INTRODUCTION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

Basement membranes are thin extracellular matrices that have important roles in the development and functions of many tissues. The major constituents of basement membranes include type IV collagens, laminins, nidogen (entactin), and proteoglycans. Interactions between these constituents and their interactions with cell surface molecules are critical for the assembly and function of basement membranes (Yurchenco, 1994; Timpl, 1996). Collagen IV and laminin can independently polymerize in a concentration-dependent manner to form separate networks in the basement membrane (Yurchenco and Furthmayr, 1984; Yurchenco et al., 1992; Yurchenco and Cheng, 1993). Connection between these independent networks is thought to be mediated by nidogen, which can bind both collagen IV and laminin with high affinity (Fox et al., 1991).

Nidogen is a 150-kDa glycoprotein that consists of two amino (G1, G2) and one carboxyl (G3) terminal globular domains that are connected by a rod domain composed primarily of endothelial growth factor (EGF) repeats (Durkin et al., 1988; Mann et al., 1989; Fox et al., 1991). Mammals express two closely related forms of nidogen, nidogen-1 and nidogen-2, that are encoded by separate genes (Kohfeldt et al., 1998). Both nidogen-1 and nidogen-2 bind type IV collagen, perlecan, and laminin with high affinity (Aumailley et al., 1993; Chung et al., 1993; Kohfeldt et al., 1998). The G3 domain of nidogen-1 binds strongly to a single EGF module in the laminin 1 chain (Mayer et al., 1993), whereas nidogen-2 shows strong interaction with LG domains of the laminin 2 chain (Kohfeldt et al., 1998; Talts et al., 1999). Binding of nidogen-1 to collagen IV and perlecan occurs through the nidogen G2 domain (Reinhardt et al., 1993). Using purified proteins, nidogen-1 was shown to form a ternary complex with type IV collagen and laminin-1 (Aumailley et al., 1993). The ability of nidogen to bind both the collagen IV and laminin networks with high affinity suggests that this linking function may be critical for basement membrane assembly.

Several lines of evidence suggest that nidogen-1 has important functions in vertebrate development and differentiation. Nidogen-1 promotes attachment of cells in culture, mediated through the single RGD sequence within its rod domain and by sequences in the G2 domain (Yelian et al., 1993; Dong et al., 1995; Wu et al., 1995). Addition of antibodies that interfere with nidogen-1 binding to laminin disrupts epithelial morphogenesis of cultured embryonic kidney, lung, or submandibular glands (Ekblom et al., 1994; Kadoya et al., 1997). Nidogen proteolysis has been correlated with loss of epithelial function during normal mammary gland involution and during gland regression induced by ectopic stromelysin-1 expression (Alexander et al., 1996). Nidogen-1 can cooperate with laminin-1 to stimulate -casein synthesis by cultured mammary epithelial cells (Pujuguet et al., 2000). These studies suggest that nidogen may have critical developmental functions in vivo.

To examine the in vivo functions of nidogen, we initiated studies of its roles in Caenorhabditis elegans development. Several basement membrane components have been shown to be essential for embryonic development in C. elegans (Kramer, 1997). Loss or alteration of type IV collagen (Guo et al., 1991; Sibley et al., 1993; Gupta et al., 1997), perlecan (Rogalski et al., 1993), or SPARC/osteonectin (Fitzgerald and Schwarzbauer, 1998) result in arrest during the mid to late stages of embryogenesis. Mutations in other basement membrane components that are required for the proper assembly of collagen IV, perlecan, or SPARC might be expected to also result in defective development.

The mechanisms of assembly of these molecules into basement membranes may differ. Perlecan is expressed in muscle cells and assembles into the basement membranes immediately adjacent to the cells that synthesize it (Moerman et al., 1996). In contrast, type IV collagen (Graham et al., 1997) and SPARC (Fitzgerald and Schwarzbauer, 1998) assemble into basement membranes associated with tissues that do not express the proteins. Epitope-tagged type IV collagen synthesized in body wall muscle cells can assemble into basement membranes that surround the pharynx, intestine, and gonad. Expression only in muscle cells was also shown to be sufficient to rescue the lethality of a type IV collagen null mutant and to provide modest fertility (Graham et al., 1997). In addition, type IV collagen is localized to certain basement membranes in C. elegans and absent from others, even though all are open to the pseudocoelom, which is the source of free collagen IV molecules. These studies indicated that a mechanism for directing type IV collagen assembly to particular locations must exist.

Laminin appears earlier than type IV collagen during mouse (Leivo et al., 1980; Dziadek and Timpl, 1985) and Drosophila development (Kuschegullberg et al., 1992) and during formation of new basement membranes in angiogenesis (Form et al., 1986). In a coculture of epithelial and mesenchymal cells, antisense inhibition of laminin expression blocked deposition of type IV collagen and nidogen at the cellular interface, indicating the importance of laminin for basement membrane assembly (De Arcangelis et al., 1996). Because nidogen can serve as a link between type IV collagen and laminin, it is possible that collagen IV could be localized by binding to nidogen that is associated with laminin already present at tissue surfaces. Thus, the earlier localized laminin could provide a pericellular binding site to concentrate collagen IV and allow its assembly. Nidogen would be required to meditate this interaction because collagen IV and laminin have not been found to directly interact.

To determine what roles nidogen might play in C. elegans development and whether it is involved in localization of type IV collagen assembly, we generated mutations in the C. elegans nidogen gene. Here we present characterization of the C. elegans nidogen gene nid-1 and analysis of mutations in it.

	MATERIALS AND METHODS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

Characterization of the nid-1 Gene

A C. elegans nidogen homologue was identified in the cosmid F54F3 (GenBank Z79696). All numbering presented is based on this GenBank sequence. Sequence and restriction analyses of RT-PCR products derived from predicted exons and six existing cDNA clones identified three alternative splice variants of the nid-1 gene. Three of the cDNAs (yk38f9, yk64d8, yk152d12) contain all exons and encode NID-1 form A, one (yk7a9) is missing exon 11 and encodes form B, and two (yk33c12, yk161f6) are missing exons 11-14 and encode form C. All other predicted intron/exon boundaries were confirmed by sequence analyses of these clones.

nid-1 Mutations

Animals were maintained as described by Brenner (Brenner, 1974). Isolation of nid-1 mutations generally followed previously described methods (Plasterk, 1995). A transposon Tc1 insertion allele of nid-1, ev608::Tc1, was identified from a library of animals carrying mut-2(r459). The Tc1 insertion site was localized to between nucleotides 3041-3042 by sequence analysis. Tc1 excision deletion alleles were identified by PCR screening populations of mut-2(r459); nid-1(ev608::Tc1) animals using primers flanking the Tc1 insertion site. Homozygous cg118 and cg119 deletion mutant animals were isolated by sib-selection. The extents of the deletions were determined by sequencing PCR-generated clones spanning the deletions. All mutations were outcrossed a minimum of seven times before further analyses.

Production of Anti-NID-1 Antiserum

A GST-NID-1 fusion construct was generated by cloning the 1.3-kb NciI-XhoI (nucleotides 7820-9521 genomic) fragment from the cDNA yk38f9 (D36924, D33953) into the pGEX-4T1 (Pharmacia Biotechnology, Piscataway, NJ) vector. This construct expresses the last 22 amino acids of the rod domain and the complete G3 domain of NID-1 fused to the carboxyl end of GST. The fusion protein was SDS-PAGE purified and used to immunize rabbits. An MBP-NID-1(G3 domain) fusion protein was generated by inserting the 1.1-kb PvuII-XhoI fragment (nucleotides 8239-9521) from the yk38f9 cDNA into the pMAL-c2 vector (New England Biolabs, Beverly, MA). The purified pMAL fusion protein was used to affinity purify the rabbit antisera as previously described (Graham et al., 1997).

Western Blot Analyses

Embryos or mixed stage animals were quick-frozen in liquid nitrogen, pulverized while frozen, and extracted with 6 M guanidine-HCl, 50 mM Tris-HCl (pH 7.4), containing 10 mM EDTA, 2 mM N-ethylmaleimide (NEM), and 2 mM PMSF (Dziadek and Timpl, 1985). Guanidine extracts were dialyzed against 7 M urea, 50 mM Tris-HCl (pH 8.6) with 10 mM EDTA, 5 mM NEM, and 5 mM PMSF at 4°C. The resulting supernatants were acetone precipitated and resuspended in dH₂O. Samples were subjected to 7% SDS-PAGE and transferred to nitrocellulose filters. Filters were reacted with affinity purified anti-NID-1 antiserum diluted 1:500 and detected using the ECL system (Amersham Pharmacia Biotechnology).

Immunocytochemistry

Embryos were purified by alkaline hypochlorite treatment and fixed with 4% paraformaldehyde as described previously (Graham et al., 1997). Larval and adult animals were prepared as described (Finney and Ruvkun, 1990). After fixation, samples were rehydrated and blocked for 1 h in PBS containing 0.1% Triton X-100 (PBS-T) and 10% normal donkey serum (NDS). Samples were then incubated overnight at room temperature with various primary antibodies diluted with PBS-T-NDS: anti-NID-1 (1:50), anti-LET-2 type IV collagen (1:100) (Graham et al., 1997), monoclonal anti-MHC-A myosin (1:1000), MH35 anti--actinin (1:500) (Francis and Waterston, 1985). Samples were washed three times with PBS-T, incubated for 1 h at room temperature with FITC-labeled donkey anti-rabbit and Texas red-labeled donkey anti-mouse secondary antibodies (Jackson ImmunoResearch, West Grove, PA) in PBS-T-NDS, and washed with PBS-T and PBS. All samples were mounted with 90% glycerol in PBS (pH 8.0) containing 1 mg/ml para-phenylenediamine and 1% DABCO. Images were captured on ASA 400 color slide film or digitally recorded with a digital camera. Digital images were deconvolved (VayTek, Inc., Fairfield, IA) to remove out-of-focus information, and contrast and brightness were adjusted using Adobe Photoshop (San Jose, CA).

nid-1 Expression Analyses

RNA was extracted from C. elegans samples using TRIZOL reagent (Life Technologies-BRL, Grand Island, NY). Poly-A-enriched RNA was isolated using batch oligo(dT)-cellulose chromatography (Sambrook et al., 1989). Six-microgram samples of poly-A-enriched RNA were fractioned on 0.8% formaldehyde-agarose gels and transferred to nylon membranes. The membranes were hybridized to ³²P-labeled nid-1 exon 11-, exon 13-, or exon 15-specific probes in modified Church buffer (50% formamide, 250 mM NaCl, 250 mM Na₂PO₄, 6% SDS, and 6% polyethylene glycol) and washed with 2× SSC, 0.1% SDS at 37°C for 45 min and with 0.2× SSC, 0.1% SDS at 60°C for 30 min.

nid-1 Phenotypic Analyses

Organization of epidermal cells was examined in jcIs1; cg118 and jcIs1; cg119 animals. jcIs1 expresses the JAM-1::GFP marker, which labels adherens junctions between cells (Mohler et al., 1998). The presence of cg118 or cg119 were confirmed by PCR analysis for the corresponding deletion. The morphologies of excretory cells and gonads were determined by observations of live nid-1 mutant animals using Nomarski DIC optics.

To determine fecundity, individual animals were picked at the L4 larval stage and were transferred daily to new plates until they stopped laying fertilized eggs. The total numbers of eggs laid each day was counted using a dissecting microscope. For the reported analysis animals were maintained at 20 ± 0.5°C, but similar results were also obtained at 24°C. To count spermatozoa, young adult animals were fixed with 4% paraformaldehyde in PBS for 4 h at 4°C, washed in PBS, postfixed with 0°C methanol for 5 min, and incubated with 100 mM DAPI for 1 h. A z-dimension stack of digital fluorescence microscopic images through the spermatheca was generated, and sperm nuclei were counted. Actin filaments were visualized using a modified rhodamine phalloidin staining procedure (Strome, 1986; Rose et al., 1997). Adult hermaphrodite gonads were extruded by decapitation in culture medium (80 mM NaCl, 20 mM KCl, 10 mM MgCl₂, 5 mM HEPES, pH 7.2, 10% FBS, 0.01% tetramisole), fixed with 3% paraformaldehyde in PBS containing 0.1% Tween-20 (PBS-TW) for 2 h at room temperature, washed in PBS-TW, and incubated with 0.33 µM rhodamine phalloidin (Molecular Probes, Inc., Eugene, OR) in PBS-TW.

	RESULTS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

The C. elegans nid-1 Nidogen Gene

A single nidogen gene nid-1 was identified in the C. elegans genome in cosmid F54F3 based on sequences generated by the genome sequencing project (Waterston and Sulston, 1995). nid-1 encodes domains similar to vertebrate nidogen G1, G2, and G3 globular domains and a rod domain of EGF repeats (Figures 1A and 2; Table 1). The region with highest amino acid sequence identity to mammalian nidogen is the G3 domain, particularly the six YWTD repeats of this domain (Springer, 1998). There is also strong similarity in the G2 and G1 domains (Figure 2; Table 1). On the basis of sequence similarity, NID-1 is not significantly more similar to mammalian nidogen-1 than to nidogen-2. However, both NID-1 and nidogen-1 have a carboxyl terminal EGF repeat that is not present in nidogen-2. The major difference between the C. elegans and mammalian nidogens is that nid-1 encodes 12 EGF repeats in the rod domain, whereas vertebrate nidogen-1 and nidogen-2 have four EGF and one or two thyroglobulin repeats in this region. The strongest sequence conservation has been maintained in the G3 and G2 domains, which mediate binding to laminin and type IV collagen, respectively.

View larger version (22K): [in this window] [in a new window]   Figure 1.   Structure of the C. elegans nid-1 nidogen gene. (A) Exons are indicated as boxes with numbers below. The nid-1A-specific exon 11 is colored dark gray, and the nid-1A- and B-specific exons 12-14 are colored light gray. The extents of the G1, G2, and G3 globular domains and the rod domain are indicated at the top of the figure. The extents of the cg118 and cg119 deletions are shown below the figure. (B) The structures of the three alternatively spliced transcripts generated from the nid-1 gene.

View larger version (85K): [in this window] [in a new window]   Figure 2.   Comparisons of nidogen protein structures and sequences. (A) Domain structures of C. elegans NID-1, mouse nidogen-1 (Mann et al., 1989), human nidogen-2 (Kohfeldt et al., 1998), and Ascidian nidogen (Nakae et al., 1993). Domains are represented as follows: , G1; , G2; , EGF; , thyroglobulin; , YWTD motif (Springer, 1998). EGF repeats that are present in all forms are colored black, the NID-1A-specific EGF is dark gray, and the NID-1A- and B-specific EGFs are light gray. The extents of domains for NID-1A are indicated at the top. (B) Sequence alignment of the G1 through G2 domains of NID-1, mouse nidogen-1, and human nidogen-2. Identical residues are shown as white letters on black background; similar residues are black letters on gray background. Levels of sequence identity and similarity are presented in Table 1. (C) Sequence alignment of the G3 domains of NID-1, mouse nidogen-1, and human nidogen-2. Residues are highlighted as described for B.

The predicted exon structure of nid-1 suggested that some of the rod domain EGF repeats could be removed by alternative splicing. We analyzed RT-PCR products and cDNA clones derived from nid-1 to demonstrate that three isoforms (nid-1A, B, C) are generated by alternative splicing (Figures 1 and 2A). All three isoforms contain the amino G1 and G2, and carboxyl G3 globular domains, but they differ in the number of EGF repeats in the rod domain. The longest form, NID-1A, contains all 12 EGF repeats in the rod domain, NID-1B is missing the single EGF repeat encoded by exon 11, and NID-1C lacks 7 EGF repeats encoded by exons 11-14. The NID-1C rod domain, containing five EGF repeats, is most similar in length to mammalian nidogen-1, which has four EGF and one thyroglobulin repeat (Figure 2A).

There is a single RGD (700-702) sequence in mouse nidogen-1, located near the end of the first rod domain EGF repeat. This RGD sequence has been shown to provide a major nidogen cell attachment activity in vitro (Mann et al., 1989; Dong et al., 1995). There is also a single RGD (714-716) present in NID-1, and it is located in the same region of the molecule, at the start of the second rod domain EGF repeat. This RGD sequence is present in all three NID-1 isoforms. There are no RGD sequences in nidogen-2 or the ascidian nidogen.

Expression of nid-1

The nid-1 isoforms are differentially expressed during development. RNA isolated from embryos contains an abundant transcript of 5.1 kb (Figure 3A). This transcript is approximately the size expected for the nid-1A splice isoform, which is predicted to be 4.8 kb before polyadenylation. The 5.1-kb transcript hybridizes to the form A-specific exon 11, demonstrating that it is the nid-1A transcript. No other transcripts are detected when exon 13 or exon 15, which would hybridize to the other isoforms, are used to probe embryo RNA. Transcripts of 5.1 and 4.2 kb are detected in approximately equal abundance in RNA isolated from mixed populations containing animals of all developmental stages (Figure 3A). The 4.2-kb transcript hybridizes to exon 15 but not to exons 11 or 13, indicating that it represents the nid-1C isoform (3.8 kb predicted size before polyadenylation).

View larger version (29K): [in this window] [in a new window]   Figure 3.   Northern and Western blot analyses of nid-1 expression in wild-type and mutant animals. (A) Detection of nid-1 transcripts in RNA extracted from wild-type (N2) and nid-1 mutant (cg118, cg119) animals. RNAs were extracted from embryos (Embryo) or populations of animals of mixed development stages (Mixed). The positions of nid-1A (open arrow) and nid-1C (filled arrow) transcripts are indicated between A and B. This blot was probed with radiolabeled nid-1 exon 15 to reveal all isoforms. Equivalent blots were also probed with exons 11 and 13. Hybridization with the RNA polymerase II gene (ama-1) was used as a loading control. The positions of size standards are indicated on the left. (B) Detection of NID-1 in extracts from wild-type (N2) and nid-1 mutant (cg118, cg119) animals. Extracts were made from embryos (Embryo) or populations of animals of mixed development stages (Mixed). The positions of putative NID-1A (open arrows), NID-1B (shaded arrows), and NID-C (filled arrows) are indicated to the right of each lane. The positions of molecular weight standards are indicated on the left.

Probing with exon 13, which would hybridize to nid-1A and nid-1B, did not reveal any transcripts beyond those detected when exon 11 was used as a probe to visualize the nid-1A transcripts. The predicted size of the nid-1B transcript (4.6 kb) is only 4% smaller than nid-1A, so it may be present in low abundance but masked by the nid-1A band. These experiments show that nid-1A is the major transcript expressed during embryogenesis and that detectable nid-1C expression does not begin until after the completion of embryogenesis.

Specific NID-1 antiserum was raised against a bacterially expressed fusion protein containing the carboxyl-terminal G3 domain that is common to all NID-1 isoforms. Western blots of embryonic C. elegans extracts show a strong immunoreactive band migrating at ~170 kDa and a weak band at 160 kDa (Figure 3B). Extracts of mixed stage animals show the 170- and 160-kDa bands as well as a strong band at 150 kDa. The 170- and 150-kDa bands are likely to be NID-1A and C isoforms, because they are similar to the predicted masses of 172 and 136 kDa, respectively, and they exhibit the same temporal characteristics as the nid-1A and nid-1C transcripts (Figure 3A). The 160-kDa band may be NID-1B, because it is close to the 166-kDa predicted mass for NID-1B. Two or more lower-molecular-weight bands are variably seen in mixed stage extracts and may represent degradation products of NID-1. These results are consistent with the RNA expression data in showing that NID-1A is the major embryonic form and NID-1C is not strongly expressed until after embryogenesis.

NID-1 Tissue Localization

Anti-NID-1 antiserum that reacts with all isoforms was used to localize the protein in whole animals. NID-1 was first detected in embryos at the beginning of morphogenesis (lima stage) localized to body wall muscle cells. As embryos elongate strong NID-1 staining is seen around body wall muscle cells and diffuse stain begins to accumulate on the surfaces of the pharyngeal and intestinal primordia (Figure 4,A and B). Once the embryo has elongated to the twofold stage, NID-1 has localized to the basal face of the body wall muscles and shows strong accumulation on the surfaces of the pharyngeal, intestinal, and gonad primordia (Figure 4, C and D). In three- and fourfold stage embryos, NID-1 accumulates to higher levels and remains localized under the four body wall muscle quadrants and on the surfaces of the pharynx, intestine, and gonad (Figure 4, E and F). In L1 larvae the intensity of staining of body wall muscle, pharynx, and intestine appears reduced, and strong staining associated with the nerve ring becomes apparent (Figure 4G). This pattern continues through the L2 and L3 larval stages, with the addition of stronger staining of the distal tip cells as they lead the growth of the gonad (Figure 4I). In late L3 to L4 stage larvae particularly strong NID-1 accumulation is seen associated with the distal tip cells and the developing somatic structures of the gonad, the spermatheca, uterus, and vulva (Figure 4, H and J).

View larger version (64K): [in this window] [in a new window]   Figure 4.   Localization of NID-1 in wild-type animals. Animals were stained with anti-NID-1 (A, C, E, and G-K) and anti-myosin A (B, D, F, and L). (A and B) A comma stage embryo shows strong accumulation of NID-1 around body wall muscle cells (arrowhead) and diffuse association with the pharyngeal and intestinal primordia. (C and D) A twofold stage embryo shows NID-1 accumulation on the basal face of body wall muscles (arrowhead) and on the surfaces of the developing pharynx (p), intestine (i), and gonad (g). (E and F) A threefold stage embryo showing NID-1 localized under the four body wall muscle quadrants (arrowheads) and on the surfaces of the pharynx (p) and intestine (i). A body wall muscle quadrant over the intestine is indicated with an arrow. (G) In an L1 larva, strong NID-1 accumulation is seen around the nerve ring (nr). Staining is also seen at the basal face of body wall muscles (arrowheads) and on the surfaces of the pharynx (p), intestine (i), and gonad (g). (H) In an L4 larva, there is strong NID-1 accumulation on the developing spermathecae (st), vulva (vu), and uterus, and at the distal tip cells (dt). There is also weaker staining on the surfaces of the pharynx and intestine, at the nerve ring (nr), and under body wall muscles. (I) In a late L2-early L3 larva, NID-1 accumulates around the distal tip cell (dt) that is leading growth of the gonad (g). (J) An L4 larva showing strong NID-1 accumulation at the distal tip cell (dt), spermatheca (st), and uterus (u). (K and L) In an adult animal, NID-1 is distributed in a weak punctate pattern under the muscle cell dense bodies and is also present between the muscle quadrants. There is stronger NID-1 accumulation at the edges of the muscle quadrants (arrows) and weak accumulation between adjacent muscle cells (arrowheads). vu, vulva.

Under the body wall muscles of larval and adult stage animals NID-1 is organized as punctate lines (Figure 4, K and L). These lines follow the rows of dense bodies within the muscle cells (Figure 5,A-C). NID-1 also accumulates strongly at the outer edges of the muscle quadrants and more weakly at the boundaries between muscle cells within each quadrant. Less organized NID-1 staining is seen in the regions between the body wall muscle quadrants, presumably associated with the epidermal basement membranes in these regions.

View larger version (81K): [in this window] [in a new window]   Figure 5.   NID-1 localization to sublateral nerves and muscle. Wild-type N2 (A-E) or nid-1(cg118) (F and G) animals were stained with anti-NID-1 (A, C-G) or anti--actinin (B and C) antibodies. (A-C) Part of a body wall muscle cell of an adult animal costained with anti-NID-1 and anti--actinin antibodies. (A) NID-1 accumulates beneath the dense body lines and along the interface between muscle cells (arrowheads). (B) The same region showing anti--actinin staining to visualize dense bodies. There is no -actinin at the muscle cell interface. (C) The merged image of A and B shows that NID-1 localizes along a line that follows the dense bodies. (D and F) Dorsal views of L2 larvae; anterior is to the left. NID-1 is seen to accumulate on the left and right dorsal sublateral nerves (arrows), the dorsal edges of the left and right dorsal muscle quadrants (dm), and the lateral edges of the dorsal muscle quadrants (arrowheads). (E and G) Left lateral views of L4 larvae showing NID-1 accumulation on the dorsal and ventral sublateral nerves (arrows) and along the lateral edges of the dorsal and ventral body wall muscle quadrants (arrowheads). vu, vulvae.

NID-1 accumulates along the four sublateral nerves that run beneath the center of each muscle quadrant (Figure 5, D and E). The sublateral nerves extend along dorso- and ventrolateral tracts from the nerve ring in the anterior of the animal to near the middle of the animal where they turn further lateral to positions coincident with the lateral edges of the body wall muscle quadrants. NID-1 accumulation on these nerves is seen in larval and adult animals and is similar in intensity to the staining at the edges of the body wall muscle quadrants. Staining along the edges of the body wall muscle quadrants appears to be associated with the muscle edges rather than the nerves in these regions because the staining closely follows the edge of the muscles and it does not display the left/right asymmetry expected for the ventral and dorsal nerve cords. As noted above, less organized NID-1 staining is also present in the regions between body wall muscle quadrants.

nid-1 Mutants

A Tc1 transposon insertion was identified near the start of exon 2 in the nid-1 gene, between the codons for Tyr54 and Met55 (nucleotides 3041-3042). Animals homozygous for the insertion are viable and fertile and display no obvious abnormal phenotypes. The Tc1 element could be removed by RNA splicing and not severely affect nid-1 function. We therefore identified two deletions caused by imprecise excisions of the Tc1 element. One deletion, cg118, removes 2582 nucleotides (3036-5617: GenBank Z79696), resulting in an in-frame fusion of the beginning of exon 2 with part of exon 8 (Figure 1A). This deletion would result in loss of all of the G2 globular domain, part of the G1 domain, and 33 amino acids from the first EGF repeat in the rod domain. The second deletion, cg119, removes 3133 nucleotides (1499-4631: GenBank Z79696), extending from 952 nucleotides before the nid-1 initiator ATG through exon 7 (Figure 1A). After outcrossing to remove extraneous mutations, homozygous cg118 and cg119 animals showed no overt abnormal phenotypes.

The effects of both deletions on nid-1 expression were examined at both the RNA and protein levels. cg118 animals contain nid-1 transcripts of ~3.2 and 2.2 kb, ~1.8 kb shorter than in wild type (N2), as predicted by the extent of the deletion (Figure 3A). The truncated nid-1A and C transcripts are present at approximately wild-type levels. No nid-1 transcripts are detectable in RNA extracted from cg119 animals, although control ama-1 RNA is detected at normal levels in these same samples. Western blots of cg118 extracts reacted with anti-NID-1 antiserum show strong bands of ~113- and 72-kDa apparent masses and a weak band of 99 kDa (Figure 3B). These sizes are similar to the 103-, 67-, and 97-kDa masses predicted for the cg118 truncated NID-1A, C, and B products, respectively. The truncated NID-1A and NID-1C proteins are present at approximately wild-type levels. No NID-1 protein is detectable on Western blots of extracts from cg119 animals reacted with anti-NID-1 antiserum (Figure 3B), and NID-1 is not detectable in these animals by immunofluorescence (Figure 6,E and F). These results show that the cg119 deletion allele is a molecular null, whereas the cg118 deletion mutant produces approximately wild-type levels of truncated NID-1 that is missing all of the G2 and part of the G1 domains.

View larger version (76K): [in this window] [in a new window]   Figure 6.   NID-1 localization in cg118 and cg119 mutant animals. Animals were stained with anti-NID-1 antiserum (A, C, E, G, H, and I) or anti-myosin A (B, D, F, and J). (A and B) Twofold stage cg118 mutant embryo shows NID-1 localization under body wall muscle (arrowheads) and on the surfaces of the pharynx (p), intestine (i), and gonad (g). (C and D) Threefold stage cg118 mutant embryo showing NID-1 accumulation at the basal surfaces (arrowheads) and edges (arrows) of body wall muscle quadrants and on the surfaces of the pharynx (p) and intestine (i). (E and F) Twofold stage cg119 mutant embryo showing complete lack of staining by anti-NID-1 antibodies. (G) L1 stage cg118 mutant larva showing NID-1 accumulation at nerve ring (nr), under body wall muscle (arrow), and on the surfaces of the pharynx (p), intestine (i), and gonad (g). (H) Late L4 stage cg118 mutant larva showing NID-1 accumulation on the developing spermatheca (st), uterus (u), and vulva (vu), and at the distal tip cell (dt). Weaker accumulation on the surface of the gonad is also seen (arrow). (I and J) Ventral view of cg118 mutant adult hermaphrodite showing NID-1 accumulation at the ventral edges of the ventral body wall muscle quadrants (arrows) and at the edges between adjacent muscle cells (arrowheads).

Effect of nid-1(cg118) Deletion on NID-1 Localization

The cg118 deletion completely removes the G2 domain, which for mouse nidogen-1 has been shown to interact with type IV collagen and perlecan (Reinhardt et al., 1993; Kohfeldt et al., 1998). To determine whether loss of the G2 domain affects the in vivo distribution of nidogen, we performed immunohistochemistry of cg118 mutant animals with anti-NID-1 antisera (Figure 6). Examination of cg118 embryos, larvae, and adults revealed no detectable differences from wild-type in the timing of NID-1 appearance, tissue localization, or levels. The cg118 truncated NID-1 accumulates on the edges of muscle quadrants and on the sublateral nerves in a manner indistinguishable from wild-type NID-1 (Figure 5, F and G). The stability and normal tissue distribution of nidogen are therefore not dependent on the presence of its G2 domain.

Effects of nid-1 Mutations on Type IV Collagen Localization

The ability of mouse nidogen-1 to form a ternary complex between type IV collagen and laminin suggests that it may influence type IV collagen assembly. The wild-type distribution of type IV collagen in C. elegans was previously described (Graham et al., 1997). We examined the distribution of type IV collagen in nid-1 deletion mutant animals using antitype IV collagen antisera (Figure 7). No difference from the wild-type localization pattern was seen in embryos, larvae, or adult homozygotes for either nid-1 deletion. The fine pattern of collagen IV localization under body wall muscles is not altered in the nid-1 mutants. We conclude that NID-1 is not required for normal localization of type IV collagen in C. elegans.

View larger version (93K): [in this window] [in a new window]   Figure 7.   Type IV collagen localization in wild-type and nid-1 mutant animals. Wild-type N2 (A-D, K, L, N, P, and Q), cg118 mutant (E-H, M, O, R, and S) or cg119 mutant (I, J, T, and U) animals were stained with anti-LET-2 type IV collagen antisera (A, C, E, G, I, K, M-P, R, T) or anti-myosin A antibodies (B, D, F, H, J, L, Q, S, and U). (A and B) Wild-type twofold stage embryo showing type IV collagen localization under body wall muscles (arrow) and on the surfaces of the pharynx (p) and intestine (i). (C, D). Wild-type threefold stage embryo showing type IV collagen localization under body wall muscle quadrants (arrowheads), and on the surface of pharynx (p) and intestine (i). (E, F). Twofold stage cg118 mutant embryo showing type IV collagen localization to body wall muscles (arrow), and on the surfaces of the pharynx (p) and intestine (i). (G and H) Threefold stage cg118 mutant embryo showing type IV collagen localization under (arrowheads) and at the edges (arrow) of body wall muscle quadrants and on the surfaces of the pharynx (p) and intestine (i). (I and J) Threefold stage cg119 embryo showing type IV collagen localization under (arrowheads) and at the edges (arrow) of body wall muscle quadrants and on the surfaces of the pharynx (p) and intestine (i). (K and L) Wild-type L1 larva showing type IV collagen localization under body wall muscle (arrow), at the nerve ring (nr), and on the surfaces of the pharynx (p), intestine (i), and gonad (g). (M) L1 stage cg118 mutant larva shows type IV collagen localization under body wall muscle (arrow), at the nerve ring (nr), and on the surfaces of the pharynx (p), intestine (i), and gonad (g). (N) Wild-type L4 stage larva showing type IV collagen localization to the surface of the gonad (arrow), the distal tip cell (dt), spermatheca (st), and uterus (u). The location of the vulva (vu) is indicated. (O) L4 stage cg119 mutant larva shows type IV collagen localization to the surface of the gonad (arrow), the distal tip cell (dt), spermatheca (st), and uterus (u). vu, vulva. (P and Q) Wild-type L4 stage larva showing type IV collagen accumulation at the interfaces between muscle cells (arrows) and in a dense body pattern under the cells. (R and S) L4 stage cg118 mutant animal showing type IV collagen accumulation at the interfaces between muscle cells (arrows) and in a dense body pattern under the cells. (T and U) L4 stage cg119 mutant animal showing type IV collagen accumulation at the interfaces between muscle cells (arrows) and in a dense body pattern under the cells.

Effects of nid-1 Mutations on Epithelial Functions

Defects in epithelial morphogenesis and function have been noted when nidogen function has been perturbed in vertebrate cultures of cells or tissues (Ekblom et al., 1994; Kadoya et al., 1997). We examined analogous tissues of C. elegans in cg118 and cg119 mutant animals. No alterations of the epidermis (hypodermis) were seen using jam-1::GFP as a marker to visualize cell-cell junctions. The excretory cell of C. elegans is analogous to the vertebrate kidney and extends long tubular processes between the epidermis and the overlying basement membrane (Nelson et al., 1983). These processes appear normal in nid-1 mutant animals. The gonad is a basement membrane covered tube that forms stereotypical anterior and posterior reflexed arms by directed migrations during development (Kimble and Hirsh, 1979). The gonad migrations occur normally in both nid-1 mutants. We conclude that nidogen is not required for the proper migrations and organization of these tissues.

Reduced Fecundity of nid-1 Mutants

Although cg118 and cg119 mutant strains display no obvious defects in morphology or motility, they do have somewhat reduced fecundity relative to the wild-type N2 strain. On average, cg119 hermaphrodites have 32% fewer offspring than N2, and cg118 hermaphrodites have 11% fewer offspring (Table 2). The reduced fecundity of the nid-1 mutants is not due to reduced numbers of sperm because they contain the same number of sperm in their spermathecae as the N2 strain (Table 2). They do not appear to have an egg-laying defect because they carry the same number of fertilized eggs in the uterus as wild-type animals. There are also no apparent defects in the structure of the spermatheca or uterus observable by Nomarski optics, and rhodamine-phalloidin staining reveals no obvious defects in the actin filaments of the uterus, spermatheca, or sheath cells (data not shown). Although the cause is unclear, the reduced fecundity of nid-1 mutants would be a substantial selective detriment in the animal's natural environment.

	DISCUSSION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

Nidogen appears as a major component of basement membranes in all organisms that have been characterized. The ability of nidogen to bind with high affinity to both type IV collagen and laminin suggested that it might be a critical molecule for assembly of basement membranes. In vitro studies in vertebrate systems have supported the potential importance of nidogen for cell adhesion and tissue morphogenesis. We have shown that loss of nidogen in C. elegans results in viable animals that are fertile and that display no overt abnormal phenotypes. Loss of other C. elegans basement membrane components, such as type IV collagen (Guo et al., 1991; Sibley et al., 1993; Gupta et al., 1997), perlecan (Rogalski et al., 1993), or SPARC (Fitzgerald and Schwarzbauer, 1998), results in embryonic lethality. If nidogen were critical for the assembly or function of these constituents of basement membranes, then its absence would also be expected to result in lethality. The fact that this is not the case provides strong evidence that nidogen is not required for these molecules to assemble into functional basement membranes.

nid-1 is the only gene in C. elegans that encodes a classical nidogen with G1, G2, rod, and G3 domains. There are, however, other genes that encode nidogen-related domains. Three predicted gene products (B0393.5, D1044.2, K03H1.5) contain G1-like domains. All three also encode predicted transmembrane domains near their carboxyl-termini, suggesting that they are cell surface transmembrane proteins. Six distinct predicted gene products contain YWTD motifs related to the nidogen G3 domain: five are predicted to be transmembrane, one is predicted to be secreted. All six also contain EGF repeats but no other similarities to nidogen. There are no other genes that encode nidogen G2-related domains. It is possible that the nidogen G1- and/or G3-related domains of these other gene products could replace NID-1 functions in nid-1 mutant animals.

Our immunolocalization studies show that NID-1 accumulates at all sites where type IV collagen was previously shown to assemble (Graham et al., 1997) but is additionally found at sites where little or no collagen IV is detected. In particular, NID-1 shows strong accumulation around the nerve ring, whereas collagen IV is only weakly detected in this region. NID-1 accumulates on the sublateral nerves, where collagen IV shows no preferential accumulation. Both collagen IV and nidogen show strong accumulation under body wall muscles during embryonic development, but the level of nidogen decreases during larval and adult stages, whereas collagen IV levels are maintained. Nidogen accumulates strongly at the edges of the muscle quadrants and only weakly at the junctions between adjacent muscle cells, whereas collagen IV shows the opposite pattern, being strongest at muscle cell junctions. NID-1 also shows diffuse accumulation on the regions of the epidermal basement membrane that lie between the body wall muscle quadrants, whereas collagen IV is absent from these regions. Thus, although collagen IV and nidogen are generally coincident, there are sites where substantial differences in their relative abundance are seen. These results indicate that there is not a one-to-one correspondence between collagen IV and nidogen in C. elegans basement membranes.

Type IV collagen assembles into all basement membranes of C. elegans except on the regions of the epidermis that are located between the body wall muscle quadrants and on the pseudocoelomic faces of the muscle quadrants (Graham et al., 1997). We had expected that nidogen would be involved in localization of collagen IV assembly, based on its ability to link collagen IV and laminin. However, we showed that type IV collagen assembles in a completely normal manner in the absence of nidogen. How is assembly of type IV collagen restricted to particular locations? Type IV collagen could be localized to cell surfaces by binding to integrins. However, in mutants for two C. elegans integrins, ina-1 -integrin (Baum and Garriga, 1997) and pat-3 -integrin (Gettner et al., 1995), type IV collagen localization appears normal (J. Kramer, unpublished results). It is possible that collagen IV assembly is dependent on other integrins or some other unidentified molecule. It is also possible that there are redundant mechanisms for collagen IV localization, such that loss of any one binding molecule will not reveal abnormalities in collagen assembly.

The mammalian nidogen-1 G2 domain binds type IV collagen and perlecan with high affinity (Reinhardt et al., 1993). The C. elegans and mammalian G2 domains show strong sequence conservation, suggesting that the C. elegans G2 domain is likely to also be capable of binding collagen IV and perlecan. The nid-1(cg118) deletion completely removes the nidogen G2 domain as well as parts of G1 and the first EGF repeat of the rod domain. The cg118-deleted NID-1 accumulates to approximately normal levels and shows normal tissue localization. Thus, the G2 domain has little or no role in nidogen stability or assembly into basement membranes. Nidogen localization therefore appears to occur independently of collagen IV and perlecan interactions.

If linking between collagen IV and laminin were a critical aspect of nidogen function, then the cg118 G2 domain deletion might be expected to act as a dominant interfering mutant. The truncated NID-1 could bind to laminin but would be unable to bind type IV collagen. Mammalian nidogen-1 was found to cooperate with laminin-1 to stimulate -casein synthesis by cultured mammary epithelial cells (Pujuguet et al., 2000). However, addition of the nidogen-1 G3 domain inhibited laminin-1 stimulation of -casein synthesis, suggesting that the presence of the laminin-binding G3 domain in the absence of other parts of nidogen dominantly interferes with laminin signaling. In coculture studies, antisense inhibition of laminin expression blocked the accumulation of type IV collagen and nidogen at the epithelial/mesenchymal interface (De Arcangelis et al., 1996), leading to the suggestion that linkage of collagen IV to laminin via nidogen was essential for its assembly. The fact that the cg118 G2 deletion nidogen does not cause dominant phenotypes indicates that its binding to laminin does not interfere with any essential processes. If nidogen was required to link type IV collagen to laminin, then the cg118 deletion NID-1 should dominantly interfere with this process.

We have identified three alternative splice isoforms of NID-1 that contain differing numbers of EGF repeats in their rod domains. NID-1A is the longest and is the predominant embryonic form, whereas the shortest, NID-1C, only appears after the completion of embryogenesis. The different numbers of EGF repeats would change the spacing between the collagen IV and laminin-binding domains of the NID-1 isoforms and could also change their binding repertoires. Alternative splice forms of other nidogen genes have not been reported. However, mammals have two nidogen genes that may provide additional diversity of nidogen function.

The alternative splice isoforms of nid-1 are reminiscent of let-2, the 2(IV) collagen gene of C. elegans. let-2 generates two alternative splice isoforms that dramatically change relative abundance immediately after the completion of embryogenesis (Sibley et al., 1993). The C. elegans perlecan gene unc-52 also produces alternatively spliced variants that differ in their expression during development (Mullen et al., 1999). The basement membranes of C. elegans appear to undergo significant changes in composition on completion of embryogenesis.

Perturbation of nidogen-1 function in cultured mammalian kidney, lung, or submandibular gland (Ekblom et al., 1994; Kadoya et al., 1997) and correlation of nidogen proteolysis with mammary gland involution (Alexander et al., 1996) have suggested that nidogen has important roles in epithelial function. We particularly looked in C. elegans nid-1 mutants at tissues that would reveal defects in epithelial morphogenesis. In nid-1 mutants no defects were seen in the excretory canals, which are analogous to the vertebrate kidney, the gonad, which forms by directed migration of a basement membrane enclosed tube, or in organization of the hypodermal cells, which form the epidermis of nematodes. We conclude that nidogen is not required for morphogenesis of these epithelia in C. elegans. It is possible that such functions for nidogen have arisen in vertebrates or that the in vitro studies are misleading as to the in vivo functions of nidogen.

We showed that nid-1 mutations cause reduced fecundity. The nid-1 null mutant showed a greater reduction (32%) than the G2 domain deletion mutant (11%). Thus, NID-1 lacking the G2 domain is able to provide some, but not all, of the nidogen function required for normal fecundity. The cause of the reduced fecundity of nid-1 mutants has not been elucidated. However, NID-1 shows strong accumulation around the developing spermatheca, uterus, and vulva. Although no defects were detected in these structures, it is possible that they do not function optimally in the absence of nidogen. The reduced fecundity that results from either of these nid-1 mutations would result in sufficient selective pressure to ensure maintenance of the gene, even in the absence of any other functions.

During preparation of this article a report appeared showing that a nid-1 mutation, ur41, can affect positioning of longitudinal nerves in C. elegans (Kim and Wadsworth, 2000). The nid-1(ur41) mutation results in no overt morphological or behavioral phenotypes, in agreement with our findings. The nerve positioning defects are only detectable using GFP markers to visualize the affected axons. The most penetrant defect reported was mispositioning of the dorsal sublateral nerves up toward the dorsal midline. Our demonstration that NID-1 accumulates on the sublateral nerves indicates that it directly associates with the affected axons. The strong accumulation of NID-1 at the edges of the body wall muscle quadrants that we demonstrated is consistent with the proposal that nidogen may influence the guidance decisions of dorsoventrally migrating axons at the interfaces between muscle and epidermis (Kim and Wadsworth, 2000).