Cloning and characterization of a second AP-2 ... - Development

Development 121, 2779-2788 (1995) Printed in Great Britain © The Company of Biologists Limited 1995

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Cloning and characterization of a second AP-2 transcription factor: AP-2β Markus Moser1, Axel Imhof1, Armin Pscherer1, Reinhard Bauer1, Werner Amselgruber2, Fred Sinowatz2, Ferdinand Hofstädter1, Roland Schüle3 and Reinhard Buettner1,* 1Institute for Pathology, University of Regensburg Medical School, D-93042 Regensburg, Germany 2Institute of Veterinary Anatomy II, Histology and Embryology, University of Munich, D-80539 Munich, 3Tumor Biology Center, Breisacherstr 117, D-79106 Freiburg, Germany

Germany

*Author for correspondence

SUMMARY AP-2 has been characterized previously as a unique 52×103 Mr transcription activator encoded by a single gene that is expressed in a restricted pattern during embryonic morphogenesis of the peripheral nervous system, face, skin and nephric tissues. Here we report the isolation of genomic and cDNA clones encoding for a second AP-2 related transcription factor, designated AP-2β. AP-2β binds specifically to a series of well-characterized AP-2 binding sites, consensus to the sequence G/CCCN3GGC, and transactivates transcription from a reporter plasmid under the control of an AP-2-dependent promoter. A C-terminal domain known to mediate homodimerization of the previously cloned AP-2α transcription activator is highly conserved and sufficient to mediate interaction between the two proteins. Northern blot and in situ hybridizations revealed that the two genes are expressed in murine embryos between days 9.5 and 19.5 p.c. Coexpression of

both mRNAs was detected in many tissues at day 13.5 and 15.5 of embryogenesis but some regions of the developing brain and face including the primordium of midbrain and the facial mesenchyme differed in their expression pattern of AP-2 genes. AP-2α and AP-2β signals in the central and peripheral nervous system overlapped with regions of developing sensory neurons. In adult tissues AP-2α expression was found mainly in the skin, eye and prostate and AP-2β expression in the kidney. In summary, our analyses of embryonic and adult mice demonstrate that two different AP-2 transcription factors are specifically expressed during differentiation of many neural, epidermal and urogenital tissues.

INTRODUCTION

neural crest-derived cell lineages and skin. Detailed northern blot and in situ hybridization studies of murine embryos revealed a temporally and spatially restricted pattern of AP-2 expression in tissues including the spinal cord, the hindbrain and midbrain junction, cranial and spinal ganglia and the facial mesenchyme. AP-2 expression was also observed in limb bud mesenchyme, in meso-metanephric mesenchyme and in surface ectoderm (Mitchell et al., 1991). A role of AP-2 in epidermal differentiation was further substantiated by studies of embryonic and adult skin in Xenopus which demonstrated that the keratin gene-regulatory factor KTF-1 is identical with or closely related to AP-2 (Snape et al., 1991; Winning et al., 1991). This factor, also known as KER1 from human keratinocytes, is involved in regulation of keratin gene promoters during epidermal differentiation (Leask et al., 1991). A recent analysis of murine embryonic skin development revealed that AP-2 mRNA is expressed in a pattern similar to, but preceding that of basal keratin mRNAs (Byrne et al., 1994). This expression pattern in combination with the observation that AP-2 has a strong inductive effect on basal keratin expression in a cellular environment that does not normally possess AP-2 activity has led to the conclusion that AP-2 is a key regulator of skin differentiation.

Transcription factors have been identified as key nuclear regulators of gene expression programmes being targeted by external signals during vertebrate development. Morphogenesis requires spatial information and induction of cell typespecific gene expression both mediated by interaction of transcription factors with response elements in gene promoters. Frequently, such master regulatory factors belong to families of several closely related transcription factors with similar properties in vitro but different spatial or temporal expression patterns in vivo. This concept has been substantiated in vivo by transgenic and gene-targeted mutation experiments involving homeobox genes (Balling et al., 1989; Kessel et al., 1990) pax genes (Gruss and Walther, 1992; Kessel and Gruss, 1990), myogenic transcription factors (Olson and Klein, 1994; Rudnicki et al., 1994; Sassoon, 1992) and retinoic acid receptors (Lufkin et al., 1993; Lohnes et al., 1993) that resulted in characteristic malformations or loss of specific cell types. Transcription factor AP-2 was purified and cloned from HeLa cells (Mitchell et al., 1987; Williams et al., 1988) and subsequently shown to have important functions in retinoidcontrolled morphogenesis and differentiation, particularly of

Key words: AP-2, transcription factor, mouse embryogenesis, cell differentiation, gene expression

2780 M. Moser and others A causal role for AP-2 in mediating retinoid-induced cell differentiation was established by analyses of teratocarcinoma cells. Treatment of both N-Tera-2 and PA-1 cells with retinoic acid results in transient induction of AP-2 mRNA levels and AP2 transcriptional activation with a maximum between 24 and 48 hours (Lüscher et al., 1989; Kannan et al., 1994). We have recently identified a differentially spliced short form of AP-2, designated AP-2B, that is a negative regulator of transcriptional activation by the previously known full-length AP-2 protein, designated AP-2A (Buettner et al., 1993). Transfection of AP2B into PA-1 cells inhibited the induction of endogenous AP-2 function by retinoic acid and resulted in a retinoid-resistant cell phenotype incapable of differentiation. We further observed that both acquisition of tumorigenicity and inhibition of cellular differentiation by an activated N-ras oncogene caused suppression of AP-2 transcriptional activity (Kannan et al., 1994). Besides activating specific patterns of gene expression during retinoid-induced cell differentiation we have shown recently that a second crucial function of AP-2 is to block induction of mycdependent genes by forming a ternary protein complex with myc/max heterodimers (Gaubatz et al., 1995). Thus, AP-2 is a key molecular switch regulating both cell proliferation and differentiation-specific gene expression. In contrast to many other families of developmentally important transcription factors AP-2 was believed to represent a unique factor encoded by a single copy gene located on chromosome 6p22.3pter (Gaynor et al., 1991). A helix-span-helix motif and a basically charged domain located in the C-terminal half of the protein were identified to mediate both homodimerization and sequence-specific DNA binding (Williams and Tjian, 1991a,b) and were shown to be necessary and sufficient for interaction with c-myc (Gaubatz et al., 1995). Transcriptional activation depends on a proline and glutamine-rich motif located on a single exon within the N terminus. Functional AP-2 binding sites with a palindromic core sequence GCCN3GGC have been identified in many gene promoters including neural, epidermal and viral enhancers. However, a number of AP-2 sites that are specifically footprinted by purified AP-2 have been found to differ significantly from this consensus sequence (Imagawa et al., 1987) raising the possibility that some AP-2 sites may be targeted by several enhancer binding proteins. Composite elements of an AP-2 site and an E-box which integrate signals from different signal transduction pathways have been clearly identified in gene promoters including the human and rat prothymosin-α and ornithin decarboxylase genes and the human and mouse spermidine synthetase, N-ras, T-cell receptor α and int-2 genes (Gaubatz et al., 1995). Elements in the SV40 enhancer and the human growth hormone gene have been found to interact with AP-2 or AP-3 and AP-2 or NF-1 in a mutually exclusive manner (Mercurio and Karin, 1989; Courtois et al., 1990). We have recently described an AP-2 binding site in the human AP2α gene promoter which confers autoregulation to the AP-2 gene and is a target for several distinct proteins (Bauer et al., 1994 and unpublished observations). When we cloned the human AP-2α gene locus we isolated genomic and subsequently complementary DNA clones that crosshybridized to the full-length human AP-2A cDNA. Here we report that these clones encode for a novel AP-2 related transcription factor, designated AP-2β, with a specific expression pattern in murine embryonic and adult tissues.

Thus, multiple AP-2 proteins translated from two different genes and derived by alternative splicing cooperate with other transcritpion factors to form combinatorial networks for regulation of cell proliferation and differentiation. For a clear terminology we refer to the two AP-2 genomic loci as AP-2α and AP-2β and to the two splice variants derived from the AP-2α gene as AP-2αA and AP-2αB. MATERIALS AND METHODS Screening of genomic and cDNA libraries Handling of lambda phages and plaque lifting was performed using standard procedures (Sambrook et al., 1989). A partial human AP2αA cDNA clone spanning nucleotides −45 to −363 with respect to the ATG translation start codon was used to screen 5×105 recombinant phages of a human genomic library in the vector lambda FixII (Stratagene, Heidelberg, Germany) representing approximately two genomes. Hybridizations were performed at intermediate stringency with the last wash in 0.5× SSC, 0.1% SDS at 55°C for 30 minutes. The human AP-2αA cDNA probe was further used to screen 3×105 recombinant phages of a commercially available murine day 13.5 embryo cDNA library cloned in the phage lambda Lexlox (AMS Biotechnology, Lugano, Switzerland). The plasmids inserted into the phages were recovered by CRE-mediated excision according to the manufacturer’s instructions and the inserts fully sequenced on both strands. Gel mobility-shift assays Both AP-2α and β proteins were modified by inserting N-terminal hexahistidine cassettes and ligated into the lacZ promoter-based expression plasmid pSK40 (Skerra and Plückthun, 1989). Conditions for expression and purification of recombinant AP-2 in E.coli using ion metal chromatography have been described in detail recently (Buettner et al., 1993; VanDyke et al., 1992). 5 ng of purified recombinant AP-2 protein was used per gel-shift assay. The following oligonucleotide sequences were hybridized to their respective antisense oligonucleotides and 32P-labeled at their 5′ termini. hMTIIA −180 site: 5′-AGGAACTGACCGCCGCGGCCCGTGTGCAGAG-3′, hMTIIA −210 site: 5′-AGGCCGAGGCGTCCCCAGGCGCAAGTGG-3′, hMTIIA −180 mutated: 5′-AGGAACTGACCGACCGCTGCCCGTGTGCAGAG-3′, hMTIIA −180 +1: 5′-AGGAACTGACCGCCGACGGCCCGTGTGCAGAG-3′, hMTIIA −180 +2: 5′-AGGAACTGACCGCCCGAACG-GCCCGTGTGCAGAG-3′, SV40: 5′-AGTAGGG GTGGAA-AGTCCCCAGGCTCCCCAG-3′, hc-myc: 5′-AGCGGCTGAGGACCCCCGAGCTGTGCTGCT-3′. Conditions for gel shiftreactions have been described previously (Buettner et al., 1993) and for competition experiments a 50 fold molar excess of unlabeled binding site was added. The purity of recombinant proteins was approximately 95% based on silver stained SDS polyacrylamide gels and the integrity of proteins was checked by western blotting the gels and immunostaining using a commercially available AP-2 antiserum (Santa Cruz Biotechnology, Santa Cruz, California). In vitro protein binding assay A fusion protein of AP-2αA and glutathione S-transferase (GST) was expressed in Escherichia coli (Smith and Johnson, 1988). 50 µl of glutathione-Sepharose beads (Pharmacia) were loaded with 20 µg of GST or GST/AP-2A fusion protein and then incubated with 10 µl of 35Smethionine labeled AP-2αA or AP-2β proteins translated in vitro using reticulocyte lysates (Promega, Madison, Wisconsin). Beads were washed five times with a buffer containing 25 mM Hepes, pH 7.9, 25 mM NaCl, 5 mM MgCl2, 0.5 mM DTT and 0.5% NP-40. Bound proteins were eluted from the beads in washing buffer sup-

Molecular cloning of AP-2β 2781 plemented with 5 mM glutathione and separated on a 10% SDS polyacrylamide gel. As a control the AP-2αA N terminus (amino acids 1165) was fused to GST and shown not to retain 35S-labeled AP-2α or AP-2β proteins (data not shown). Transient transfections and CAT assays To measure AP-2 transactivation a pBLCAT2-based chloramphenicol-acetyl-transferase reporter containing the trimerized hMTIIA-180 site in front of a minimal TK promoter (Buettner et al., 1993) was cotransfected with AP-2αA and AP-2β expression plasmids into F9 cells. Transient transfections were performed using standard calcium phosphate precipitations (Graham and van der Eb, 1973). F9 cells were cultured in 1:1 Ham’s F12/DMEM supplemented with 10% fetal calf serum. CAT assays were performed with amounts of protein extracts equalized by cotransfecting an LTR-lacZ expression plasmid (Gorman et al., 1982). For expression of AP-2αA and AP-2β, fully encoding murine cDNAs were ligated into the cytomegalovirus-based plasmid pCMX (Umesono et al., 1991). RNA isolation, northern blots, probes and RT-PCR Whole embryos were minced in liquid nitrogen and total cellular RNA was extracted (Sambrook et al., 1989). Oligo d(T) cellulose chromatography was performed twice using commercially available spin columns (Pharmacia). 8 µg of twice poly(A)+-selected RNA was loaded per lane on 1.2% agarose formaldehyde gels and blotted onto nylon membranes according to standard protocols (Sambrook et al., 1989). The probes used for specific hybridizations of AP-2α and AP2β mRNA are a BamHI/BglII fragment spanning nucleotides 365-789 of the human AP-2αA cDNA and a SmaI fragment spanning nucleotides 343-668 of the murine AP-2β cDNA. For northern probes these fragments were excised from a gel and labeled with [32P]dCTP to a specific activity of 2×109 cpm/µg using a T7 random primer kit (BioRad, Richmond, Califormia). For 35S-labeled riboprobes these fragments were cloned into the plasmid pBluescript and transcribed with a specific activity of 3×107 cpm/µg RNA using T7 and T3 RNA polymerase (Stratagene). RT-PCR amplifications were performed as described previously (Buettner et al., 1993). For amplification of AP-2α mRNA, primers 5′-CCC AAG CCA TAG CTC GAG ACT C-3′ and 5′-GAA GTC ACA GAT TTG GGA GGG A-3′ were used, resulting in a 531 bp fragment spanning residues 66-597 of the cDNA. For AP-2β mRNA amplification, primers 5′-ACC AGC AAC GGG ACG GCA CGG-3′ and 5′-TGG CGG AGA CAG CAT TGC TGT TG-3′ were used resulting in a 518 bp fragment spanning residues 61-579 of the cDNA with respect to the ATG protein start codon. For RNA obtained from whole embryos a second nested PCR was performed. Nested primers for AP-2α were 5′-CTG GGC ACT GTA GGT CAA TCT C-3′ and 5′-GGA CGT CCT CGA TGG CGT GAG G-3′ resulting in a 388 bp fragment and primers for AP-2β were 5′-GAA GTG GGC TCA GAA GCC GGC T-3′ and 5′-AAT GAC TGA CTG GTC CAA TAG G-3′ resulting in a 259 bp fragment. As a control S16 ribosomal protein RNA or β-actin mRNA was coamplified (Byrne et al., 1994). For all PCR, including the nested reactions, 25 cycles of 1 minute at 95°C 1 minute at 64°C and 1 minute at 72°C were performed. All PCR products were Southern blotted and hybridized with AP-2α and AP2β specific probes to faciliate quantification. In situ hybridizations At the appropriate time after mating NMRI mice were perfused with 4% paraformaldehyde/0.5% glutaraldehyde in 0.15 M cacodylate buffer, pH 7.4 and the embryos were dissected and postfixed for 2 hours in the same solution. In situ hybridization of paraffin-embedded sections was performed essentially as described by Hogan et al. (1986). Briefly, slides were pretreated with proteinase K (1-10 µg/ml) for 30 minutes at 37°C fixed in 4% paraformaldehyde/PBS (pH 7.0) for 5 minutes, washed twice in H2O and acetylated in acetic anhydride diluted 1:400 in 0.1 M triethanolamine (pH 8) for 10 minutes at room temperature. Finally, slides

were washed twice with H2O, dehydrated in ethanol, air dried and prehybridized for 4 hours at 50°C in 50% formamide, 10% dextran sulfate, 10 mM Tris pH 8, 10 mM sodium phosphate, pH 7, 2× SSC, 5 mM EDTA, pH 8, 150 µg/ml tRNA, 10 mM DTT, 10 mM β-mercaptoethanol. Hybridizations were performed in the same mix supplemented with 5×104 cpm/µl cpm of sense or antisense riboprobe at 50°C overnight. The slides were washed twice at 52°C in 50% formamide/2× SSC for 30 minutes and again twice in 2× SSC for 5 minutes. After RNase A treatment (20 µg/ml) for 30 minutes at 37°C slides were washed again five times in 50% formamide/2× SSC at 65°C for 1 hour or overnight at 55°C, finally rinsed in 2× SSC, dehydrated, coated with Kodak NTB 2 emulsion and exposed for 2 weeks.

RESULTS Isolation of AP-2 related genomic and complementary DNA clones As described recently (Bauer et al., 1994) we used a partial EcoRI/BamHI human AP-2α cDNA probe to screen a human genomic library and we have isolated 4 independent phages covering the entire AP-2α gene locus. In addition two other weakly hybridizing phages had inserts incompatible with the other clones regarding their restriction pattern (Buettner et al., 1994). The full 19 kb insert of one of these crosshybridizing phages was sequenced, revealing 5 short regions homologous to exons 2-6 of the AP-2α gene. All of these five regions were flanked by consensus splice donor and acceptor sites and interrupted by intervening intron sequences with no homology to the AP-2α gene. As these five putative exons encoded an open reading frame we concluded that we had isolated partial genomic clones of a gene distinct from AP-2α. With respect to its high sequence homology to AP-2α and equivalent genomic organization this gene was designated AP-2β. To isolate a fully encoding AP-2β cDNA clone we used the human AP-2α cDNA probe to screen further a murine cDNA library derived from a total embryo at day 13.5 of gestation. Altogether 7 independent cDNA clones were obtained, 4 of which represented murine AP-2α cDNA clones (Moser et al., 1993). Three other phagemids contained cDNA inserts 96% identical with the respective nucleic acid residues of the five putative human AP-2β exons. As the predicted open reading frames in the murine and the human clones exhibited an even higher degree of identity (98%) we concluded that we had isolated the murine cDNA corresponding to the human genomic locus AP-2β. Fig. 1 displays the sequence of the longest AP-2β cDNA clone covering an open reading frame of 448 amino acids flanked by a 5′ untranslated sequence of 213 bp and a 3′ untranslated sequence of 174 bases (EMBL data library accession number X78197). The predicted amino acids of AP-2β and AP2α are indicated underneath the codons in Fig. 1 and residues conserved between the two proteins are linked by a dash. There is an overall identity of 76% and a similarity of 85% based on amino acid level and an identity of 70% based on nucleic acid level between the murine AP-2α and AP-2β sequences. Inspection of the two protein sequences revealed the highest degree of homology (92%) in the region from amino acids 217 to 448 covering the DNA binding and dimerization motifs. Also the region between amino acids 203 and 227 in the AP-2α protein necessary for interaction with the c-myc proto-oncogene is highly conserved in AP-2β (88% homology). In contrast, the two proteins differ significantly between amino acids 21 to 217

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Fig. 1. Nucleotide and deducted amino acid sequence of the murine AP-2β cDNA. The murine AP-2α amino acid sequence (Moser et al., 1993) is indicated underneath the AP-2β sequence and conserved residues are denoted by vertical dashes.

(58% identity), a region that has been identified as harbouring the transactivation domain (Williman and Tjian, 1991a). However, most of the proline and glutamine residues in this area are located at identical positions in AP-2β. A putative cAMP dependent protein kinase recognition site (RRSLSP from aa 244 to 248; Kemp and Pearson, 1990), a short amino-terminal peptide (from aa 1 to 13) and a strongly charged C-terminal peptide (DKEEKHRK from 441 to 448) are motifs well conserved between the AP-2α and AP-2β proteins. Sequence-specific DNA binding, transcriptional activation and interaction between AP-2α and AP-2β To compare the DNA binding properties of AP-2α and AP-2β both proteins were tagged with a hexahistidine cassette at their N termini to allow for binding to zinc-chelate sepharose (VanDyke et al., 1992) and expressed in E. coli. Soluble bacterial extracts were purified by ion metal chromatography resulting in

recombinant AP-2α and AP-2β proteins of approximately 95% purity assessed by silver stained SDS-PAGE (data not shown). Purified AP-2α and AP-2β proteins were then tested by gel mobility-shift assays for their ability to bind to several synthetic AP-2 sites covering DNaseI-footprinted regions in the human metallothionein IIa gene promoter from residue −191 to −164 (hMTIIA −180) and from residue −227 to −201 (hMTIIA −210), in the SV40 promoter from residue 250 to 222 (SV40) and in the human c-myc gene promoter from residue −668 to −640 (hc-myc). We found the classification of these sites according to their relative affinity as determined with purified AP-2 from HeLa cell nuclear extracts (Imagawa et al., 1987) to be highly reproducible with bacterially expressed recombinant AP-2 proteins. Both AP-2α and AP-2β bound strongly to the hMTIIA −180 and SV40 AP-2 recognition sites with similar binding affinities (Fig. 2A) and interacted weakly with the hMTIIA −210 and hc-myc sites. The specificity of

Molecular cloning of AP-2β 2783 1987). As verified by the gel-shift assays shown in Fig. 2A both AP-2α and AP-2β proteins bind strongly to this element. Fully encoding murine AP-2α and AP-2β cDNAs were subcloned into the cytomegalovirus-based expression plasmid pCMXpL1 and cotransfected with the AP-2 reporter into F9 murine teratocarcinoma cells. We observed approximately 15fold transactivation by AP-2α and slightly weaker 12-fold transactivation by AP-2β (fig. 2B). These results demonstrate that both proteins AP-2α and AP-2β transactivate transcription from an AP-2 sensitive promoter in F9 cells. The AP-2α and β gel-shift complexes shown in Fig. 2A differed slightly in their electrophoretic mobility and a mixture of the two proteins resulted in an intermediate complex. This observation prompted us to ask whether the two proteins interact in vitro. Therefore, AP-2α was expressed and purified as a fusion protein with glutathione-S-transferase (Fig. 3A) and immobilized at glutathione-sepharose. The matrix was loaded with in vitro translated [35S]methionine-labeled AP-2α and AP-2β protein, respectively, washed and eluted with free glutathione. As shown in figure 3B both AP-2α and AP-2β were specifically retained from the reticulocyte lysate by the GSTAP-2α fusion protein and did not bind to the columns in the absence of immobilized AP-2α protein. Further controls revealed that a C-terminal fragment spanning amino acids 166437 was sufficient to retain both AP-2α and AP-2β proteins in contrast to the first 165 amino acids that did not interact with any of the two 35S-labeled proteins (data not shown).

Fig. 2. DNA binding and transcriptional activation by AP-2α and AP-2β proteins. (A) Gel-shifts using 5 ng recombinant AP-2α or β proteins or a 1:1 mixture of both proteins. Oligomeric AP-2 binding sites from the SV40 promoter (SV40), the human metallothionein IIA gene (hMTIIA) and the human c-myc gene (hc-myc) are described in detail in Materials and methods. (B) AP-2α and AP-2β activate transcription from an AP-2 sensitive promoter in F9 cells. Shown are CAT assays from cells transfected with 2 µg of the plasmids indicated above. RSV-CAT: rRous sarcoma virus LTR-based CAT expression vector. TK-CAT: minimal thymidine kinase promoterbased CAT vector. AP2/TK-CAT: trimerized AP-2 binding site inserted into TK-CAT. α,β: CMV promoter-based AP-2α and AP-2β expression plasmids.

binding was verified by introducing C to A and G to T mutations into the two half sites of the hMTIIA −180 palindromic core sequence (hMTIIA −180 mut) and by increasing their spacing from N3 to N4 (hMTIIA −180+1 and hMTIIA −180+2). Both types of alterations, point mutations and increased spacing of binding half sites, abolished binding by AP-2α and AP-2β proteins entirely, suggesting that the specific protein to DNA contacts and the spatial requirements of the binding site are very similar. As most sequence differences between AP-2α and AP-2β were found within the N-terminal half of the proteins including the transactivation domain we compared the ability of the two genes to transactivate transcription via binding to an AP-2 recognition site. As an AP-2 sensitive reporter, we subcloned the trimerized hMTIIA −180 site in front of the minimal TK promoter in the plasmid pBLCAT2 (Luckow and Schütz,

Expression of AP-2α and AP-2β during murine embryogenesis To compare the expression patterns of AP-2α and AP-2β in mouse embryos we subcloned partial cDNA fragments that specifically detected AP-2α and AP-2β mRNAs. Thorough control hybridizations confirmed that these probes did not crossreact under conditions of high or moderate stringency (data not shown). 32P-radiolabeled α- and β-specific probes with identical specific activity were then used to hybridize northern blots of total embryonic poly(A)+ RNA prepared from a series of mouse embryos between gestational days 9.5 to 19.5. The results shown in figure 4A revealed that a single AP-2β mRNA, approximately 6 kb in size, was expressed in embryos between day 11.5 and 17.5. In contrast a complex pattern of multiple AP-2α mRNAs including two most abundant 3 and 2 kb transcripts were detected between day 9.5 until birth. Again these northern blots confirmed that the two α- and β-specific cDNA probes did not crosshybridize. We and others have observed earlier multiple transcripts of the AP-2α gene (Buettner et al., 1993; Byrne et al., 1994) which are generated by differential splicing and usage of multiple polyadenylation signals (Bauer et al., 1994). To ensure that the signals obtained by the northern blot shown in Fig. 4A represented AP-2α mRNAs we performed RT-PCR reactions using specific AP-2α and AP-2β primer pairs. We were able to amplify levels of AP2α and AP-2β mRNA fragments in good agreement with the Northern blot signals from day 9.5 until birth (Fig. 4B). We then transcribed sense and antisense 35S-labeled riboprobes from the AP-2α and AP-2β-specific cDNA fragments to determine the tissue-specific expression pattern of the two AP2 genes by in situ hybridization of paraffin-embedded murine embryos. Sections obtained from embryos of days 13.5 and 15.5

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Fig. 3. In vitro interaction of AP-2α and AP-2β proteins. (A) Western blot analysis of bacterially expressed GST-AP-2α fusion protein and his-tagged AP-2α and β proteins. +/− indicate purifications from soluble E. coli lysates in the presence and absence of 1 mM IPTG. (B) Interaction of in vitro translated 35S-labeled AP2α and AP-2β proteins with a GST-AP-2α affinity matrix.

p.c. were analyzed because significant levels of both genes were detected by northern blots at these gestational stages. Both AP-2α and β signals were abundant in cells coating the neural tube in day 13.5 embryos. A few representative slides are shown in Fig. 5 and a summary of serial vertical and transverse sections is presented in Table 1. Expression started anteriorly in a distinct layer of cells located within the roof of the midbrain and extended downwards into the area around the fourth ventricle including the midbrain/cerebellar junction, the primordia of cerebellum, pons and medulla oblongata. Expression continued posteriorly in the spinal cord, the dorsal root ganglia, further in the prevertebral sympathic ganglia and the ganglion nodosum (Fig. 5). Positive cells in the spinal cord were found mainly in the dorsolateral mantle layer of the grey matter where the sensory axons of the dorsal root ganglia enter and are connected to the second sensory neurons (data not shown). It is interesting that the signals were confined strictly to differentiating cells underneath the layer of dividing cells coating the inner surface of the dorsal central canal. Expression outside the central nervous system was observed in the skin, the kidneys and in many areas of the facial mesenchyme (Table 1). Both AP-2α and β signals in the skin were strictly confined to the ectodermally derived epithelium including hair follicles and sharply discontinued at the border with the entodermally derived oral epithelium (see Fig. 5A).

Fig. 4. Northern blot and RT-PCR analyses of AP-2α and AP-2β expression during murine embryogenesis. (A) Northern blots of RNA isolated from whole murine embryos between day 9.5 and 19.5 p.c. Shown are autoradiographs resulting from specific AP-2α and AP-2β probes and below the respective agarose gels to verify loading of RNA. (B) AP-2α and AP-2β RT-PCR amplification of murine embryo RNA between days 9.5 and 19.5. Nested primer pairs were used to amplify specifically AP-2α and AP-2β (see Materials and Methods) and rRNA (S16) coamplified to demonstrate equivalence of mRNA levels.

These results agree well with a recent very thorough study of AP-2 expression in developing skin (Byrne et al., 1994) and show that both genes are coexpressed in skin and hair follicles. Similarly, AP-2 signals in the kidney resulted from coexpression of both genes in the metanephric tubules and vesicles (Table 1 and data not shown). No expression was observed in the undifferentiated mesenchyme still present in the nephric medulla. To a large extent, regions of AP-2α-specific hybridization signals overlapped with the pattern of AP-2β expression (Fig. 5B,C). However several distinct differences in the expression pattern of the two genes were observed in structures of the midbrain and facial mesenchyme. AP-2β-specific signals were observed to be abundant in the dorsal and anteriolateral primordium of midbrain where AP-2α expression was scarce. In contrast AP-2α mRNA was expressed in a distinct region of the anterior midbrain. As in the spinal cord we found that the layer of undifferentiated mitotic cells coating the innermost surface of the neural tube did not express any AP-2 mRNAs (Fig. 5D-G). Expression of AP-2 in the facial mesenchyme was associated with areas of developing sensory structures. Both messages were detected with different intensities around the olfactory epi-

Molecular cloning of AP-2β 2785

Fig. 5. In situ hybridizations of vertical sections obtained from murine embryos at day 13.5 and 15.5. (A) Embryo at stage E13.5 hybridized to AP-2β cRNA. Note signals in the facial skin (sk), tongue, around the nasal cavity (of), the midbrain (mb) and spinal cord. (B) E13.5 embryo hybridized to AP-2β and to AP-2α. (C) cRNA. Note differences of the hybridization pattern in the midbrain in structures around the oral cavity and the ganglion nodosum (gn). bp: branchial pouch; sc, spinal cord; cb, cerebellum; drg, dorsal root ganglion. (D) Sagittal sections through the midbrain and hindbrain junction of E15.5 embryos hybridized to AP-2β and to AP-2α. (E) cRNA. (F,G) Lateral sections through the midbrain and hindbrain of E15.5 embryos hybridized to AP-2β (F) and to AP-2α (G) cRNA. Scale bars, (A-C) 1 mm; (D-G) 0.3 mm.

2786 M. Moser and others thelium (Fig. 5A-C). Especially in embryos at day 15.5 it appeared that AP-2β signals were significantly more abundant than AP-2α signals and that the latter were much more scattered around the nasal cavity (data not shown). Below the surface of the tongue and around the first branchial pouch we observed specifically AP-2β signals but not expression of AP-2α mRNA both on the sagittal and the transverse sections. The skin and whisker pads around the oral orificium which represent structures important for palpation abundantly expressed both mRNAs. As described in detail previously (Byrne et al., 1994) AP-2 expression in the body skin occurred only as discrete patches at that developmental stage. Strong signals were further observed in the cornea and around the eye (Table 1; data not shown). Expression of AP-2α and AP-2β in adult tissues and cell lines Expression of both AP-2 mRNAs decreased significantly after birth and we were not able to measure AP-2 expression in adult tissues by means of northern blot analysis, with the exception of very faint AP-2α-specific signals in skin (data not shown). Therefore, pairs of primers were designed to allow for amplification of AP-2 mRNAs by RT-PCR. As shown in figure 6 these primers amplify specifically AP-2α or AP-2β cDNA fragments when plasmid DNA is used as a template in control reactions. AP-2α expression was detected in a wide variety of cell lines including HeLa cells derived from a cervical carcinoma and HaCat cells representing human keratinocytes. Interestingly, all melanoma cell lines we examined including murine B16 and human Mel Im cells expressed AP-2α mRNA. Further, Cos and CV-1 representing nephric epithelial cells, the urothelial cell line RT-4 and PA-1 human teratocarcinoma cells were positive. AP-2 mRNA levels observed in RT-4 cells were low but reproducible. No signals were obtained from human umbilical vein endothelial cells (HUVEC) and the prostate cancer cell line LNCaP. Besides the expected PCR product of 531 bp, we coamplified a second slightly smaller PCR product that hybridized

intensively with the AP-2α-specific cDNA probe. Subcloning and subsequent sequencing of all AP-2α and β RT-PCR products revealed that the small AP-2α PCR product was derived by aberrant internal priming of the AP-2α sense primer within the AP-2α mRNA and did not represent a differentially spliced mRNA or an aberrantly primed AP-2β product. Consistently, this smaller product was not observed in a nested PCR reaction as shown in Fig. 4B. So far we have not yet detected a cell line expressing significant levels of AP-2β mRNA. When RNA from adult tissues obtained 2 months after birth was used as a template and moderate PCR conditions were applied (25 cycles, one round of amplification) we observed AP-2 expression in the eye, skin, kidney, prostate, thymus, skeletal muscle and very weakly in the brain (cerebrum). These results agree well with previous data obtained by RNase protection (Mitchell et al., 1991) except for spleen that was negative in our study. Interestingly, the ratio of AP-2α and AP2β mRNAs varied significantly as AP-2α was the more abundant message in skin, prostate, thymus and skeletal muscle whereas abundance of AP-2β mRNA was observed in the kidney. The only AP-2 product that was amplified from brain represented very low levels of AP-2β mRNA. Equal amounts of both α- and β-specific products were amplified from the eye but other tissues including spleen, lung, liver, small and large intestine did not express any of the two AP-2 mRNAs. DISCUSSION Molecular cloning of a second AP-2 related transcription factor Previously, AP-2 has been characterized as a unique tran-

Table 1. Summary of in situ hybridizations AP-2β

AP-2α

Neural tissues forebrain midbrain cerebellum hindbrain spinal cord dorsal root ganglia sympathicus

Tissue

− +++ + + ++ + +

− + ++ + ++ + −

Facial mesenchyme olfactory epithelium tongue first branchial pouch eye

++ + ++ +

+ − − +

Peripheral tissues kidney skin liver spleen heart

++ ++ − − −

++ ++ − − −

Hybridization signals of vertical and transverse sections of murine E13.5 and E15.5 were semi-quantified using four categories: negative (−), weakly positive (+), moderately positive (++) and strongly positive (+++).

Fig. 6. Expression of AP-2α and AP-β in cell lines and adult tissues measured by RT-PCR. (A) Cell lines are HeLa (Hela cells), HaCat (human keratinocytes), 6928 (PA-1 human teratocarcinoma cells), HUVEC (human umbilical vein enothelial cells), Cos and CV-1 (simian epithelial kidney cells), RT-4 (human bladder carcinoma cells), LNCaP (human prostate carcinoma cells) and B16 and Mel Im (melanoma cells). (B) Tissues are indicated above. To control equivalence of mRNA S16 RNA was coamplified.

Molecular cloning of AP-2β 2787 scription factor encoded by a single gene (Williams et al., 1988). No apparent homology to any DNA binding protein other than faint structural similarities to the proline-rich transactivation domains of NF-1 (Mermod et al., 1989) and OTF-2 (Gerster et al., 1990) have been described. The first evidence for a gene family of AP-2 related transcription factors was obtainted by a study of AP-2 expression in Xenopus embryos (Winning et al., 1991). Three different AP-2 mRNA transcripts were detected on northern blots that were regulated differentially during the middle and later stages of embryogenesis. These results prompted the authors to examine genomic DNA on Southern blots, revealing that between 2 and 4 different AP2 related genes exist in Xenopus. Now we report the isolation of a second AP-2 related transcription factor, AP-2β, from a murine embryo cDNA library. A comparison of the partially available intron-exon organization with the recently mapped complete AP-2α gene structure strongly suggests that the two genes originated from a gene duplication of a common ancestor. Currently we are isolating and sequencing the 5′ flanking region of the AP-2β gene and it will be interesting to determine whether cis-regulatory elements that were identified in the AP-2α gene promoter including an autoregulatory AP-2 site (Bauer et al., 1994) are also present in the AP-2β promoter. Our analyses of the AP-2β protein have revealed that all known functional AP-2α protein domains, i.e. dimerization, DNA-binding and transactivation motifs, are conserved. The results from the CAT assays demonstrate that this is also true for the least conserved domain, the proline and glutamine-rich transactivation motif. It has been shown both for P-Q-rich and for acidic-blob-type transactivators that overall structural features are more important than individual amino acids (Ruden et al. 1991; Pei and Shih 1991; Mitchell and Tjian 1989) allowing more non-conserved amino acid exchanges in that protein region. While the precise structural requirements of P-Q-rich transactivators are still unclear, almost all P and Q residues of AP-2α and AP-2β are conserved and therefore likely to be functionally relevant. It was noted by Winning et al. that the Xenopus AP-2 protein lacks a short amino-terminal peptide sequence when compared to the human protein. Interestingly, this sequence encoded by the small first exon is well conserved between AP-2α and AP-2β and also between the murine and the human AP-2α genes. It therefore appears that during evolution a novel exon with an unknown function was added after the separation of amphibia. Expression of different AP-2 mRNAs during embryogenesis The pattern of AP-2α and AP-2β mRNAs detected in mouse embryos at day 13.5 is in good agreement with the AP-2 tissue distribution described previously (Mitchell et al., 1991; Byrne et al., 1994). These studies indicated that AP-2 plays a major role in neural tube formation, skin maturation and urogenital development. However, the probes that were used in earlier experiments were not able to discriminate between the two mRNAs. In order to determine specifically tissue-specific patterns of AP-2α and AP-2β mRNA expression we have therefore designed two partial cDNA probes that were shown by control hybridizations to react specifically with the respective genes. Since a number of different AP-2α transcripts derived by alternative splicing and usage of different

polyadenylation signals have been described (Buettner et al., 1993; Bauer et al., 1994) we did not use 3′ untranslated regions from the cDNA as probes but rather chose sequences from the second to the fourth coding exons. We are not aware of any AP-2 transcript that will not be detected by these probes and we have ruled out, by careful control hybridizations, that these probes crossreact. Further, this region differed enough to design specific primer pairs for RT-PCR reactions. The specificity of these reactions was controlled by cloning and sequencing the amplified products. Even though colocalization of AP-2α and AP-2β signals was detected in many tissues we observed distinct patterns of expression in developing facial structures and in the midbrain. AP-2 mRNA expression in the midbrain has not been described in the previous studies (Mitchell et al., 1991) a difference that is most likely due to the later stage of development that we examined. Consistently, we even found an expanded region of intense AP-2β-specific signals in the midbrain at day 15.5 (Fig. 5D-G). Inversely, AP-2α appears to be the predominantly expressed gene in the cerebellum, but there is also significant coexpression of AP-2β. There are two common features to AP-2 expression in the CNS. Firstly, all signals were confined to differentiating neuroblasts located underneath undifferentiated mitotic cells coating the central canal. However neuroblast differentiation per se does not appear to be dependent on AP-2 since cells in the anterior CNS did not express any AP-2 messages. Therefore, it is possible that AP-2 genes specify certain neural cell differentiation fates. Secondly, the pattern of AP-2 signals suggests that AP-2 plays a role in the development of cells important for sensory perception. The signals in the dorsal root ganglia, the dorsolateral grey matter and the medulla oblongata overlap with the location of the somata of sensory neurons. This observation is further supported by the pattern of signals in the facial mesenchyme. The signals around the olfactory epithelium, in the tongue, around the oral cavity, in the whisker pads, the eye and the first branchial pouch were detected in areas of developing neuroepithelial sensory structures. Therefore, we propose that the expression of AP-2 in neuroepithelial derivatives is associated with differentiating cells involved in sensory perception. The two other regions of significant AP-2 expression were skin and urogenital tissues both in embryos and adult animals. Consistently, we were also able to amplify AP-2 mRNAs from cell lines representing skin, kidney and urinary bladder epithelial cells. Importantly AP-2 expression was not only detected in keratinocytes but also in melanocytic cell lines, indicating that AP-2 plays other roles in skin differentiation beyond regulation of keratinization. In summary, our data demonstrate the presence of two different AP-2 transcription factors, albeit at different ratios, in neuroepithelial, epidermal and urogenital tissues. If the two proteins do not differ in their molecular function, why are two genes needed? It has been realized recently that besides activating transcription a second important function of AP-2 is to interact with and modulate other transcription factors including the c-myc proto-oncogene (Gaubatz et al., 1995). It is therefore possible that the two genes modulate different key regulatory factors during cell differentiation. Alternatively, duplication of the AP-2 gene locus during evolution has resulted in two different gene promoters, allowing more flexibility in gene

2788 M. Moser and others expression regulation and signal response. Gene-targeted knockout experiments are now under way for both AP-2α and AP-2β genes and this will determine whether the two genes have different functions in cell differentiation or morphogenesis. We thank Michael Tainsky (Houston) and Harald Schnürch (Martinsried) for many discussions and help with the photography. Silvia Seegers, Monika Kerscher and Anke Buettner provided excellent technical assistance and help with the manuscript. A. I. is a predoctoral fellow of the Graduiertenkolleg Therapieforschung: Onkologie (Regensburg). This work was supported by grants from the DFG (Bu 672/2-2) and the Wilhelm Sander-Stiftung to R. B.

REFERENCES Balling, R., Mutter, G., Gruss, P. and Kessel, M. (1989). Craniofacial abnormalities induced by ectopic expression of the homeobox gene Hox-1.1 in transgenic mice. Cell 58, 337-347. Bauer, R., Pscherer, A., Imhof, A., Moser, M. Kopp, H., Seegers, S., Kerscher, M., Tainsky, M.A., Hofstaedter, F. and Buettner, R. (1994). The genomic structure of the human AP-2 gene. Nucl. Acids Res. 22, 1413-1420. Buettner, R., Kannan, P., Imhof, A., Bauer, R., Yim, S. O., Glockshuber, R., VanDyke, M. W. and Tainsky, M. A. (1993). An alternatively spliced mRNA from the AP-2 gene encodes a negative regulator of transcriptional activation by AP-2. Mol. Cell. Biol. 13, 4174-4185. Buettner, R., Moser, M., Pscherer, A., Imhof, A., Bauer, R. and Hofstaedter, F. (1994). Molekulare Klonierung eines neuen AP-2 Transkriptionsfaktors, AP-2β, und seine Funktion in der Zelldifferenzierung. Verh. Dtsch. Ges. Path. 78, 38-42. Byrne, C., Tainsky, M. A. and Fuchs, E. (1994). Programming gene expression in developing epidermis. Development 120, 2369-2383. Courtois, S. J., Lafontaine, D. A., Lemaigre, F. P., Durviaux, S. M. and Rousseau, G. G. (1990). Nuclear factor-I and activator protein-2 bind in a mutually exclusive way to the overlapping promoter sequences and transactivate the human growth hormone gene. Nucl. Acids Res. 18, 57-64. Gaubatz, S., Imhof, A., Dosch, R., Werner, O., Mitchell, P., Buettner, R. and Eilers, M. (1995). Transcription factor AP-2 negatively regulates gene activation by myc. EMBO J. 14, 1508-1519. Gaynor, R. B., Muchardt, C., Xia, Y. R., Kliask, I., Mohandas, T., Sparkes, R. S. and Lusis, A. J. (1991). Localization of the gene for the DNA-binding protein AP-2 to human chromosome 6p22. 3-pter. Genomics 10, 1100-1102. Gerster, T., Balmaceda, C. G. and Roeder, R. G. (1990). The cell typespecific octamer transcription factor OTF-2 has two domains required for the activation of transcription. EMBO J. 9, 1635-1643. Gorman, C. M., Moffat, L. M. and Howard, B. (1982). Recombinant genomes which express chloramphenicol acetyl-transferase in mammalian cells. Mol. Cell. Biol. 2, 1044-1053. Graham, F. L. and van der Eb, A. J. (1973). A new technique for the assay of infectivity of human adenovirus 5 DNA. Virology 52, 456-467. Gruss, P. and Walther, C. (1992). Pax in development. Cell 69, 719-722. Hogan, B., Costantini, F. and Lacy, E. (1986). Manipulation of the mouse embryo. A laboratory manual. Cold Spring Harbor, New York: Cold Spring Harbor Laboratory Press. Imagawa, M., Chiu, R. and Karin, M. (1987). Transcription factor AP-2 mediates induction by two different signal transduction pathways: protein kinase C and cAMP. Cell 51, 251-260. Kannan, P., Buettner, R., Chiao, P. J., Yim, S. O., Sarkiss, M. and Tainsky, M. A. (1994). N-ras oncogene causes AP-2 transcriptional self-interference, which leads to transformation. Genes Dev. 8, 1258-1269. Kemp, B. E. and Pearson, R. B. (1990). Protein kinase recognition sequence motifs. Trends Biochem. Sci. 15, 342-346. Kessel, M., Balling, R. and Gruss, P. (1990). Variations of cervical vertebrae after expression of a Hox-1. 1 transgene in mice. Cell 61, 301-308. Kessel, M. and Gruss, P. (1990). Murine developmental control genes. Science 249, 374-379. Leask, A., Byrne, C. and Fuchs, E. (1991). Transcription factor AP-2 and its role in epidermal-specific gene expression. Proc. Natl. Acad. Sci. USA 88, 7948-7952.

Lohnes, D., Kastner, P., Dierich, A., Mark, M., LeMeur, M. and Chambon, P. (1993). Function of the retinoic acid receptor gamma in the mouse. Cell 73, 643-658. Lufkin, T., Lohnes, D., Mark, M., Dierich, A., Gorry, P., Gaub, M. P., LeMeur, M. and Chambon, P. (1993). High postnatal lethality and testis degeneration in retinoic acid receptor alpha mutant mice. Proc. Natl. Acad. Sci. USA 90, 7225-7229. Lüscher, B., Mitchell, P. J., Williams, T. and Tjian, R. (1989). Regulation of transcription factor AP-2 by the morphogen retinoic acid and by second messengers. Genes Dev. 3, 1507-1517. Mercurio, F. and Karin, M. (1989). Transcription factors AP-3 and AP-2 interact with the SV40 enhancer in a mutually exclusive manner. EMBO J. 8, 1455-1460. Mermod, N., O’Neill, E. A., Kelly T. J. and Tjian, R. (1989). The proline-rich transcriptional activator of CTF/NF-1 is distinct from the replication and DNA binding domain. Cell 58, 741-753. Mitchell, P. J., Wang, C. and Tjian, R. (1987). Positive and negative regulation of transcription in vitro: enhancer-binding protein AP-2 is inhibited by SV40 T antigen. Cell 50, 847-861. Mitchell, P. J. and Tjian, R. (1989). Transcriptional regulation in mammalian cells by sequence-specific DNA binding proteins. Science 245, 371-378. Mitchell, P. J., Timmons, P. M., Hebert, J. M., Rigby, P. W. and Tjian, R. (1991). Transcription factor AP-2 is expressed in neural crest lineages during mouse embryogenesis. Genes. Dev. 5, 105-119. Moser, M., Pscherer, A., Bauer, R., Imhof, A., Seegers, S., Kerscher, M. and Buettner, R. (1993). The complete cDNA sequence of the murine AP-2 transcription factor. Nucl. Acids Res. 21, 4844. Olson, E. N. and Klein, W. H. (1994). bHLH factors in muscle development: dead lines and commitments, what to leave in and what to leave out. Genes Dev. 8, 1-8. Pei, D. Q. and Shih, C. H. (1991). An attenuator domain is sandwiched by two distinct transactivation domains in the transcription factor C/EBP. Mol. Cell. Biol. 11, 1480-1487. Ruden, D. M., Ma, J., Li, Y. and Ptashne, M. (1991). Generating yeast transcription activators containing no yeast protein sequences. Nature 350, 250-252. Rudnicki, M. A., Schnegelsberg, P. N., Stead, R. H., Braun, T., Arnold, H. H. and Jaenisch, R. (1993). MyoD or Myf-5 is required for the formation of skeletal muscle. Cell 75, 1351-1359. Sambrook, J., Fritsch, E. F. and Maniatis, T. (1989). Molecular Cloning: A Laboratory Manual, 2nd ed. Cold Spring Harbor, New York: Cold Spring Harbor Laboratory Press. Sassoon, D. A. (1992). Myogenic regulatory factors: dissection of their role and regulation during vertebrate embryogenesis. Dev. Biol. 156, 11-23. Skerra, A. and Plückthun, A. (1989). Assembly of functional immunoglobulin Fv fragment in Escherichia coli. Science 240, 1038-1041. Smith, D. B. and Johnson, K. S. (1988). Single step purification of polypeptides expressed in E. coli as fusion proteins with glutathione-Stransferase. Gene 67, 31-40. Snape, A. M., Winning, R. S. and Sargent, T. D. (1991). Transcription factor AP-2 is tissue-specific in Xenopus and is closely related or identical to keratin transcription factor 1 (KTF-1). Development 113, 283-293. Umesono, K., Murakami, K. K., Thompson, C. C. and Evans, R. M. (1991). Direct repeats as selective response elements for the thyroid hormone, retinoic acid, and vitamin D3 receptors. Cell 65, 1255-1266. VanDyke, M. W., Sirito, M. and Sawadogo, M. (1992). Single-step purification of bacterially expressed polypeptides containing an oligohistidine domain. Gene 111, 99-104. Williams, T., Admon, A., Lüscher, B. and Tjian, R. (1988). Cloning and expression of AP-2, a cell-type-specific transcription factor that activates inducible enhancer elements. Genes Dev. 2, 1557-1569. Williams, T. and Tjian, R. (1991a). Analysis of the DNA-binding and activation properties of the human transcription factor AP-2. Genes Dev. 5, 670-682. Williams, T. and Tjian, R. (1991b). Characterization of a dimerization motif in AP-2 and its function in heterologous DNA-binding proteins. Science 251, 1067-1071. Winning, R. S., Shea, L. J., Marcus, S. J. and Sargent, T. D. (1991). Developmental regulation of transcription factor AP-2 during Xenopus laevis embryogenesis. Nucl. Acids Res. 19, 3709-3714. (Accepted 12 May 1995)