Biol Reprod Email Content Delivery
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow My Folders
Right arrow Download to citation manager
Right arrow reprints & permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Cerdà, J.
Right arrow Articles by Franke, W. W.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Cerdà, J.
Right arrow Articles by Franke, W. W.
Agricola
Right arrow Articles by Cerdà, J.
Right arrow Articles by Franke, W. W.
Biology of Reproduction 61, 692-704 (1999)
©Copyright 1999 Society for the Study of Reproduction, Inc.

Articles

Cadherin-Catenin Complexes During Zebrafish Oogenesis: Heterotypic Junctions Between Oocytes and Follicle Cells1

Joan Cerdà2,a, Sonja Reidenbacha, Silke Prätzela, and Werner W. Frankea

a Division of Cell Biology, German Cancer Research Center, D-69120 Heidelberg, Germany


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
During vertebrate oogenesis, the germ cells and associated somatic cells remain connected by a variety of adhering junctional complexes. However, the molecular composition of these cellular structures is largely unknown. To identify the proteins forming the heterotypic adherens junctions between oocytes and follicle cells in the zebrafish (Danio rerio), the cDNAs encoding {alpha}E-catenin and plakoglobin were isolated. Using these cDNAs, in combination with the previously isolated ß-catenin cDNA, and antibodies specific for {alpha}- and ß-catenin, plakoglobin, and N- and E-cadherin, we found differences in catenin and plakoglobin gene expression during oogenesis. The immunolocalization of these plaque proteins, as well as of cadherins, in the ovarian follicle indicated an enrichment of {alpha}- and ß-catenin and of E-cadherin-like protein(s) in the oocyte cortex, notably at sites of oocyte-follicle cell contacts, suggesting the presence of hitherto unknown heterotypic adherens junctions between these cells. By contrast, plakoglobin and N-cadherin localization was restricted to cell-cell contacts in the follicle cell layer. During oocyte maturation, mRNAs for {alpha}E- and ß-catenin and plakoglobin accumulated, and all three plaque-forming proteins were stored in unfertilized eggs, either in complexed forms with cadherins or as free cytoplasmic pools. These findings suggest possible roles of these junctional proteins during early embryogenesis.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Cell-cell adhesion within a tissue is mediated by different cell junctions among which the adherens junctions and desmosomes constitute prominent structures forming semistable intercellular contacts (for details and references, see [1]). Characteristically, all these adhering junctions are composed of transmembrane glycoproteins, members of the cadherin family, and cytoplasmic plaque proteins that anchor the cytoskeleton to the cadherin. However, whereas the adherens junctions are formed by "classical" cadherins (e.g., E-, N-, and P-cadherin) that associate with specific plaque proteins, such as {alpha}- and ß-catenins and plakoglobin, vinculin, {alpha}-actinin, and in some cases also protein p120cas (reviewed in [1, 2]), the desmosomes are clusters of desmosomal cadherins (desmogleins and desmocollins), which in the cytoplasm assemble (in addition to the common plaque protein plakoglobin) several desmosome-specific proteins, including desmoplakins I and II and plakophilins [1, 3–9].

Important roles of these junctions and their constitutive molecules have recently been demonstrated during embryogenesis and in tissue morphogenesis [10–17]. However, the nature and function of cell adhesion structures in the vertebrate ovarian follicle, particularly those connecting the oocyte with the somatic (follicle or granulosa) cells, are still unclear. Besides the existence of gap junctions, electron microscopy studies of the oocyte-follicle complexes of various vertebrates have documented the presence of a variety of junctional structures, tentatively defined as adherens or "desmosome-like" junctions [18–24]. The occurrence of functional gap junctions between oocytes and the surrounding follicle cells has been well established by fluorescent dye transfer and metabolic coupling assays (e.g., [25–29]), and conclusive evidence of the importance of the signaling through these channels for successful oogenesis and ovulation has been recently obtained for connexin 37 in the mouse [30].

By contrast, little attention has been paid to other types of oocyte-follicle cell contacts that appear in fact to be complex (e.g., [31]). Remarkably, the molecular composition of the adhesion structures connecting the oocyte with the follicle cells is largely unknown. In Xenopus laevis, an antibody against XB/U-cadherin has also reacted with oocyte-follicle cell contacts in the region of the forming vitelline membrane [32]. During oocyte maturation, when the oocyte microvilli retract and detach from the follicle cells, this cadherin is retrieved from the oocyte surface into an internal pool, concomitant with an enhanced translation of maternal mRNAs encoding this cadherin as well as EP/C-cadherin [32–35], which are subsequently used for the formation of junctions between blastomeres [36–38]. Plakoglobin mRNA is also accumulated in X. laevis oocytes and eggs [39], whereas in the mouse this protein has been detected only after fertilization [40]. However, the location of plakoglobin and catenins within the vertebrate ovarian follicle remains to be elucidated.

In recent years, the zebrafish (Danio rerio) has emerged as an excellent model for developmental genetics in vertebrates—systematic genome-wide mutagenesis screens having allowed the identification of genes essential during embryogenesis (e.g., [41, 42]). Given the advantages of this model, similar approaches have been recently undertaken in order to recover mutations affecting ovarian and testicular growth and development (e.g., [43]). In the present work, as a first step toward the identification of molecular constituents of the heterologous cell junctions in the ovarian follicle, we have isolated the zebrafish orthologs of the mammalian plakoglobin and catenins. The patterns of expression of the mRNAs and the resulting proteins during oogenesis, as well as their localization in the ovarian follicle, are shown.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Animals and Cell Culture

Wild-type zebrafish, purchased from a local pet store, were raised and kept under standard laboratory conditions at about 28°C [44]. The monolayer cell cultures used were human A431 carcinoma cells (American Type Culture Collection, Rockville, MD; cf. [45]) and zebrafish ZF4 fibroblastoidal cells [46] maintained under standardized conditions and grown near confluence.

Collection of Ovarian Follicles and Eggs

The ovary was dissected from females anesthetized in MS-222 (Sigma Chemical Co., St. Louis, MO) and immediately placed in 60% Leibovitz L-15 culture medium with L-glutamine (Sigma) containing 100 µg/ml gentamicin and adjusted to pH 7.5 [47]. Oocyte development was staged according to Selman et al. [47]: stage I or primary growth stage (<= 140-µm diameter); stage II or cortical alveolus stage (0.14 to 0.34-mm diameter); stage III or vitellogenic stage (0.34 to 0.69-mm diameter); stage IV or oocyte maturation stage (0.69 to 0.73-mm diameter); and stage V or ovulated oocytes, i.e., unfertilized eggs (>= 0.73-mm diameter). Ovarian follicles at stage I and II were isolated from the ovary by incubating ovarian fragments in 10 mM EDTA for approximately 30 min; stage III follicles were isolated using fine watchmaker's forceps under the stereomicroscope. To induce oocyte maturation and collect oocytes at stage IV, small ovarian fragments with follicles containing stage III oocytes were incubated in L-15 medium in the presence of 1 µg/ml of the maturation-inducing steroid, 17{alpha},20ß-dihydroxy-4-pregnen-3-one (17,20ßP), for up to ~6 h at 27–28°C [47]; and occurrence of oocyte maturation was scored by the incidence of germinal vesicle breakdown. Follicles containing both maturing and matured oocytes (stage IV) were then manually isolated. Finally, unfertilized eggs were obtained by gently stripping the ventral side of gravid females during the early morning. Ovarian fragments, isolated follicles, and eggs were frozen in liquid nitrogen and stored at -80°C until biochemical analysis.

Cloning of Zebrafish {alpha}E-Catenin and Plakoglobin

Total RNA was extracted from several ovaries and used as a template to synthesize dT-primed cDNA [9]. The cDNA synthesized from 10 µg total RNA was subjected to the polymerase chain reaction (PCR) employing degenerate oligonucleotides corresponding to the first and second conserved regions in the amino acid sequences of Drosophila {alpha}-catenin, mouse {alpha}E-catenin, and chicken {alpha}N-catenin [48]. The sequences of the degenerate oligonucleotides were as follows: forward 1: 5'-CA(A/G) GT(T/A) AC(T/C/A) AC(T/C/A) CT(G/C/T) GT(A/T) AA-3'; forward 2: 5'-(C/A)G(T/A) CA(G/A) CA(A/G) GA(A/G) CT(T/A/G) AA(G/A) GAT-3'; and reverse: 5'-AAT (T/G)AT (T/A)AC (T/C)TG (C/A/G)GG (A/G)CA (C/A)AG-3'. The PCR reaction was performed with 10 µM of each forward and reverse primer under the following cycle: 1 cycle at 95°C for 5 min; 30 cycles at 95°C for 1 min, 50°C for 2 min, and 72°C for 3 min; and a final cycle at 72°C for 7 min. The resulting cDNA fragments were ligated into pBluescriptII (SK-; Stratagene, La Jolla, CA) and sequenced on both strands using an ABI 373 DNA sequencer (Applied Biosystems, Foster City, CA). After sequence comparison using the software program package HUSAR (Heidelberg Unix Sequence Analysis Resources), one clone (O1-2; 155 bp) was identified as the zebrafish ortholog of human {alpha}E-catenin.

To isolate the full-length zebrafish {alpha}E-catenin cDNA, the EcoRI/XhoI-fragment of clone O1-2 was used to screen a zebrafish post-somitogenesis embryo cDNA library in {lambda}-ZAP II (constructed by R. Riggleman, K. Helde, and D. Grunwald, Dept. Human Genetics, Eccles Institute, University of Utah, Salt Lake City, UT) at high stringency. The same library was screened at low stringency with a fragment of 1500 nucleotides (nt) corresponding to amino acids 243 to 744 of human plakoglobin [49]. For each screening, approximately 106 phages were plated and screened with the corresponding {alpha}-32P random prime-labeled probe. Several positive phagemids were excised (ExAssist Helper Phase; Stratagene) and sequenced on both strands, and two poly(A)+ tail-bearing cDNA clones, clone A68 (2962 nt) and clone 9-4 (2664 nt), were found to contain sequence encoding {alpha}E-catenin and plakoglobin, respectively. However, when the protein sequence of clones A68 and 9-4 was translated and aligned to human {alpha}E-catenin [50] and X. laevis plakoglobin [39], respectively, it was evident that these clones were missing the putative N-terminus of the protein including the initiation methionine. Therefore, deletion fragments corresponding to the 5'-end of these clones were used as probes for the screening of an embryo neurula-stage random-primed cDNA library in {lambda}-Zap II (a gift of Prof. José Campos-Ortega, Institute for Developmental Biology, University of Cologne, Germany). After this second screening, another two partial cDNA clones were isolated, clone A54 (2088 nt) and clone P-7 (967 nt), which contained overlapping sequence to clones A68 and 9-4 of 1261 and 312 nt, respectively. These new clones contained a "Kozak sequence" [51] before the start codon, indicating that they encoded the N-terminus. Accordingly, the full-length {alpha}E-catenin cDNA was obtained by assembling clones A54 and A68, and clones P-7 and 9-4 were assembled to obtain the complete plakoglobin cDNA. The nucleotide sequence of the cDNAs encoding zebrafish {alpha}E-catenin and plakoglobin are available in GenBank/EMBL/DDBJ under accession nos. AF099737 and AF099738, respectively.

Analysis of Gene Expression

For Northern blotting, the poly(A)+ RNA was purified using the Oligotex mRNA minikit (Quiagen GmbH, Hilden, Germany), separated on formaldehyde-containing 1% agarose gel, and blotted to nylon membranes that were hybridized with 32P-labeled antisense cRNAs as described previously [9]. For detection of {alpha}E-catenin, the antisense cRNA probe was synthesized from EcoRI-linearized clone A68 with T7 RNA polymerase; for ß-catenin, clone pZ1-7H [12] was linearized with BstEII and antisense cRNA synthesized with SP6 RNA polymerase; and for plakoglobin, clone 9-4 was linearized with EcoRI and antisense cRNA synthesized with T7.

The reverse transcription (RT)-PCR assay was performed on total RNA isolated from ~100 ovarian follicles at selected developmental stages with the RNAeasy kit (Quiagen). The first-strand cDNA was synthesized from 5–10 µg RNA, and the controls omitted the RT enzyme. For the PCR, the cDNA from each stage was used in a standard PCR reaction buffer containing 1 µM of forward and reverse oligonucleotide primers: for {alpha}E-catenin, forward 5'-GCA TTC CTG TTC CTT AGT CAC C-3' (nt 3059–3080) and reverse 5'-TCA TGC TTC AAC ATT ATG CAT T-3' (nt 3282–3304); for ß-catenin, forward 5'-CCT GCA TTG TGA TTT TGG C-3' (nt 2886–2901) and reverse 5'-ATA AAA GCA AAA CTG GCC CC-3' (nt 3487–3506); and for plakoglobin, forward 5'-ATT CTC ACT TCT GGG GGA GG-3' (nt 2514–2534) and reverse 5'-AAA ATC TCC ACA CAA ACT GCG-3' (nt 3049–3070). The PCR conditions were 1 cycle at 95°C for 5 min; 35–40 cycles at 95°C for 1 min, 60°C for 2 min, and 72°C for 4 min; and a final cycle at 72°C for 7 min. As a control to ensure that the amplified bands corresponded to {alpha}E-catenin, ß-catenin, and plakoglobin, clones A68, pZ1–7H, and 9-4, respectively, were used as a template with the forward and reverse primers in the PCR. These controls were co-electrophoresed with the samples from the developmental series. Primers to the zebrafish ß-actin [52] were used as a PCR control under the same conditions; forward 5'-CCG TGA CCT GAC TGA CTA CCT-3' (nt 552–573) and reverse 5'-AAG CAT TTG CGG TGG ACG ATG-3' (nt 1110–1130). All samples were electrophoresed on 1.5–2% nondenaturing polyacrylamide gels and photographed with Polaroid (Cambridge, MA) black-and-white print film type 667 (Sigma).

Hybridization in situ on frozen sections of the ovary was performed as described previously [53]. The 35SrCTP-labeled antisense probes were synthesized as indicated for Northern analysis. The controls were hybridized with sense probes generated from clone A68 (PmlI-linearized and transcribed with T3) or from clone 9-4 (NcoI-linearized and transcribed with T3). Pictures were taken using a confocal laser-scanning microscope, Zeiss LSM 410 UV (Carl Zeiss, Jena, Germany).

Antibodies

The following monoclonal antibodies (mAbs) and rabbit antisera were used: anti-{alpha}-catenin clone 5 against mouse {alpha}-catenin (1:100–1:500; Transduction Laboratories, Lexington, KY); anti-{alpha}-catenin rabbit antisera against human/mouse {alpha}-catenin (1:1000; Sigma); anti-ß-catenin clone 14 against mouse ß-catenin (1:1000; Transduction Laboratories); anti-human plakoglobin clone 15 (1:1000; Transduction Laboratories) and PG5.1 against bovine plakoglobin (1:5; [3]); anti-E-cadherin clone 36 against human E-cadherin (1:50–1:100; Transduction Laboratories); pan-cadherin rabbit antisera against a synthetic peptide corresponding to the C-terminal amino acids of chicken N-cadherin (1:750; Sigma); and anti-N-cadherin rabbit antisera (R-851) against zebrafish N-cadherin (5 µg/ml; [54]).

Protein Extraction and Immunoprecipitation

Total protein extracts from cultured cell lines and tissues were obtained by homogenizing the samples directly in SDS-PAGE sample buffer [55], heated at 95°C for 5–10 min, and treated with 1 µl of benzonase (Merck, Darmstadt, Germany). Proteins were extracted from follicles and eggs by homogenizing the samples in Nonidet P-40 (NP-40)-containing lysis buffer (1% NP-40, 1 mM CaCl2, 150 mM NaCl, 10 mM Tris, pH 7.4, 0.5 mg/ml PMSF, and a cocktail of protease inhibitors [Mini EDTA-free; Boehringer, Mannheim, Germany]), followed by a centrifugation at 12 000 x g for 10 min at 4°C. Alternatively, the samples were initially permeabilized in lysis buffer containing digitonin (0.5 M 3-[N-morpholino]propanesulfonic acid, pH 6.8, 1 mM EGTA, 1 mM MgCl2, 0.005% digitonin, 0.5 mg/ml PMSF, and a cocktail of protease inhibitors) by pipetting up and down the samples for 3–5 min, followed by a centrifugation at 120 000 x g for 1 h at 4°C; the resulting supernatant and pellet were then homogenized in NP-40-containing buffer as described above.

For immunoprecipitation, the whole lysates or their digitonin-soluble and -insoluble fractions were precleared by incubation with free protein-A or protein-G sepharose beads for 30–60 min; they were subsequently incubated overnight at 4°C with 5–10 µl of the antibodies. Freshly prepared beads were then absorbed to the lysates for 1 h at 4°C, and bead-coupled antibodies were separated by a short centrifugation at 12 000 x g and washed with cold homogenization buffer followed by PBS. Bound proteins were eluted with SDS-PAGE sample buffer at 95°C and processed for immunoblotting.

SDS-PAGE and Immunoblotting

Proteins were separated by SDS-PAGE (7%); they were electroblotted on nitrocellulose membranes using 10 mM borate (pH 8.8) as a transfer buffer. After blocking incubation in Tris-buffered saline with 0.1% Tween (TBST) and 5% milk powder for 1–3 h, the membranes were incubated with the antibodies in TBST for 1–2 h at room temperature or overnight at 4°C. Bound antibodies were detected with horseradish peroxidase-coupled goat secondary antibodies (diluted 1:10 000) using the enhanced chemiluminescence method (ECL; Amersham, Braunschweig, Germany).

Sedimentation Analysis

Extracts from approximately 50 unfertilized eggs were layered onto 11-ml 5–30% (w:w) sucrose gradients prepared in lysis buffer without detergent. After centrifugation at 200 000 x g for 16 h at 4°C in a SW40 rotor (Beckmann Instruments, Palo Alto, CA), the gradients were fractionated from top to bottom in 400-µl aliquots. The fractions were analyzed by SDS-PAGE and immunoblotting. Reference proteins of known S values were centrifuged in parallel (catalase, 232 kDa and 11.8 S; aldolase, 158 kDa and 7.4 S; and BSA, 68 kDa and 4.9 S).

Immunofluorescence and Electron Microscopy

Immunofluorescence microscopy was done on cryostat and paraffin sections. For cryostat sections, tissues were embedded in Tissue-Tek (OCT 4583 embedding compound; Miles, Elkhart, IN) and frozen in isopentane precooled in liquid nitrogen. Tissues were sectioned (4–5 µm) and were fixed either with acetone for 10 min at -20°C or with 2% formaldehyde in PBS for 15 min followed by 5-min incubation in 50 mM NH4Cl. After fixation, sections were shortly blocked with 5% goat serum and incubated with the primary antibodies for 30–60 min, and with secondary antibodies coupled to cyanine 3.29-OSu goat (1:500; Dianova, Hamburg, Germany) for 20 min. For paraffin sections, ovarian fragments were fixed in 4% formaldehyde in PBS for 7 h, dehydrated, and embedded in paraffin. The samples were sectioned (5 µm), dried overnight at 60°C, dewaxed, and rehydrated. Sections were then transferred into urea buffer (0.1 M Tris-HCl, 5% urea, pH 9.4), microwaved at 600 W five times, 3–4 min each, and cooled to room temperature. The sections were blocked with 5% goat serum and 2% milk powder and then incubated with the primary and secondary antibodies diluted in 2% milk powder.

For standard electron microscopy, ovarian fragments were processed as previously described [56]. For immunogold electron microscopy, cryosections were mounted on coverslips, fixed in 2% formaldehyde for 12 min, and permeabilized with 0.05% saponin for 5 min. After blocking with 5% goat serum + 0.1% BSA-C (Bio Trend, Cologne, Germany) for 15 min, the primary antibodies diluted in PBS + 0.1% BSA-C were applied for approximately 2 h. After washes, the specimens were incubated with secondary antibodies coupled to 1.4-nm-diameter colloid gold particles that were applied overnight followed by silver enhancement. Secondary fixation with glutaraldehyde, dehydration, and embedding have been described previously [56].

Immunofluorescence was observed and documented with a Zeiss Axiophot photomicroscope, and electron micrographs were taken using a Zeiss electron microscope EM 910.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Isolation of Zebrafish {alpha}E-Catenin and Plakoglobin cDNAs

We isolated {alpha}E-catenin and plakoglobin cDNAs by RT-PCR and screening of embryonic zebrafish expression libraries. By recombination of two overlapping cDNAs, a full-length clone was isolated that contained an open reading frame of 3777 nt and a predicted translation product of 907 amino acid residues and a molecular weight of 100 000. The amino acid sequence of this molecule showed 90% overall identity with murine (Fig. 1A; cf. [57]) and human [50] {alpha}E-catenin, and 82% and 61% identity with mouse {alpha}N-catenin [58] and Drosophila {alpha}-catenin [48], respectively. Because of the high sequence similarity of the zebrafish cDNA to mammalian {alpha}E-catenins, extending throughout the entire length of the molecule (Fig. 1A), we concluded that the isolated clone was the zebrafish ortholog, and designated it zebrafish {alpha}E-catenin.



View larger version (41K):
[in this window]
[in a new window]
  		FIG. 1. Amino acid sequence similarity of zebrafish {alpha}E-catenin and plakoglobin to mouse {alpha}E-catenin and human plakoglobin. A) Schematic diagram showing the three domains (boxes a, b, and c) conserved among {alpha}-catenin and vinculin [48], and percentage of identity in the amino acid sequences of each domain between mouse {alpha}E-catenin (mus{alpha}E; [57]) and zebrafish {alpha}E-catenin (dan{alpha}E). The amino acid sequences from mouse {alpha}E-catenin employed as immunogen for the production of the anti-{alpha}-catenin mAb 5 (Immunogen 1, amino acid position 729-906; Transduction Laboratories) and rabbit antisera (Immunogen 2, amino acid position 890–901; Sigma) are indicated. The predicted amino acid sequence alignment of mouse and zebrafish {alpha}E-catenin at the C-terminus is also shown. The dashes indicate identical amino acids; the boxed residues indicate the conserved amino acid changes; and the dot is the gap introduced to optimize alignment. The asterisks denote the known stop codon. B) Schematic diagram of the different domains of human plakoglobin (homoPG; [49]) including the 13 armadillo repeats (numbered boxes), and percentage of identity in the amino acid sequence of each domain with zebrafish plakoglobin (danPG) predicted amino acid sequence. The epitope-containing region (ECR) of mAb PG5.1 [76], as well as the amino acid sequence of human plakoglobin employed as immunogen for the production of the anti-plakoglobin mAb 15 (Immunogen 3, amino acid position 553–738; Transduction Laboratories), is indicated. At the bottom, amino acid sequence comparison of human and zebrafish plakoglobin at the C-terminus (the symbols are as in A).

To isolate the cDNA encoding the zebrafish ortholog of plakoglobin, a cDNA fragment of human plakoglobin was used as a probe to screen a post-somitogenesis embryo cDNA library at low stringency. A partial-length clone, lacking the N-terminus, was isolated and sequenced. The corresponding 5'-end clone was then isolated by a second screening of a neurula embryo cDNA library at high stringency. The combination of the two overlapping clones resulted in a full-length cDNA that contained an open reading frame of 3304 nt, encoding a protein of 729 amino acids (molecular weight 80 000). The predicted protein was positively identified as the zebrafish ortholog of plakoglobin, as it showed 70% and 71% overall sequence identity with human [49] and X. laevis [39] plakoglobin, respectively, and only 67% identity with zebrafish ß-catenin [12]. The zebrafish plakoglobin showed a particularly high degree of identity (70–93%) with its human ortholog in the 13 "arm" repeats (Fig. 1B; cf. [59]).

Interestingly, the 3'-untranslated region (UTR) of both zebrafish cDNAs encoding {alpha}E-catenin and plakoglobin revealed three and two potential nuclear polyadenylation sequences (AAUAAA), respectively, and a number of potential cytoplasmic polyadenylation sequences (CPEs) (see nucleotide sequences AF099737 and AF099738 at GenBank/EMBL/DDBJ). This feature of the zebrafish plakoglobin cDNA contrasts with the human and mouse orthologs, which show only one [49] or no [40] CPEs.

{alpha}E-Catenin, ß-Catenin, and Plakoglobin Were Differentially Expressed in the Oocyte During Oogenesis

Northern blot experiments on total RNA from the whole ovary, using cRNA probes corresponding to the 3'-UTR of each mRNA, were carried out to determine the expression of the genes encoding {alpha}E-catenin, ß-catenin, and plakoglobin (Fig. 2A). These experiments pointed to the existence of a single mRNA species of ~3.9, ~3.3, and ~3.4 kilobases (kb) for each of the genes ({alpha}E and ß-catenin and plakoglobin, respectively), indicating that all three plaque proteins were indeed synthesized in the zebrafish ovary.



View larger version (56K):
[in this window]
[in a new window]
  		FIG. 2. Expression of zebrafish {alpha}E-catenin, ß-catenin, and plakoglobin mRNAs during oogenesis. A) Northern blot hybridization of poly(A)+ RNA (2–4 µg/lane) from the whole ovary with antisense cRNA probes to {alpha}E- and ß-catenin and plakoglobin. The positions (bars) of the reference RNAs are indicated by lines at the left margin (from top: 9.5, 7.5, 4.4, 2.4, and 1.3 kb). B) RT-PCR analysis of the gene expression for {alpha}E- and ß-catenin and plakoglobin during oogenesis. The analysis was performed on first-strand cDNAs synthesized from RNA isolated at different developmental stages. Plus or minus over each line refers to the presence or absence of RT in the cDNA synthesis. Approximately equivalent amounts of cDNA were being assayed as evident from the consistent level of the ß-actin gene. On the left is indicated the length of the DNA markers (Bluescript digested with HinfI).

RT-PCR assays, employing total RNA extracts from isolated ovarian follicles and unfertilized eggs, and sets of primers corresponding to the 3'-UTR regions for each cDNA, were used to determine the synthesis of {alpha}E- and ß-catenin and plakoglobin mRNAs during oogenesis (Fig. 2B). Results from RNA samples taken from different developmental stages of oogenesis indicated that the expected polynucleotide band was amplified and detected from stage I–II follicles up to unfertilized eggs. However, an increased expression of {alpha}E- and ß-catenin and plakoglobin appeared to occur during oocyte maturation (stage IV), followed by dramatic decreases in the unfertilized egg—except in the case of {alpha}E-catenin, for which the levels of mRNA remained very similar to those observed in maturing follicles. The relatively consistent pattern of expression of the ß-actin gene served as a control to show that equivalent amounts of cDNA were assayed in the developmental series.

To determine the spatial pattern of mRNA synthesis for {alpha}E- and ß-catenin and plakoglobin during oogenesis, we performed in situ hybridization on cryostat sections from the ovary (Fig. 3). For the three plaque proteins (Fig. 3, A–C), a positive reaction was observed in the ooplasm of oocytes at the phase of primary growth (stage I), which gradually decreased as the oocytes progressed into the cortical alveolus stage (stage II)—the hybridization signal being almost undetectable in oocytes at vitellogenic stages (stage III). However, in the somatic cells (follicle cells and theca) surrounding the oocyte, no clear labeling was observed with any probe. Control sections hybridized with sense probes (Fig. 3D) also showed negative reaction.



View larger version (153K):
[in this window]
[in a new window]
  		FIG. 3. Confocal laser-scanning micrographs of the distribution of {alpha}E-catenin (A), ß-catenin (B), and plakoglobin (C) mRNAs in follicles at different developmental stages visualized by in situ hybridization on cryostat sections from the ovary with antisense probes; D) sense (control) cRNA probes. In A the germinal vesicle is indicated by an asterisk, whereas the somatic cell layers surrounding the oocyte are indicated by arrowheads. I, Primary growth stage; II, early cortical alveolus stage; II*, mid-cortical alveolus stage; III, vitellogenic stage. Bars = 50 µm.

Antibodies Against Mammalian {alpha}-Catenin, ß-Catenin, and Plakoglobin Reacted with the Corresponding Proteins in the Zebrafish

Given the high sequence homology between the zebrafish catenins and their mammalian orthologs, we tested whether antibodies against mammalian {alpha}- and ß-catenin and plakoglobin also reacted with the respective zebrafish proteins (Fig. 4). On immunoblots (Fig. 4A) of proteins from zebrafish ZF4 cell culture (lanes 2), skin (lanes 3), and liver (lane 4), anti-{alpha}-catenin (mAb 5), anti-ß-catenin (mAb 14), and anti-plakoglobin (PG5.1) antibodies reacted specifically with components with molecular masses of ~105, ~94, and ~82 kDa, respectively (for comparison, see also the reaction with the same antibodies in A431 human carcinoma cells [lanes 1]). On immunofluorescence microscopy using zebrafish adult tissue sections of skin and liver (Fig. 4B, a–d), or cultured ZF4 cells (not shown), the antibodies reacted specifically at the cell borders, the reaction being identical to that observed in A431 cells (not shown). For {alpha}-catenin and plakoglobin, the same results were obtained when a rabbit anti-{alpha}-catenin and mAb 15 were employed (not shown). These results, therefore, proved that the antibodies tested could be used to identify unambiguously the {alpha}- and ß-catenin and plakoglobin orthologs of the zebrafish.



View larger version (87K):
[in this window]
[in a new window]
  		FIG. 4. Cross-reaction of antibodies against mammalian catenins with the same components from the zebrafish. A) Immunoblots of protein extracts from human A431 (lanes 1) and ZF4 culture cells (lanes 2) and from diverse tissues (lanes 3, skin; lane 4, liver; lane 5, ovarian follicles) probed with antibodies against {alpha}-catenin (mAb 5), ß-catenin (mAb 14), plakoglobin (mAb PG5.1), and E-cadherin (mAb 36) as indicated. The position (bars) of molecular weight markers is indicated on the left (from top to bottom [x 10-3]: 205, 116, 97, and 66). B) Immunolocalization of the components detected in the immunoblots on cryostat sections from different adult tissues (a, mAb 5 on skin; b, mAb 14 on liver; c, mAb PG5.1 on skin; d, mAb 36 on skin). Identical results on immunoblots and immunofluorescence microscopy were obtained with anti-{alpha}-catenin and pan-cadherin rabbit antibodies, and with the plakoglobin mAb 15 (data not shown).

Antibodies against human E-cadherin, as well as pan-cadherin rabbit antibodies and mAbs (not shown) against chicken N-cadherin, were also tested as controls. Interestingly, the anti-E-cadherin mAb 36 (Fig. 4A) did not react with any component in protein extracts from ZF4 cell culture but recognized a higher molecular mass component (~150 kDa) than that observed in A431 cells (~120 kDa) in protein extracts from the skin (lane 3) and from ovarian follicles (lane 5). This ~150-kDa polypeptide band was also detected by the pan-cadherin antibodies (not shown). Immunolocalization on cryosections from the zebrafish skin with these antibodies was confined to cell-cell contacts (Fig. 4B, d), as also seen in A431 cells (not shown), which is consistent with the concept of a membrane-associated protein in the junctions between zebrafish cells.

Catenins Were Complexed with Cadherins in the Ovarian Follicle During Oogenesis

Using the antibodies previously characterized, as well as antibody R-851 against zebrafish N-cadherin, the cadherin-catenin interactions in ovarian follicles and eggs were investigated by immunoprecipitation followed by immunoblotting (Fig. 5). Detergent extracts were prepared from isolated stage III follicles (Fig. 5A) or unfertilized eggs (Fig. 5B), and immunoprecipitates with anti-{alpha}-catenin, anti-ß-catenin, and anti-plakoglobin were analyzed for the presence of cadherins and catenins.



View larger version (46K):
[in this window]
[in a new window]
  		FIG. 5. Analysis of cadherin-catenin complexes in ovarian follicles (A) and unfertilized eggs (B) by immunoprecipitation. Whole lysates from ~200 stage III follicles or from ~100 unfertilized eggs were immunoprecipitated (IP) with anti-{alpha}-catenin (mAb 5), anti-ß-catenin (mAb 14), and anti-plakoglobin (PG5.1) mAbs (lanes 2) and analyzed by immunoblotting with the indicated antibodies. For the immunoprecipitation controls (lanes 1), mouse mAb against human fibronectin was employed in parallel. In unfertilized eggs, N-cadherin was not detected in the ß-catenin IP; in the plakoglobin IP, neither E-cadherin-like protein(s) nor N-cadherin was detected (not shown). The position (bars) of molecular weight markers is indicated on the left (from top to bottom [x 10-3]: 205, 116, 97, and 66).

Immunoprecipitated proteins from ovarian follicle extracts revealed the specific polypeptides reacting with catenin antibodies, indicating the presence of all three catenins (Fig. 5A). In addition, when anti-{alpha}-catenin was used for immunoprecipitation, both ß-catenin and plakoglobin were detected in the precipitates, whereas immunoprecipitates with anti-ß-catenin reacted only with {alpha}-catenin and not with plakoglobin. Immunoprecipitates with either anti-{alpha}-catenin, anti-ß-catenin, or anti-plakoglobin also contained E-cadherin-like protein(s) as well as N-cadherin (see also Fig. 5A for controls, lane 1). These results suggested the presence of catenin-cadherin complexes in zebrafish ovarian follicles, containing either N- or E-cadherin-like protein.

In unfertilized eggs (Fig. 5B), {alpha}- and ß-catenin as well as plakoglobin were also present, whereas E-cadherin-like component(s) and N-cadherin were not detected under the conditions employed (not shown). However, immunoprecipitates with ß-catenin also contained {alpha}-catenin and even showed a weak reaction with E-cadherin antibodies, thus also indicating the presence of E-cadherin-like protein(s) in the egg, although possibly in low amounts and—at least sometimes—complexed to {alpha}- and/or ß-catenin. In the same egg extracts, plakoglobin coprecipitated in low amounts with {alpha}-catenin, as judged by the very low signal intensity (Fig. 5B; see lane 2 after immunoprecipitation with plakoglobin antibody). Neither ß-catenin nor E-cadherin was found in the precipitates (not shown).

In order to examine possible changes of the cadherin-catenin complex during oogenesis, digitonin-soluble and -insoluble fractions from follicles at different developmental stages, and from unfertilized eggs, were precipitated with catenin antibodies; the precipitates were analyzed for the presence of {alpha}- and ß-catenin, plakoglobin, and an E-cadherin-like glycoprotein(s) (Fig. 6A). The digitonin-insoluble fractions from all follicle stages contained considerable amounts of all three catenins in complexes with E-cadherin-like component(s), while catenins not complexed to cadherins were detected in the specific soluble fractions in which heterodimers of {alpha}- with ß-catenin, as well as of {alpha}-catenin with plakoglobin, were also observed (not shown). The proportion between soluble and insoluble plaque proteins did not appear to change with advancing development. In unfertilized eggs, however, the proportion of solubilized plakoglobin with respect to the cadherin-complexed fraction was apparently higher than that of {alpha}- and ß-catenin.



View larger version (28K):
[in this window]
[in a new window]
  		FIG. 6. A) Coprecipitation of {alpha}- and ß-catenin and plakoglobin with E-cadherin-like components from digitonin-soluble (lane S) and -insoluble (lane I) extracts of developing follicles and unfertilized eggs (the developmental stages are indicated). Immunoprecipitates (IP) obtained with anti-{alpha}-catenin (mAb 5), anti-ß-catenin (mAb 14), or anti-plakoglobin (PG5.1) were analyzed by immunoblotting for these proteins, and for E-cadherin-like components using the anti-E-cadherin mAb 36. The controls immunoprecipitated in parallel with fibronectin mAb did not show any of the specific components (not shown). B) Separation of E-cadherin-like component(s), {alpha}-catenin, ß-catenin, and plakoglobin by sucrose gradient centrifugation. The fractions (fraction number from top to bottom) were analyzed for E-cadherin-like protein(s), {alpha}- and ß-catenin, and plakoglobin with the same antibodies as in A. Note that the polypeptide bands appeared as broad bands or displaced to the bottom of the gel because of the yolk present in the samples that was not removed prior to the gradient centrifugation. The arrowheads indicate the peak distribution of protein standards (from left to right: BSA, 4.9 S; aldolase, 7.4 S; catalase, 11.8 S).

To address the question whether the detergent-extractable ooplasmic catenins and plakoglobin are free or complexed, sedimentation velocity experiments were performed: extracts from approximately 50 eggs were fractionated by sucrose gradient centrifugation, and the individual fractions were assayed by SDS-PAGE and immunoblotting for {alpha}- and ß-catenin, plakoglobin, and E-cadherin-like components (Fig. 6B). In these analyses, complexes containing E-cadherin-like protein were recovered in fractions 11–15, which also contained both {alpha}- and ß-catenin, as well as some plakoglobin. In addition, considerable proportions of plakoglobin, as well as some ß-catenin, sedimented with lower S values (5–7S; fractions 6–9), indicative of monomers or dimers (for analysis of 7S plakoglobin homodimers, see [60, 61]).

In summary, these experiments indicated that in unfertilized eggs, major proportions of {alpha}- and ß-catenin appeared in complexes with cadherin, while the bulk of plakoglobin seemed to occur as a free cytoplasmic form.

Immunolocalization of Plakoglobin and Catenins in the Ovarian Follicle

Immunofluorescence and immunoelectron microscopy were used to determine sites of enrichment of plaque proteins and cadherins. Figure 7 summarizes the immunofluorescence results on paraffin or cryostat sections through the ovary.



View larger version (126K):
[in this window]
[in a new window]
  		FIG. 7. Immunofluorescence microscopy on paraffin (A–D and F–H) and frozen (E) sections from the ovary after reaction with anti-{alpha}-catenin (rabbit antisera; A, B), anti-ß-catenin (mAb 14; C, D), anti-plakoglobin (mAb 15; E), anti-E-cadherin (mAb 36; F, G), and anti-N-cadherin (R-851; H) antibodies. A–C and G) Phase-contrast optics; (A'–C', B'', G', and D–F, H) epifluorescence. A, A') Section through early stage II follicles showing immunostaining for {alpha}-catenin in the oocyte cortical ooplasma (the germinal vesicle is indicated by an asterisk, and the arrowheads indicate the follicle cells). B, B', B'') Section through two adjacent follicles at stage III displaying punctate reaction in follicle cells (arrowhead) and in the cortical ooplasm (arrow). The oocyte is indicated by an asterisk. B'') {alpha}-Catenin staining within the microvilli from the vitelline envelope (white arrowhead). C, C') Follicles from stage I to stage II with increasing immunostaining corresponding to ß-catenin in the oocyte cortex and in follicle cells (white arrowhead). D) Stage III follicles showing ß-catenin reaction at the follicle cell borders (arrow), as well as a punctate decoration in the oocyte cortex (arrowhead). E) Section through a late stage II follicle showing positive reaction for plakoglobin only in follicle cell-cell contacts. F and G, G') Follicles at stage I and II displaying a continuous immunostaining for E-cadherin-like molecule(s) at the oocyte cortex (F), which became restricted into a punctate decoration at intercellular contacts between follicle cells and the oocyte (arrowhead) in stage III follicles (G'); the surrounding follicle cells (arrow) also showed E-cadherin-like proteins at the cell boundaries. H) Localization of N-cadherin at intercellular contacts in the follicular cell layer of stage III follicles. Bar = 50 µm.

Positive reactions for {alpha}- and ß-catenin, as well as for E-cadherin-like protein, were found in follicles at different stages of development (Fig. 7, A–D, F, G). Immunostaining for {alpha}- and ß-catenin and E-cadherin was already observed in follicles of the primary growth phase (stage I), and appeared as a continuous decoration in the cortical ooplasm (Fig. 7, C and F). As oocytes advanced into the cortical alveolus stage (stage II), characterized by the formation of the cortical alveoli and the vitelline envelope (i.e., zona radiata), the staining for these proteins in the oocyte cortex adjacent to the vitelline envelope became stronger, accentuating the zone of contacts between the oocyte and the surrounding follicle cells (Fig. 7, A, C, F). When the follicles reached growth stage III, characterized by the accumulation of yolk, the pattern of immunostaining of {alpha}- and ß-catenin and E-cadherin-like components changed dramatically, now resulting in a conspicuous punctate decoration at the oocyte border and at the cell-cell contacts with the surrounding follicle cells (Fig. 7, B, D, G). In addition, the signals for these junctional proteins appeared in the microvilli projecting into the vitelline envelope (Fig. 7, B-B'', G). By contrast, plakoglobin immunostaining using two different mAbs (PG5.1 and mAb 15) in the oocyte was not so pronounced at any developmental stage examined, whereas it was strongly delineating the cell-cell borders of the surrounding follicle cells (Fig. 7E). Similarly, reaction with N-cadherin antibodies was restricted to the cell-cell contacts in the follicle cell layer (Fig. 7H) and was detected only from stage II onward.

When manually isolated stage III follicles were induced to mature in vitro with the maturation-inducing steroid 17,20ßP and then analyzed by immunofluorescence microscopy, maturation was resumed, as indicated by germinal vesicle breakdown, and the punctate pattern of {alpha}- and ß-catenin as well as of E-cadherin-like proteins at the oocyte-follicle cell boundaries disappeared (Fig. 8, A–C). By contrast, the immunostaining at the follicle cell contacts remained intense. Similarly, the plakoglobin staining at the follicle cell boundaries was not visibly affected by the treatment with the hormone (not shown).



View larger version (73K):
[in this window]
[in a new window]
  		FIG. 8. Immunofluorescence microscopy on paraffin sections from follicles matured in vitro with 17,20ßP. Sections were immunostained with the same antibodies as in Figure 7. Note the absence of {alpha}-catenin (A), ß-catenin (B), and E-cadherin-like protein (C, phase-contrast optics; C', epifluorescence) from the intercellular junctions between the oocyte and follicle cells, whereas the signals for these components at the junctions between follicle cells remained unchanged. The plakoglobin staining at follicle cell contacts also remained unchanged during oocyte maturation (not shown). C shows the change in the contour of the yolk bodies (asterisks) that occurs during oocyte maturation (compare with Fig. 7G). At this oocyte stage, the method of staining employed seemed to cause an increased autofluorescence of the yolk platelets and cortical alveoli (arrow), which was more evident with the anti-{alpha}-catenin rabbit antisera (A). Bar = 50 µm.

Conventional electron microscopy, as well as immunoelectron microscopy using immunogold labeling, was applied to localize catenins and cadherins within the ovarian follicle (Fig. 9). Typical desmosomes with intermediate filament bundles attached (brackets in Fig. 9A), adherens junctions, and gap junctions (not shown) were frequently observed between follicle cells. In stage I follicles, short oocyte projections, presumably microvilli in statu nascendi, formed focal contacts with the surrounding follicle cells (Fig. 9B). In stage II, microvillous projections, now containing microfilament bundles, extended through the "pore canals" of the vitelline envelope to form junctional contacts with follicle cells (Fig. 9, C–E).



View larger version (135K):
[in this window]
[in a new window]
  		FIG. 9. Ultrastructure of junctional complexes in the ovarian follicle (A–E), and immunogold localization of {alpha}-catenin, ß-catenin, plakoglobin, or E-cadherin-like protein at these junctions using anti-{alpha}-catenin rabbit antisera (F, I), anti-ß-catenin mAb 14 (J), PG5.1 (G, H), and anti-E-cadherin mAb 36 (K). Normal fixed (A–E) or saponin-permeabilized (F–K) cryosections. A) Stage I follicle with typical-appearing desmosomes (brackets) between two follicle cells surrounding the oocyte, which projected short microvilli toward the overlying follicle cells. The vitelline envelope (asterisk) was not yet formed. B) By approximately the same stage, the oocyte started to make "focal-type"-like contacts with the surrounding follicle cells. C) Lower-power electron micrograph of a stage II follicle. The oocyte elongated microvilli through pore canals located in the vitelline envelope (arrowheads) toward a single layer of follicle cells. External to the follicle cell layer were a basal lamina, the vascularized theca layer, and a thin surface epithelium. D, E) In stage II follicles, the points of contact between oocyte and follicle cell processes appeared associated with electron-dense material (brackets), and microfilaments along the microvilli can also be observed (arrows). F) Localization of {alpha}-catenin at adherens junctions between follicle cells. G, H) Plakoglobin immunoreaction in both desmosome (G) and adherens junction (H) plaques (brackets) in follicle cells. The desmosome-associated bundles of intermediate filaments are indicated by arrows in (G). I) Stage II follicle showing strong signal for {alpha}-catenin at the oocyte cortex adjacent to the vitelline envelope (asterisk); the signal at the overlying follicle cells was not as strong as in the oocyte. J) Immunogold localization of ß-catenin at points of contact between follicle cell and oocyte microvilli. K) Localization of E-cadherin-like protein at the cortex of an early stage II oocyte and in its follicle cells. Bars = 1 µm (A–C), 0.5 µm (D–K). O, oocyte; FC, follicle cells; BL, basal lamina; T, theca; SE, surface epithelium; V, vitelline envelope.

Immunogold localization revealed the presence of {alpha}- and ß-catenin as well as of E-cadherin-like proteins in adherens junctions, but not in desmosomes, between follicle cells (Fig. 9F). Plakoglobin immunoreaction was found in the plaques of both adherens junctions and desmosomes (Fig. 9, G and H). Interestingly, the entire oocyte cortex of stage II follicles showed an intense immunogold reaction with anti-{alpha}-catenin (Fig. 9I), anti-ß-catenin (not shown), and anti-E-cadherin antibodies (Fig. 9K), indicating high concentrations of these proteins in the cortical ooplasm. The points of contact between oocyte microvilli and follicle cells were strongly labeled for ß-catenin (Fig. 9J), as well as for {alpha}-catenin and E-cadherin-like protein (not shown).


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
In the present study we have identified the zebrafish homologues of {alpha}E-catenin and plakoglobin and have shown that these proteins, together with their complex partners in adherens junctions—i.e., ß-catenin and an E-cadherin equivalent—are abundantly synthesized and stored during oogenesis. Our finding that these proteins can be localized to specific junctional structures connecting the oocyte with the follicle epithelial cells points to a function in establishing follicle architecture; and the maintenance of these proteins in the unfertilized egg suggests an additional function as a maternal pool contributing, among other things, to junction formation during early embryogenesis.

{alpha}-Catenins are cadherin-associating proteins of vertebrates composed at least of two members, {alpha}E and {alpha}N-catenin, which are distantly related in amino acid sequence to vinculin and are required for organizing cadherins and tight junctions, and hence essential in multicellular structures (see, e.g. [62–64]). The zebrafish ortholog of mammalian {alpha}E-catenin presents 90% identity with {alpha}E-catenin and 82% identity with {alpha}N-catenin of the mouse (cf. [57, 65]), suggesting the presence of at least two {alpha}-catenin isoforms also in the zebrafish. This was corroborated when a different partial-length cDNA clone of 2711 nt with 90–93% identity to chick [65], mouse [58], and human [66] {alpha}N-catenin was isolated from a cDNA library derived from adult zebrafish (data not shown).

ß-Catenin and plakoglobin are other constitutive proteins of the plaques associated with cadherins in adherens junctions (see Introduction), and plakoglobin is the only protein known to be common to both adherens junctions and desmosomes [3]. Both proteins are unrelated to the {alpha}-catenins, but they share more than 60% identity and belong to a large multigene family, which includes the product of the Drosophila segment polarity armadillo, the tumor suppressor APC protein, protein p120, plakophilins 1–3, and the neural plakophilin-related arm-repeat protein (NPRAP)—all characterized by variable numbers of repeats of a motif of ~42 amino acids, called arm-repeats (see [1, 9, 59]). The zebrafish plakoglobin shows ~70% identity with human [49] and X. laevis [39] plakoglobin, which, however, is only observed in the 13 arm-repeats. Interestingly, the sequence identity of plakoglobin and ß-catenin [12] in the zebrafish is 67%, i.e., similar to that between mammalian plakoglobin and ß-catenin [3, 67]. These findings, together with the molecular features of zebrafish {alpha}E-catenin, suggest that the divergence of catenins throughout evolution already took place during the appearance of lower vertebrates.

Remarkably, the pattern of synthesis of {alpha}- and ß-catenin as well as of E-cadherin-related proteins in the oocyte appears to correlate with the establishment of heterotypic cell contacts between oocytes and follicle cells throughout folliculogenesis. In stages in which microvilli-like cell processes project from the oocyte surface through the developing vitelline "membrane" and form focal contacts with the follicle cells ([23, 47], Fig. 9B), related to the puncta adhaerentia of other cells (cf. [1]), {alpha}- and ß-catenin, as well as E-cadherin-like proteins, accumulated in the oocyte cortex, in close proximity to the oocyte-follicle cell boundary. When oocytes advanced into vitellogenesis (stage III) and their microvilli elongated through the pore canals of the fully formed vitelline envelope, the localization of both catenins and E-cadherin-like protein in the oocyte became restricted to a more punctate pattern, reflecting the points of contact between oocytes and follicle cells. Their coimmunolocalization at the relatively rare sites of plaque-bearing junctions between oocytes and follicle cells strongly suggests a role of these components in the heterologous adhesion of both cell types during oogenesis. This finding thus confirms and extends previous reports for X. laevis XB/U-cadherin [32]. These complexes obviously also play a role in anchoring the actin filament bundles of the developing oocyte (for amphibia see [20, 68]). This view would be also in agreement with our finding that ß-catenin coprecipitated with {alpha}-catenin and E-cadherin-like proteins in both developing follicles and unfertilized eggs, resembling the situation in the mouse where {alpha}- and ß-catenin appear complexed to E-cadherin in mature oocytes [40]. However, since our immunoprecipitation experiments were carried out with follicle-enclosed oocytes, the contribution of the follicle cell complexes to the immunoprecipitates could not be separately determined; thus conclusive biochemical evidence for the presence of cadherin-catenin complexes at the oocyte cell membrane is still lacking.

Interestingly, in oocytes of stages I and II, {alpha}- and ß-catenin as well as E-cadherin-like protein were labeled more strongly in the oocyte cortex than in the surrounding follicle cells. As already mentioned, at these stages the oocyte projects microvilli toward the follicle cells that eventually elongate considerably, reaching up to 8 µm in length, also extending deeply into the spaces between adjacent follicle cells; in contrast, those projecting from the follicle cells are much less numerous and frequently do not reach the oocyte surface [23, 47]. For a long time it has been believed that follicle cells are the main regulators of oocyte growth and differentiation in vertebrates, but a number of studies support the hypothesis of an active role of the oocyte in regulating the function of the surrounding follicle cells (see [69, 70] and references therein). Therefore, examination of the emergence of the adhesion pattern between germ and somatic cells during early oogenesis, including investigation of the origin of the signaling pathways involved, would be of considerable interest.

Unlike that found for {alpha}E- and ß-catenin, plakoglobin mRNA synthesis in the oocyte was not closely correlated with an increased localization of this protein in the cortical ooplasm (for control, note that this protein was specifically localized in adherens junction and desmosome plaques in the surrounding follicle cells). Whether the changes observed in plakoglobin synthesis during oocyte growth and its low concentration in the cortex are influenced by mRNA or protein masking [71], or by proteolysis, remains to be seen (for X. laevis oocytes, see [39]).

During zebrafish oocyte maturation, mRNAs for catenins and plakoglobin seem to be accumulated, and this appears to be associated with the dissociation of {alpha}- and ß-catenin from the oocyte membrane. Concomitantly, the removal of E-cadherin-related protein has also been noted, and this has also been reported in X. laevis oocytes for cadherins and ß1-integrin [32, 72]. Moreover, in X. laevis an increased translation of mRNAs encoding XB/U- and EP/C-cadherin as well as ß1-integrin [32–35, 38, 72] has also been reported and suggested to contribute to a maternal pool important for the adhesion of blastomeres. Similarly, Ohsugi et al. [40] have reported that {alpha}- and ß-catenin, as well as E-cadherin, are stored in unfertilized mouse eggs.

The presence of ß-catenin in zebrafish eggs is also consistent with a previous report on this species [12], as well as on X. laevis early embryos (e.g., [73]), and with the observation of a transient nuclear localization of maternal ß-catenin in blastula-stage zebrafish embryos [74]. The maternal pool of plakoglobin has also been described in X. laevis [14, 39], and in this species its functional importance has been demonstrated by the finding that depletion of maternal plakoglobin causes a loss of cell adhesion and a distorted shape in blastula-stage embryos [75]. With the present study, we have established the basis for similar function-oriented experiments during zebrafish embryogenesis.


   ACKNOWLEDGMENTS
 
We are grateful to Drs. R. Moon, J. Campos Ortega, D. Grunwald, and B. Geiger for providing clones, cDNA libraries, and antibodies. We also thank Dr. Michael Brand for use of his fish-breeding facility, and Dr. Harald Herrmann for constructive discussions and reading of the manuscript. The excellent help of C. Grund and of A. Hunziker is greatly appreciated.


   FOOTNOTES
 
1 Financial support was provided by the Deutsche Forschungsgemeinschaft (DFG). Participation of J.C. was financed by a postdoctoral fellowship from the European Commission (Training and Mobility of Researchers Program). Back

2 Correspondence: waken{at}ctv.es Back

Accepted: April 19, 1999.

Received: February 11, 1999.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Schmidt A, Heid HW, Schäfer S, Nuber UA, Zimbelman R, Franke WW. Desmosomes and cytoskeletal architecture in epithelial differentiation: cell type-specific plaque components and intermediate filament anchorage. Eur J Cell Biol 1994; 65:229–245.[Medline]
  2. Takeichi M. Morphogenetic roles of classic cadherins. Curr Opin Cell Biol 1995; 7:619–627.[CrossRef][Medline]
  3. Cowin P, Kapprell H-P, Franke WW, Tamkun J, Hynes RO. Plakoglobin: a protein common to different kinds of intercellular adhering junctions. Cell 1986; 46:1063–1073.[CrossRef][Medline]
  4. Kapprell H-P, Owaribe K, Franke WW. Identification of a basic protein of Mr 75000 as an accessory desmosomal plaque protein in stratified and complex epithelia. J Cell Biol 1988; 106:1679–1691.[Abstract/Free Full Text]
  5. Buxton RS, Magee AI. Structure and interactions of desmosomal and other cadherins. Semin Cell Biol 1992; 3:157–167.[Medline]
  6. Garrod DR. Desmosomes and hemidesmosomes. Curr Opin Cell Biol 1993; 5:30–40.[CrossRef][Medline]
  7. Koch PJ, Franke WW. Desmosomal cadherins: another growing multigene family of adhesion molecules. Curr Opin Cell Biol 1994; 6:682–687.[CrossRef][Medline]
  8. Mertens C, Kuhn C, Franke WW. Plakophilins 2a and 2b: constitutive proteins of dual localization in the karioplasm and the desmosomal plaque. J Cell Biol 1996; 135:1009–1025.[Abstract/Free Full Text]
  9. Schmidt A, Langbein L, Rode M, Prätzel S, Zimbelmann R, Franke WW. Plakophilins 1a and 1b: widespread nuclear proteins recruited in specific epithelial cells as desmosomal plaque components. Cell Tissue Res 1997; 290:481–499.[CrossRef][Medline]
  10. Heasman J, Crawford A, Goldstone K, Garner-Hamrick P, Gumbiner B, McCrea P, Kintner C, Noro CY, Wylie C. Overexpression of cadherins and underexpression of ß-catenin inhibit dorsal mesoderm induction in early Xenopus embryos. Cell 1994; 79:791–803.[CrossRef][Medline]
  11. Larue L, Ohsugi M, Hirchenhain J, Kemler R. E-Cadherin null mutant embryos fail to form a trophectoderm epithelium. Proc Natl Acad Sci USA 1994; 91:8263–8267.[Abstract/Free Full Text]
  12. Kelly GM, Erezyilmaz DF, Moon RT. Induction of a secondary embryonic axis in zebrafish occurs following the overexpression of ß-catenin. Mech Dev 1995; 53:261–273.[CrossRef][Medline]
  13. Haegel H, Larue L, Ohsugi M, Fedorov L, Herrenknecht K, Kemler R. Lack of ß-catenin affects mouse development at gastrulation. Development 1995; 121:3529–3537.[Abstract]
  14. Karnovsky A, Klymkowsky M. Anterior axis duplication in Xenopus induced by the over-expression of the cadherin-binding protein plakoglobin. Proc Natl Acad Sci USA 1995; 92:4522–4526.[Abstract/Free Full Text]
  15. Ruiz P, Brinkmann V, Ledermann B, Behrend M, Grund C, Thalhammer C, Vogel F, Birchmeier C, Gunther U, Franke WW, Birchmeier W. Targeted mutation of plakoglobin in mice reveals essential functions of desmosomes in the embryonic heart. J Cell Biol 1996; 135:215–225.[Abstract/Free Full Text]
  16. Bierkamp C, McLaughlin KJ, Swarz H, Huber O, Kemler R. Embryonic heart and skin defects in mice lacking plakoglobin. Dev Biol 1996; 180:780–785.[CrossRef][Medline]
  17. Wylie C, Kofron M, Payne C, Anderson R, Hosobuchi M, Joseph E, Heasman J. Maternal ß-catenin establishes a "dorsal signal" in early Xenopus embryos. Development 1996; 122:2987–2996.[Abstract]
  18. Zamboni L. Fine morphology of the follicle wall and follicle cell-oocyte association. Biol Reprod 1974; 10:125–149.[Abstract]
  19. Anderson E, Albertini DF. Gap junctions between the oocyte and the companion follicle cells in the mammalian ovary. J Cell Biol 1976; 71:680–686.[Abstract/Free Full Text]
  20. Franke WW, Rathke PC, Seib E, Trendelenburg MF, Osborn M, Weber K. Distribution and mode of arrangement of microfilaments structures and actin in the cortex of the amphibian oocyte. Eur J Cell Biol 1976; 14:111–130.
  21. Toshimori K, Yasuzumi F. Gap junctions between microvilli of an oocyte and follicle cells in the teleost (Plecoglosus altivelis). Z Mikrosk-Anat Forsch (Leipz) 1979; 93:458–464.
  22. Browne CL, Werner W. Intercellular junctions between the follicle cells and oocytes of Xenopus laevis. J Exp Zool 1984; 230:105–113.[CrossRef][Medline]
  23. Kessel RG, Roberts RL, Tung HN. Intercellular junctions in the follicular envelope of the teleost, Brachydanio rerio. J Submicrosc Cytol 1988; 20:415–424.
  24. Motta PM, Makabe S, Naguro T, Correr S. Oocyte follicle cells association during development of human ovarian follicle. A study by high resolution scanning and transmission electron microscopy. Arch Histol Cytol 1994; 57:369–394.[Medline]
  25. Gilula NB, Epstein ML, Beers WH. Cell-to-cell communication and ovulation: a study of the cumulus-oocyte complex. J Cell Biol 1978; 78:58–75.[Abstract/Free Full Text]
  26. Moor RM, Cran DG. Intercellular coupling in mammalian oocytes. In: Johnson MH (ed.), Development in Mammals, vol 4. Amsterdam: Elsevier, North-Holland; 1980: 3–37.
  27. Eppig JJ. The relationship between cumulus cell-oocyte coupling, oocyte meiotic maturation, and cumulus expansion. Dev Biol 1982; 89:268–272.[CrossRef][Medline]
  28. Racowski C. Effect of forskolin on the spontaneous maturation and cyclic AMP content of rat oocyte-cumulus complexes. J Reprod Fertil 1984; 72:107–116.[Abstract/Free Full Text]
  29. Cerdà JL, Petrino TR, Wallace RA. Functional heterologous gap junctions in Fundulus ovarian follicles maintain meiotic arrest and permit hydration during oocyte maturation. Dev Biol 1993; 160:228–235.[CrossRef][Medline]
  30. Simon AM, Goodenough DA, Li E, Paul DL. Female infertility in mice lacking connexin 37. Nature 1997; 385:525–529.[CrossRef][Medline]
  31. Albertini DF, Rider V. Patterns of intercellular connectivity in the mammalian cumulus-oocyte complex. Microsc Res Tech 1994; 27:125–133.[CrossRef][Medline]
  32. Müller AHJ, Angres B, Hausen P. U-cadherin in Xenopus oogenesis and oocyte maturation. Development 1992; 114:533–543.[Abstract/Free Full Text]
  33. Choi YS, Sehgal R, McCrea P, Gumbiner B. A cadherin-like protein in eggs and cleaving embryos of Xenopus laevis is expressed in oocytes in response to progesterone. J Cell Biol 1990; 110:1575–1582.[Abstract/Free Full Text]
  34. Herzberg F, Wildermuth V, Wedlich D. Expression of XBcad, a novel cadherin, during oogenesis and early development. Mech Dev 1991; 35:33–42.[CrossRef][Medline]
  35. Kühl M, Wedlich D. XB/U-Cadherin mRNA contains cytoplasmic polyadenylation elements and is polyadenylated during oocyte maturation in Xenopus laevis. Biochim Biophys Acta 1995; 1262:95–98.[Medline]
  36. Ginsberg D, DeSimone D, Geiger B. Expression of a novel cadherin (EP-cadherin) in unfertilized eggs and early Xenopus embryos. Development 1991; 111:315–325.[Abstract]
  37. Angres B, Müller AHJ, Kellermann J, Hausen P. Differential expression of two cadherins in Xenopus laevis. Development 1991; 111:829–844.[Abstract]
  38. Müller H-AJ, Kühl M, Finnemann S, Schneider S, van der Poel SZ, Hausen P, Wedlich D. Xenopus cadherins: the maternal pool comprises distinguishable members of the family. Mech Dev 1994; 47:213–223.[CrossRef][Medline]
  39. Fouquet B, Zimbelmann R, Franke WW. Identification of plakoglobin in oocytes and early embryos of Xenopus laevis: maternal expression of a gene encoding a junctional plaque protein. Differentiation 1992; 51:187–194.[CrossRef][Medline]
  40. Ohsugi M, Hwang S-Y, Butz S, Knowles BB, Solter D, Kemler R. Expression and cell membrane localization of catenins during mouse preimplantation development. Dev Dynam 1996; 206:391–402.[CrossRef][Medline]
  41. Haffter P, Granato M, Brand M, Cullins MC, Hammerschmidt M, Kane DA, Odenthal J, van Eeden JM, Jiang Y-J, Heisenberg C-P, Kelsh RN, Furutani-Seiki M, Vogelsang E, Beuchle D, Schach U, Fabian C, Nüsslein-Volhard C. The identification of genes with unique and essential functions in the development of the zebrafish, Danio rerio. Development 1996; 123:1–36.[Abstract]
  42. Driever W, Solnica-Krezel L, Schier AF, Neuhauss SCF, Malicki J, Stemple DL, Stainier DYR, Zwartkruis F, Abdelilah S, Rangini Z, Belak J, Boggs C. A genetic screen for mutations affecting embryogenesis in zebrafish. Development 1996; 123:37–46.[Abstract]
  43. Bauer MP, Goetz FW. Zebrafish mutagenesis: a screen for reproductive mutants. Biol Reprod 1998; 58(suppl 1):104 (abstract 104).
  44. Westerfield M. The Zebrafish Book. Edition 2.1. Eugene, Oregon: University of Oregon Press; 1995.
  45. Troyanovsky SM, Eshkind LG, Troyanovsky RB, Leube RE, Franke WW. Contributions of cytoplasmic domains of desmosomal cadherins to desmosome assembly and intermediate filament anchorage. Cell 1993; 72:561–574.[CrossRef][Medline]
  46. Driever W, Rangini Z. Characterization of a cell line derived from zebrafish (Brachydanio rerio) embryos. In Vitro Cell Dev Biol 1993; 29A:749–754.
  47. Selman K, Wallace RA, Sarka A, Qi X. Stages of oocyte development in the zebrafish, Brachydanio rerio. J Morphol 1993; 218:203–224.[CrossRef]
  48. Oda H, Uemura T, Shiomi K, Nagafuchi A, Tsukita S, Takeichi M. Identification of a Drosophila homologue of {alpha}-catenin and its association with the armadillo protein. J Cell Biol 1993; 121:1133–1140.[Abstract/Free Full Text]
  49. Franke WW, Goldschmidt MA, Zimbelman R, Mueller HM, Schiller DL, Cowin P. Molecular cloning and amino acid sequence of human plakoglobin, the common junctional plaque protein. Proc Natl Acad Sci USA 1989; 86:4027–4031.[Abstract/Free Full Text]
  50. Rimm DL, Kebriacel P, Morrow JS. Molecular cloning reveals alternative splice forms of human {alpha}(E)-catenin. Biochem Biophys Res Commun 1994; 203:1691–1699.[CrossRef][Medline]
  51. Kozak M. Point mutations define a sequence flanking the AUG initiator codon that modulates translation by eukaryotic ribosomes. Cell 1986; 44:283–292.[CrossRef][Medline]
  52. Kelly G, Reversade B. Characterization of a cDNA encoding a novel band 4.1-like protein in zebrafish. Biochem Cell Biol 1999; (in press).
  53. Rogers MA, Langbein L, Prätzel S, Moll I, Krieg T, Winter H, Schweizer J. Sequences and differential expression of three novel human type II hair keratins. Differentiation 1997; 61:187–194.[CrossRef][Medline]
  54. Bitzur S, Kam Z, Geiger B. Structure and distribution of N-cadherin in developing zebrafish embryos: morphological effects of ectopic over-expression. Dev Dyn 1994; 201:121–136.[Medline]
  55. Laemmli UK. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 1970; 227:680–685.[CrossRef][Medline]
  56. Rose O, Grund C, Reinhardt S, Starzinski-Powitz A, Franke WW. Contactus adherens, a special type of plaque-bearing adhering junction containing M-cadherin, in the granule cell layer of the cerebellar glomerulus. Proc Natl Acad Sci USA 1995; 92:6022–6026.[Abstract/Free Full Text]
  57. Herrenknecht K, Ozawa M, Eckerskorn C, Lottspeich F, Lenter M, Kemler R. The uvomorulin-anchorage protein alpha catenin is a vinculin homologue. Proc Natl Acad Sci USA 1991; 88:9156–9160.[Abstract/Free Full Text]
  58. Uchida N, Shimamura K, Miyatani S, Copeland NG, Gilbert DJ, Jenkins NA, Takeichi M. Mouse alpha N-catenin: two isoforms, specific expression in the nervous system, and chromosomal localization of the gene. Dev Biol 1994; 163:75–85.[CrossRef][Medline]
  59. Peifer M, Berg S, Reynolds AB. A repeating amino acid motif shared by proteins with diverse cellular roles. Cell 1994; 76:789–791.[CrossRef][Medline]
  60. Kapprell H-P, Cowin P, Franke WW. Biochemical characterization of the soluble form of the junctional plaque protein, plakoglobin, from different cell types. Eur J Biochem 1987; 166:505–517.[Medline]
  61. Duden R, Franke WW. Organization of the desmosomal plaque proteins in cells growing at low calcium concentration. J Cell Biol 1988; 107:1049–1063.[Abstract/Free Full Text]
  62. Watabe M, Nagafuchi A, Tsukita S, Takeichi M. Induction of polarized cell-cell association and retardation of growth by activation of the E-cadherin-catenin adhesion system in a dispersed carcinoma line. J Cell Biol 1994; 127:247–256.[Abstract/Free Full Text]
  63. Rimm DL, Koslov ER, Kebriaei P, Cianci CD, Morrow JS. Alpha1(E)-catenin is an actin-binding and -bundling protein mediating the attachment of F-actin to the membrane adhesion complex. Proc Natl Acad Sci USA 1995; 92:8813–8817.[Abstract/Free Full Text]
  64. Watabe-Uchida M, Uchida N, Imamura Y, Nagafuchi A, Fujimoto K, Uemura T, Vermeulen S, van Roy F, Adamson ED, Takeichi M. {alpha}-Catenin-vinculin interaction functions to organize the apical junctional complex in epithelial cells. J Cell Biol 1998; 142:847–857.[Abstract/Free Full Text]
  65. Hirano S, Kimoto N, Shimoyama Y, Hirohashi S, Takeichi M. Identification of a neural {alpha}-catenin as a key regulator of cadherin function and multicellular organization. Cell 1992; 70:293–301.[CrossRef][Medline]
  66. Claverie JM, Hardelin P, Legouis R, Levilliers J, Bougueleret L, Mattei MG, Petit C. Characterization and chromosomal assignment of human cDNA encoding a protein related to the murine 102-kDa cadherin-associated protein (alpha-catenin). Genomics 1993; 15:13–20.[CrossRef][Medline]
  67. Butz S, Stappert J, Wessig H, Kemler R. Plakoglobin and ß-catenin: distinct but closely related. Science 1992; 257:1142–1144.[Free Full Text]
  68. Dumont JN, Brummett AR. Oogenesis in Xenopus laevis (Daudin). V. Relationships between developing oocytes and their investing follicular tissues. J Morphol 1978; 155:73–98.[CrossRef][Medline]
  69. Wallace RA. Interactions between somatic cells and the growing oocyte of Xenopus laevis. In: McLaren M, Wylie CC (eds.), Current Problems in Germ Cells. Cambridge: Cambridge University Press; 1983: 285–306.
  70. Gosden R, Krapez J, Briggs D. Growth and development of the mammalian oocyte. Bioessays 1997; 19:875–882.[CrossRef][Medline]
  71. Stebbins-Boaz B, Richter JD. Translational control during early development. Crit Rev Eukaryotic Gene Expr 1997; 7:73–94.[Medline]
  72. Müller AHJ, Gawantka V, Ding X, Hausen P. Maturation induced internalization of ß1-integrin by Xenopus oocytes and formation of the maternal integrin pool. Mech Dev 1993; 42:77–88.[CrossRef][Medline]
  73. Schneider S, Herrenknecht K, Butz S, Kemler R, Hausen P. Catenins in Xenopus embryogenesis and their relation to the cadherin-mediated cell-cell adhesion system. Development 1993; 118:629–640.[Abstract]
  74. Schneider S, Steinbeisser H, Warga RM, Hausen P. ß-Catenin translocation into nuclei demarcates the dorsalizing centers in frog and fish embryos. Mech Dev 1996; 57:191–198.[CrossRef][Medline]
  75. Kofron M, Spagnuolo A, Klymkowsky M, Wylie C, Heasman J. The roles of maternal {alpha}-catenin and plakoglobin in the early Xenopus embryo. Development 1997; 124:1553–1560.[Abstract]
  76. Wahl J, Sacco PA, McGranahan-Sadler TM, Sauppé LM, Wheelock MJ, Johnson KR. Plakoglobin domains that define its association with the desmosomal cadherins and the classical cadherins: identification of unique and shared domains. J Cell Sci 1996; 109:1143–1154.[Abstract]



This article has been cited by other articles:


Home page
 Biol. Reprod.Home page
D. Baron, R. Houlgatte, A. Fostier, and Y. Guiguen
Large-Scale Temporal Gene Expression Profiling During Gonadal Differentiation and Early Gametogenesis in Rainbow Trout
Biol Reprod, November 1, 2005; 73(5): 959 - 966.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow My Folders
Right arrow Download to citation manager
Right arrow reprints & permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Cerdà, J.
Right arrow Articles by Franke, W. W.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Cerdà, J.
Right arrow Articles by Franke, W. W.
Agricola
Right arrow Articles by Cerdà, J.
Right arrow Articles by Franke, W. W.

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS