenorhabditis elegans muscle structure and function caused by mutation of troponin I. Biophys. J. 86:991–1001.[Medline]
Cassata, G., G. Shemer, P. Morandi, R. Donhauser, B. Podbilewicz, and R. Baumeister. 2005. ceh-16/engrailed patterns the embryonic epidermis of Caenorhabditis elegans. Development. 132:739–749.
Chen, H., and A. Engelman. 1998. The barrier-to-autointegration protein is a host factor for HIV type 1 integration. Proc. Natl. Acad. Sci. USA. 95:15270–15274.
Decostre, V., R. Ben Yaou, and G. Bonne. 2005. Laminopathies affecting skeletal and cardiac muscles: clinical and pathophysiological aspects. Acta Myol. 24:104–109.[Medline]
Encalada, S.E., J. Willis, R. Lyczak, and B. Bowerman. 2005. A spindle checkpoint functions during mitosis in the early Caenorhabditis elegans embryo. Mol. Biol. Cell. 16:1056–1070.
Ercan, S., P.G. Giresi, C.M. Whittle, X. Zhang, R.D. Green, and J.D. Lieb. 2007. X chromosome repression by localization of the C. elegans dosage compensation machinery to sites of transcription initiation. Nat. Genet. 39:403–408.[CrossRef][Medline]
Fay, D.S., and M. Han. 2000. Mutations in cye-1, a Caenorhabditis elegans cyclin E homolog, reveal coordination between cell-cycle control and vulval development. Development. 127:4049–4060.[Abstract]
Francis, R., and R.H. Waterston. 1991. Muscle cell attachment in Caenorhabditis elegans. J. Cell Biol. 114:465–479.
Fridkin, A., E. Mills, A. Margalit, E. Neufeld, K.K. Lee, N. Feinstein, M. Cohen, K.L. Wilson, and Y. Gruenbaum. 2004. Matefin, a C. elegans germ-line specific SUN-domain nuclear membrane protein, is essential for early embryonic and germ cell development. Proc. Natl. Acad. Sci. USA. 101:6987–6992.
Furukawa, K., S. Sugiyama, S. Osouda, H. Goto, M. Inagaki, T. Horigome, S. Omata, M. McConnell, P.A. Fisher, and Y. Nishida. 2003. Barrier-to-autointegration factor plays crucial roles in cell cycle progression and nuclear organization in Drosophila. J. Cell Sci. 116:3811–3823.
Gorjánácz, M., E.P.F. Klerkx, V. Galy, R. Santarella, C. López-Iglesias, P. Askjaer, and I.W. Mattaj. 2006. C. elegans BAF-1 and its kinase VRK-1 participate directly in postmitotic nuclear envelope assembly. EMBO J. 26:132–143.[CrossRef][Medline]
Haithcock, E., Y. Dayani, E. Neufeld, A.J. Zahand, N. Feinstein, A. Mattout, Y. Gruenbaum, and J. Liu. 2005. Age-related changes of nuclear architecture in Caenorhabditis elegans. Proc. Natl. Acad. Sci. USA. 102:16690–16695.
Haraguchi, T., T. Koujin, M. Segura-Totten, K.K. Lee, Y. Matsuoka, Y. Yoneda, K.L. Wilson, and Y. Hiraoka. 2001. BAF is required for emerin assembly into the reforming nuclear envelope. J. Cell Sci. 114:4575–4585.[Medline]
Henderson, S.T., D. Gao, S. Christensen, and J. Kimble. 1997. Functional domains of LAG-2, a putative signaling ligand for LIN-12 and GLP-1 receptors in Caenorhabditis elegans. Mol. Biol. Cell. 8:1751–1762.[Abstract]
Herndon, L.A., P.J. Schmeissner, J.M. Dudaronek, P.A. Brown, K.M. Listner, Y. Sakano, M.C. Paupard, D.H. Hall, and M. Driscoll. 2002. Stochastic and genetic factors influence tissue-specific decline in ageing C. elegans. Nature. 419:808–814.[CrossRef][Medline]
Holaska, J.M., K.K. Lee, A.K. Kowalski, and K.L. Wilson. 2003. Transcriptional repressor germ cell-less (GCL) and barrier-to-autointegration factor (BAF) compete for binding to emerin in vitro. J. Biol. Chem. 278:6969–6975.
Hresko, M.C., L.A. Schriefer, P. Shrimankar, and R.H. Waterston. 1999. Myotactin, a novel hypodermal protein involved in muscle-cell adhesion in Caenorhabditis elegans. J. Cell Biol. 146:659–672.
Lee, K.K., T. Haraguchi, R.S. Lee, T. Koujin, Y. Hiraoka, and K.L. Wilson. 2001. Distinct functional domains in emerin bind lamin A and DNA-bridging protein BAF. J. Cell Sci. 114:4567–4573.[Medline]
Lee, M.S., and R. Craigie. 1998. A previously unidentified host protein protects retroviral DNA from autointegration. Proc. Natl. Acad. Sci. USA. 95:1528–1533.
Lehmann, R. 2001. Cell migration in invertebrates: clues from border and distal tip cells. Curr. Opin. Genet. Dev. 11:457–463.[CrossRef][Medline]
Lin, C.W., and A. Engelman. 2003. The barrier-to-autointegration factor is a component of functional human immunodeficiency virus type 1 preintegration complexes. J. Virol. 77:5030–5036.
Liu, J., T. Rolef-Ben Shahar, D. Riemer, P. Spann, M. Treinin, K. Weber, A. Fire, and Y. Gruenbaum. 2000. Essential roles for Caenorhabditis elegans lamin gene in nuclear organization, cell cycle progression, and spatial organization of nuclear pore complexes. Mol. Biol. Cell. 11:3937–3947.
Liu, J., K.K. Lee, M. Segura-Totten, E. Neufeld, K.L. Wilson, and Y. Gruenbaum. 2003. MAN1 and emerin have overlapping function(s) essential for chromosome segregation and cell division in C. elegans. Proc. Natl. Acad. Sci. USA. 100:4598–4603.
Mackenzie, J.M.J., R.L. Garcea, J.M. Zengel, and H.F. Epstein. 1978. Muscle development in Caenorhabditis elegans: mutants exhibiting retarded sarcomere construction. Cell. 15:751–762.[CrossRef][Medline]
Margalit, A., M. Segura-Totten, Y. Gruenbaum, and K.L. Wilson. 2005. Barrier-to-autointegration factor is required to segregate and enclose chromosomes within the nuclear envelope and assemble the nuclear lamina. Proc. Natl. Acad. Sci. USA. 102:3290–3295.
Margalit, A., A. Brachner, J. Gotzmann, R. Foisner, and Y. Gruenbaum. 2007. Barrier-to-autointegration factor - a BAFfling little protein. Trends Cell Biol. 17:202–208.[CrossRef][Medline]
Merz, D.C., G. Alves, T. Kawano, H. Zheng, and J.G. Culotti. 2003. UNC-52/perlecan affects gonadal leader cell migrations in C. elegans hermaphrodites through alterations in growth factor signaling. Dev. Biol.<
cells in heterozygous animals (C) but not in homozygous animals (D). (E–H) AJM-1–GFP expression in vulva precursor cells (VPCs) of wild-type (E and G) and gk324/gk324 (F and H) animals. At the early L3 stage, all six VPCs were visualized in both wild-type (E) and gk324/gk324 worms (F). In wild-type worms at late L3, VPCs formed a normal vulva structure (G; 18-cell stage). In most gk324/gk324 worms, the VPCs either did not divide at all (not depicted) or had a partial division of P6.p (arrowhead in H). Arrows in E–H indicate VPCs. (I–K) Vulva formation in baf-1–null homozygous animals is rescued by the transgenic expression of GFP–BAF-1, as shown at early L4 (arrow in I), late L4 (arrow in J), and young adult (arrow in K). Bars, 10 µm.
