TFAP2A is required for normal expression of melanocyte differentiation genes

Zebrafish homozygous for a strong loss-of-function mutation in tfap2a (i.e., lockjaw allele, hereafter tfap2a^-/- mutants) lack detectable anti-TFAP2A immunoreactivity [28,47] and exhibit approximately one-third reduction of embryonic melanocytes, impaired melanocyte migration, and delayed melanization relative to wildtype and tfap2a^+/- siblings (Fig 1A and 1B) [27,30,48]. In order to better characterize the melanization phenotype, we compared tfap2a^-/- mutants to tfap2a^+/- siblings over a ten-hour period, starting with the first emergence of melanocytes around 28 hours post fertilization (hpf) (S1 Fig). While melanocytes are initially pale in both genotypes, individual melanocytes in the tfap2a^+/- siblings pigmented more quickly than individual melanocytes in tfap2a^-/- mutants. This supports earlier evidence that, in addition to reduced numbers and migration, melanocytes in tfap2a^-/- mutants have defects in differentiation [30].

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Fig 1. Melanocyte differentiation genes are sensitive to the loss of TFAP2A.

(A, B) Lateral images of tfap2a^+/- and tfap2a^-/- mutant zebrafish at 36 hpf. tfap2a^-/- mutants have fewer melanocytes and delayed pigmentation relative to their wildtype or tfap2a^+/- siblings. (C) Fold changes calculated from microarray expression levels for 20 zebrafish melanocyte genes. Top: 11 genes with expression levels between 0.2–0.55-fold in tfap2a^-/- mutants compared to WT siblings. Bottom: 9 genes with expression levels reduced no further than the number of melanocytes in tfap2a^-/- mutants. (D) qRT-PCR validation of zebrafish microarray results for 13 melanocyte genes (Mann-Whitney U, *p<0.05, **p<0.01). (E) qRT-PCR validation of microarray results for 12 melanocyte genes in mouse melan-a cells transfected with control siRNA or each of two siRNAs targeting Tfap2a (siA, si5). Significance evaluated only when both siA and si5 altered gene expression in the same direction relative to siCtrl (one-way ANOVA with Bonferroni correction, ****p<0.0001).

https://doi.org/10.1371/journal.pgen.1006636.g001

To extend previous analyses of gene expression in the melanocytes of tfap2a^-/- mutants [30,34,48,49], we generated expression profiles of tfap2a^-/- mutant zebrafish and their wildtype siblings at 36 hpf. Prior to harvesting RNA, we decapitated animals to eliminate the retinal pigmented epithelium, which appears to be normally pigmented in tfap2a^-/- mutants (Fig 1A and 1B). Generating cDNA and probing microarrays revealed that 2,337 unique Ensembl transcripts (corresponding to 2,324 genes) are differentially expressed in the trunks of tfap2a^-/- mutants versus siblings (FDR p<0.05). Of these, the expression levels of 124 transcripts in tfap2a^-/- mutants are decreased to ≤0.7-fold of wildtype levels and 358 transcripts are increased to ≥1.25-fold (expression profile in S1 Table). We referred to zebrafish gene expression patterns at an online database (ZFIN) to identify 19 genes annotated as “melanoblast,” “melanocyte,” or “pigment cell” [50]. An additional gene, slc24a4a, is annotated as “neural crest,” but is expressed in a pattern resembling that of dct [51]. Most of these 20 genes encode proteins that have known roles in melanocyte differentiation (Fig 1C, S2 Table). In tfap2a^-/- mutants, 11 of these genes were expressed between 0.2- to 0.55-fold of wildtype levels, a much greater fraction of melanocyte genes than expected by chance (hypergeometric test, p<0.0001), and were therefore considered Tfap2a-dependent (Fig 1C). The levels of several others, including trpm1a, were not significantly changed or were reduced by no more than expected from the one-third decrease in melanocyte cell number, and we considered these to be Tfap2a-independent. qRT-PCR validation of microarray results confirmed that expression of multiple pigmentation genes depends on Tfap2a in zebrafish (Fig 1D).

Because TFAP2A is expressed in multiple tissues, including the skin where melanocytes reside, the Tfap2a-dependent gene expression in zebrafish melanocytes could reflect an indirect requirement for Tfap2a rather than a cell-autonomous one. To clarify this issue, we conducted a microarray expression profile of immortalized mouse melanocytes (melan-a cells) [52] depleted of Tfap2a. We identified two independent siRNAs that reduced expression of Tfap2a to below 0.25-fold of the level in cells transfected with control siRNAs. Comparison of microarray profiles showed that the expression levels of 30 genes were consistently decreased (≤0.7-fold) and the expression levels of 38 genes were consistently increased (≥1.4-fold) in cells transfected with either Tfap2a-targeted siRNA relative to control siRNAs (S3 Table). Among the 30 genes with decreased expression, Mc1r and Slc24a4 are implicated in pigmentation. qRT-PCR verified significant reduction of these two genes, as well as Dct, Mlph, and Pmel, which narrowly missed the threshold cut-off in the array analysis (Fig 1E). Of note, several of the other genes with decreased expression upon knockdown of Tfap2a have been reported in the literature as activated (Aldh1a3, Igf2bp1, Ephb2, Pbk) or downregulated (Qpct, Wfdc1) in melanoma [53–59]. In summary, expression of a subset of melanocyte differentiation genes, including Slc24a4, Mc1r, Mlph, Pmel, and Dct, was TFAP2A-dependent in both mouse melanocytes and zebrafish trunks, while expression of Mitf orthologs was TFAP2A-independent. Certain pigmentation genes, including oca2 and slc45a2, appeared Tfap2a-dependent in zebrafish but not in mouse melanocytes, perhaps reflecting a more thorough depletion of Tfap2a in the former, or potentially distinct homeostatic mechanisms induced by siRNA versus gene mutation. Nonetheless, these results indicate that TFAP2A, whether directly or indirectly, regulates the expression of genes involved in melanocyte differentiation.

TFAP2A occupancy in mouse and human melanocytes

To determine the direct transcriptional targets of TFAP2A in melanocytes, we conducted chromatin immunoprecipitation followed by high-throughput sequencing (ChIP-seq) in mouse melan-a cells (two replicates) and in human primary melanocytes (one replicate). Anti-TFAP2A immunoreactivity is strong and concentrated in the nucleus of human primary melanocytes (S2A Fig) but appears weaker and more diffuse in several melanoma cell lines (S2B–S2E Fig), consistent with a reduction of TFAP2A RNA levels in melanoma [45]. ChIP-seq detected 16,305 TFAP2A-bound loci in mouse melanocytes, and 13,690 TFAP2A-bound loci in human melanocytes (hereafter, TFAP2A peaks). De novo motif analysis [60] of sequences precipitated by the anti-TFAP2A antibody from mouse or human melanocytes revealed that a known TFAP2A binding site (MA0003.2, JASPAR) is strongly enriched and tends to be centrally located within peaks (S3A and S3C Fig). Anti-TFAP2A ChIP followed by quantitative PCR (ChIP-qPCR) confirmed TFAP2A binding enrichment for selected peaks at genes of interest in mouse or human melanocytes, as well as in the M21 melanoma cell line, which has detectable TFAP2A expression (S3B, S3D and S3E Fig).

Published comparisons of ChIP-seq results for a given transcription factor in mouse and human melanocytes have suggested rapid divergence of binding events during evolution [61–63]. Consistent with these studies, we found that only about 11% of TFAP2A peaks in human primary melanocytes coincide with the orthologs of TFAP2A peaks lifted over from mouse (S4 Table), and conversely, about 9% of the TFAP2A peaks identified in mouse melanocytes coincide with peaks lifted over from human (S5 Table). TFAP2A peaks shared between species are enriched near promoters (S6 Table). In contrast to the modest concordance of peaks and orthologous sequences, the concordance of genes associated with TFAP2A peaks in the two species is very high, as shown below.

Relationship between TFAP2A occupancy and defined regulatory elements

In both mouse and human melanocytes, TFAP2A peaks are more likely to be found within genes, including introns, than in intergenic regions (Fig 2A, S4A Fig). Overall, genes with higher expression in melanocytes are enriched for promoter-proximal peaks of TFAP2A (mouse GSE87051, human GSM958174 [64]) (Fig 2B, S4B Fig). To assess patterns of TFAP2A binding at enhancers, we compared the TFAP2A ChIP-seq data from mouse melan-a cells to a published profile of candidate enhancers also in these cells, which were defined by H3K4me1 peaks flanking a p300 peak [65]. Remarkably, 70% (1,752 of 2,489) of enhancer elements marked in this way overlap with a TFAP2A peak (hypergeometric test, p<0.0001). Conversely, about 10% of TFAP2A peaks are fully marked as enhancers, while another 35% are partially marked (i.e., either overlapping p300 or flanked by at least one H3K4me1 peak). Fig 2C illustrates this pattern upstream of the melanocyte differentiation gene Slc45a2, as well as a TFAP2A peak at the promoter (additional examples in S5A–S5F Fig). In agreement with estimates that the median distance between promoters and cis-acting enhancers is 15kb, genes with high expression are enriched for a TFAP2A peak overlapping the active enhancer signature at a distance of 5–50kb from the TSS (Fig 2D) [66]. However, the promoter-proximal TFAP2A peaks are just as likely to possess a TFAP2A binding site as the promoter-distal peaks (about 50% in both cases). This is in contrast to observations from mouse chondrocytes, where promoter-distal SOX9 peaks are more likely to contain a SOX9 binding site than promoter-proximal ones [67]. Altogether, the binding profile of TFAP2A indicates that it acts at both enhancers and promoters of melanocyte genes.

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Fig 2. TFAP2A binds active enhancers and promoters in mouse melanocytes.

(A) Pie chart showing distribution of mouse TFAP2A peaks with respect to genomic features. TSS, transcription start site; TTS, transcription termination site. (B) Distance from TSS to the nearest TFAP2A peak for genes in three expression categories: highest 1000, median 1000, or lowest 1000. Promoter-proximal TFAP2A peaks are enriched at highly expressed genes (RNA-seq on mouse melan-a cells at GSE87051). (C) Examples of active enhancer signatures defined by H3K4me1 peaks flanking a p300 peak [65], and partial enhancer signatures, overlapping TFAP2A peaks upstream of the melanocyte differentiation gene Slc45a2. (D) Distance from TSS to the nearest TFAP2A peaks that overlap the active enhancer signature for genes in three expression categories: highest 1000, median 1000, or lowest 1000. (E) Overlap of TFAP2A ChIP-seq peaks in mouse melanocytes with published TFAP2A ChIP-seq peaks in mouse kidney and epididymis cells [68]. 34% of melanocyte peaks were shared with one or both of the other cell types. (F) MEME-ChIP analysis of unique peaks from each cell type. Melanocyte-unique peaks are significantly enriched for SOX10 and M-Box MITF binding motifs, while kidney-unique and epididymis-unique peaks are not. *All motifs shown are a result of de novo MEME-ChIP enrichment analysis except the M-Box, which we specifically searched using the Analysis of Motif Enrichment (AME) tool [109].

https://doi.org/10.1371/journal.pgen.1006636.g002

TFAP2A-bound chromatin elements are enriched for distinct binding motifs in different cell types

Because TFAP2A is widely expressed, the extent to which TFAP2A peaks would be melanocyte-specific was unclear. We found that about 15% of TFAP2A peaks in mouse melanocytes overlap those reported in either primary mouse kidney or primary mouse epididymis cells (but not both), and 19% are shared by all three cell types [68] (Fig 2E). MEME-ChIP analysis [69] showed that melanocyte-unique TFAP2A peaks are enriched for the binding motifs of transcription factors active in melanocytes, including SOX10, FOS/JUN, TEAD, and the M-box binding site for MITF [70] (Fig 2F). These transcription factors are also highly enriched in candidate melanocyte enhancers [65]. Kidney-unique and epididymis-unique peaks showed less significant or no enrichment for these binding motifs, although the E-box, recognized by MITF and many other bHLHZip transcription factors, was enriched in all three cell types. The enrichment of binding sites for melanocyte transcription factors like SOX10 and MITF in melanocyte-unique peaks suggests that TFAP2A binds cell-type specific loci in addition to generic ones; binding to any particular locus is presumably a function of cofactor availability and chromatin accessibility [reviewed in 71].

Many TFAP2A-dependent melanocyte genes are direct targets of TFAP2A

We used Genomic Regions Enrichment of Annotations Tool (GREAT) [72] to identify genes associated with TFAP2A peaks in mouse and human melanocytes, with an assignment rule of basal promoter plus 100kb distal, and found that these genes are enriched for ontology terms relevant to melanocyte differentiation, including “pigmentation”, “melanosome”, and “melanoma” (S6A and S6B Fig). However, TFAP2A can both activate and repress gene expression [73]. To find genes likely to be activated by TFAP2A, we integrated our TFAP2A ChIP-seq with H3K27ac ChIP-seq data from human melanocytes (GSM1127072 [64]) and mouse melanocytes [14], as H3K27ac marks active regulatory elements [reviewed in 74,75]. We found that 55% of TFAP2A peaks in human melanocytes and 58% of TFAP2A peaks in mouse melanocytes either overlap H3K27ac peaks or are flanked by H3K27ac peaks, and we refer to them hereafter as active TFAP2A peaks (human Fig 3A, mouse S6C Fig, S7 Table). Analysis with GREAT revealed that genes associated with active TFAP2A peaks include 15 of the 30 genes with significantly decreased expression in Tfap2a-depleted mouse melanocytes, among them Mc1r and Slc24a4 (S8 Table). Similarly, orthologs of eight out of ten Tfap2a-dependent melanocyte genes in zebrafish tfap2a^-/- mutants (excepting slc24a5 and tyr) are associated with active TFAP2A peaks in mouse or human melanocytes. These genes are candidates to be direct transcriptional targets of TFAP2A, suggesting that the phenotype of delayed melanization in zebrafish tfap2a^-/- mutants can be explained in part by a direct effect on certain melanocyte differentiation effector genes (e.g., dct, mlpha, mc1r, and pmela), and indirect regulation of others (e.g., slc24a5 and tyr).

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Fig 3. TFAP2A peaks are associated with genes involved in pigmentation.

(A-C) Density-based clustering of H3K27ac signal at (A) TFAP2A peaks, (B) MITF peaks, and (C) TFAP2A peaks that overlap MITF peaks in human melanocytes (H3K27ac data from GSM1127072 [64]), MITF peaks from [18]). (D) Overlap between genes associated with active TFAP2A peaks and genes associated with active MITF peaks in human melanocytes. (E) Typical enhancers (gray) and super-enhancers (colored) in human melanocytes that overlap neither TFAP2A nor MITF peaks, TFAP2A peaks only, MITF peaks only, or both TFAP2A and MITF peaks. Labels identify melanocyte genes of interest. (F) Diagram of the TRPM1 -700 bp promoter element depicting the positions of four TFAP2A binding sites (A1–A4) and the previously reported E-box 1 MITF binding site (E1). (G) Luciferase assays in M21 melanoma cells. Deletion of all four TFAP2A binding sites (ΔAP2) significantly reduced reporter activity compared to the intact TRPM1 -700bp element (Student’s t-test, **p = 0.01).

https://doi.org/10.1371/journal.pgen.1006636.g003

TFAP2A and MITF co-occupy regulatory elements associated with pigmentation genes

The presence of TFAP2A peaks at a majority of active melanocyte enhancers, as well as the enrichment of the MITF binding motif in TFAP2A peaks, implies that TFAP2A binds many of the same regulatory elements as MITF in melanocytes. To evaluate overlap between TFAP2A and MITF across the genome, we compared our human TFAP2A ChIP-seq results to a set of 16,572 MITF ChIP-seq peaks also from human primary melanocytes [18]. 5,367 (39%) of TFAP2A peaks are shared with MITF, overlapping by at least one base pair. Integrating MITF and H3K27ac ChIP-seq data yielded 61% of MITF peaks (Fig 3B) and 76% of TFAP2A/MITF shared peaks (Fig 3C) that overlap, or lie between, H3K27ac peaks and are thus considered to be active. Using GREAT, we found that about 77% of the genes associated with active TFAP2A peaks are also associated with active MITF peaks, a highly significant overlap (hypergeometric test, p<0.0001) (Fig 3D). Furthermore, 79% of these genes are associated with active TFAP2A/MITF shared peaks, suggesting that TFAP2A and MITF are co-bound at many, but not all, shared targets. GO term analysis [76,77] revealed that the subset of genes associated with both TFAP2A and MITF peaks are enriched for the terms “melanosome” and “pigment granule” (p = 9.08E-07), as well as “DNA repair” (p = 5.98E-08), “mitotic cell cycle process” (p = 5.59E-09), “regulation of cell proliferation” (p = 2.40E-03), and “regulation of cell differentiation” (p = 2.60E-03) (all p-values Bonferroni corrected, S9 Table). This supports a regulatory role for TFAP2A not only in differentiation, but across other categories of genes proposed to be regulated by MITF in melanocytes and melanoma, as with the MITF rheostat [22].

To assess the overlap between targets of TFAP2A and MITF with respect to pigmentation, we focused on a list of 170 genes that cause coat color phenotypes in mice [78], adding TRPM1 based on its role in the coat color phenotype of appaloosa horses [79–82]. Orthologs of 97 genes on this list are associated with active TFAP2A peaks in human melanocytes and/or active TFAP2A peaks in mouse melanocytes (Table 1, asterisks). Of these, 72 genes are also associated with active MITF peaks (Table 1), 46 being active shared TFAP2A/MITF peaks (Table 1, bold). We then examined overlap of MITF and TFAP2A binding at clusters of closely spaced enhancers, sometimes called stretch or super-enhancers (SEs) [83], that are linked to cell type-specific gene expression [84,85]. Following published methods, we used H3K27ac data to identify 652 SEs in human primary melanocytes [85] (S7 Fig). Of these, 530 (81%) are bound by both MITF and TFAP2A (Fig 3E, S10 Table). Interestingly, genes involved in melanocyte differentiation, including those encoding proteins expressed in the melanosome, are associated with SEs bound by MITF only (e.g. TYR, MLANA, SLC24A5, DCT) and SEs bound by both MITF and TFAP2A (MLPH, OCA2, TRPM1, MC1R). Exceptions to this pattern include KIT, which is associated with one SE bound solely by TFAP2A and one SE bound solely by MITF, and TYRP1, which is associated with an SE bound by neither (Fig 3E). Taken together, these results show that TFAP2A and MITF bind regulatory elements associated with melanocyte differentiation effectors. Notably, 409 (63%) of all SEs are bound by TFAP2A peaks that overlap with MITF peaks. It remains to be determined whether TFAP2A and MITF exhibit cooperative binding at these loci.

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Table 1. Mouse coat color genes associated with peaks of TFAP2A and MITF that overlap active enhancer marks (H3K27ac).

https://doi.org/10.1371/journal.pgen.1006636.t001

While several of the pigmentation genes associated with active TFAP2A and MITF peaks showed TFAP2A-dependent expression in both zebrafish and mouse, we also noted many apparently TFAP2A-independent genes on this list. One possible explanation is that the presence of a TFAP2A peak does not signify contribution of TFAP2A to the activity of a given regulatory element. Alternatively, activity of a redundantly-expressed TFAP2 paralog may compensate for the loss of TFAP2A. To rule out the first possibility, we focused on the gene TRPM1, which has a promoter-proximal TFAP2A peak in both mouse and human melanocytes but is expressed at high levels in melanocytes in a TFAP2A-independent manner. In addition, expression of TRPM1 is a sensitive readout of MITF activity levels [79], and a minimal TRPM1 promoter, which has an MITF peak in melanoma cells [17] and primary melanocytes [18], has activity in melanoma cells that is lost upon deletion of the MITF binding sites [86]. We engineered the TRPM1 promoter into a vector suitable for quantitative luciferase reporter assays and created variants with mutations in either the E-Box MITF binding site shown previously to be most important for promoter activity (ΔE1) [86], or four predicted TFAP2 binding sites (ΔAP2) (Fig 3F). These constructs, together with a control vector, were transfected into M21 melanoma cells, which express TFAP2A (S2C Fig). The intact TRPM1 promoter drove much higher luciferase expression than the empty reporter vector, while the ΔE1 promoter variant drove about 50% of intact, and the ΔAP2 variant drove about 10% of intact (p = 0.01) (Fig 3G). These results show that TFAP2A directly activates the TRPM1 promoter, supporting the hypothesis that other TFAP2 paralogs are able to compensate for the absence of TFAP2A at certain melanocyte genes.

Mouse Tfap2a / Tfap2b double conditional mutants are depleted of melanocytes

Testing whether TFAP2 paralogs function redundantly in melanocyte development requires simultaneous depletion of all such paralogs expressed in melanocytes. In mouse melanocytes, Tfap2a and Tfap2b have the highest and second highest expression, respectively, while Tfap2c and Tfap2e are undetectable [87,88]. To determine whether Tfap2 paralogs function redundantly in murine melanocyte development, we generated double conditional mutants (DCM) using a previously published Wnt1-Cre transgenic line [89] and conditional alleles of Tfap2a [24] and Tfap2b (EVO and TW, in preparation). We then utilized two approaches to assess melanocyte development in DCM embryos, corresponding single conditional mutant (SCM) embryos, and control embryos. First, embryos in which the Rosa26-reporter (r26r)-allele [90] was also incorporated were dissected at embryonic day 12.0 (E12.0) and subsequently stained for β-galactosidase (β-gal) activity. The r26r-allele used in combination with the Wnt1-Cre transgene results in β-gal positive staining of premigratory neural crest cells and subsequent derivatives (Fig 4A–4D, S8A–S8D Fig). β-gal-positive melanoblasts and corresponding melanocytes migrate ventrolaterally from the dorsal neural tube and can be identified by their position just below the developing surface ectoderm, most easily observed dorsal to the hindlimb. Examination of control (Fig 4A), Tfap2a SCM (Fig 4B), and Tfap2b SCM (Fig 4C) embryos revealed roughly equivalent numbers of stained cells with a similar distribution. In contrast, Tfap2a/Tfap2b DCM embryos have many fewer β-gal-positive cells in this location (Fig 4D). Second, embryos were processed for in situ hybridization with Pmel [91] and Dct [92,93] riboprobes, detecting melanoblasts and differentiated melanocytes (Fig 4E–4L). As with the r26r experiments, this staining labeled similar numbers of Pmel-positive and Dct-positive cells in control (Fig 4E and 4I), Tfap2a SCM (Fig 4F and 4J), and Tfap2b SCM (Fig 4G and 4K) embryos, but far fewer cells in Tfap2a/Tfap2b DCM embryos (Fig 4H and 4L). The absence of Pmel-positive and Dct-positive melanoblasts in DCMs was evident from the time these cells emerged in control embryos at E10.5 and E11.5, suggesting that the reduced melanoblast number in DCMs is not the result of impaired melanoblast migration (S8E–S8L Fig). Because TFAP2 paralogs have been shown to function during the early stages of neural crest induction [33,34], we next tested whether the observed reduction in melanocytes could be explained by a disruption in this step. Both lineage tracing with the r26r-reporter line (S9A–S9F Fig) and in situ hybridization with a Sox10 riboprobe at E9.5 (S9G and S9H Fig) revealed relatively normal neural crest induction in DCMs, as in controls. Consistent with this observation, α-neurofilament immunostaining (Fig 4M and 4N) and lineage tracing (Fig 4O and 4P, S10A–S10D Fig) identified the initial formation of an alternate trunk neural crest derivative, dorsal root ganglia (DRG), in both DCMs and controls. However, similar to the melanocyte lineage, the neural crest-derived enteric nervous system (ENS) was disrupted in Tfap2a SCM embryos and completely failed to populate the gastrointestinal tract of DCM embryos (S10E–S10L Fig). Broadly, these analyses suggest that in mouse embryos, paralogous proteins TFAP2A and TFAP2B act redundantly subsequent to neural crest induction within distinct neural crest lineages. The virtual absence of melanoblasts in Tfap2a/Tfap2b DCM embryos implies that they contribute to specification and differentiation of the melanocyte lineage, similar to the functions of MITF.

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Fig 4. TFAP2A and TFAP2B redundantly regulate murine melanocyte development.

(A-D) Lateral views of E12.0 control (A), Tfap2a SCM (B), Tfap2b SCM (C), or Tfap2a/Tfap2b DCM (D) mouse embryos processed for β-galactosidase (β-gal) staining. β-gal+ cells are a result of recombination of the r26r-allele by the Wnt1-Cre transgene, labeling neural crest cells and derivatives. (E-L) In situ hybridization expression patterns for Pmel (E-H, dorsal views) or Dct (I-L, lateral views) in control (E, I), Tfap2a SCM (F, J), Tfap2b SCM (G, K), or Tfap2a/Tfap2b DCM (H, L) E12.0 mouse embryos. Insets in (A-L) show higher magnification images just above the hindlimb in a similar viewing plane as the low magnification image. (M, N) Lateral views of the trunk processed for ɑ-neurofilament immunoreactivity in an E10.5 control (M) or Tfap2a/Tfap2b DCM (N), revealing developing dorsal root ganglia (a subset labeled with arrowheads). (O, P) Immunofluorescence of cryosections (cross-sectional plane at the level of the hindlimb) of an E12.0 control (O) or Tfap2a/Tfap2b DCM (P), containing a Tomato-reporter (rTMr) allele labeling neural crest cells and derivatives (arrowheads highlight a subset of melanocytes in the ventrolateral pathway). Abbreviations: DCM, double conditional mutant; drg, dorsal root ganglia; hl, hindlimb; nt, neural tube; SCM, single conditional mutant (A = Tfap2a or B = Tfap2b). Scale bars = 250μM, except M, N = 500μM.

https://doi.org/10.1371/journal.pgen.1006636.g004

Zebrafish mitfa;tfap2a double mutants display a synergistic genetic interaction

Given the widespread co-occupancy of regulatory elements by TFAP2A and MITF, we predicted that mitfa;tfap2a double mutant zebrafish would have a greater-than-additive melanocyte phenotype consistent with synergistic interaction between these genes. To test this, we used the hypomorphic allele mitfa^z25 to reduce Mitfa levels without eliminating the melanocyte lineage, as occurs with total loss-of-function alleles such as mitfa^w2 [94,95]. Compared to wildtype embryos at 72 hpf (Fig 5A), mitfa^z25/z25 homozygotes have fewer and less dendritic melanocytes (Fig 5B). Melanocytes are further reduced, punctate, and noticeably paler in mitfa^w2/z25 trans-heterozygous embryos (Fig 5C). Incrosses of mitfa^w2/z25;tfap2a^+/- double heterozygous adults and mitfa^+/+;tfap2a^+/- single heterozygous adults yielded various combinations of mitfa;tfap2a genotypes. Embryos were sorted into phenotypic bins at 72 hpf, photographed individually, and genotyped for both mitfa^z25 (when appropriate) and tfap2a. Relative to wildtype embryos (Fig 5A), mitfa^+/+;tfap2a^-/- homozygous mutants exhibited a clear reduction of melanocytes in the ventral stripe caudal to the tail, reflecting decreased melanocyte migration and/or proliferation (Fig 5G). However, mitfa^+/+;tfap2a^+/- heterozygous mutants could not be distinguished from wildtypes (compare Fig 5A and 5D). Quantification of ventral tail melanocytes (count area indicated by brackets in Fig 5A–5I) revealed that there is some variation within each genotype, but we detected no genotype/phenotype correlation between tfap2a^+/+ and tfap2a^+/- embryos (Fig 5J, one-way ANOVA). In contrast, both the mitfa^z25/z25 (Fig 5B, 5E and 5H) and mitfa^w2/z25 (Fig 5C, 5F and 5I) backgrounds had three distinguishable phenotypic groups corresponding to the tfap2a genotype, with tfap2a^+/- embryos showing significant differences in the number of ventral tail melanocytes compared to both tfap2a^+/+ and tfap2a^-/- embryos (Fig 5K, one-way ANOVA). Thus, the phenotype of double mutants appears to be more severe than the combination of the phenotypes in single mutants. The biochemical underpinning of this genetic interaction is unknown; Mitfa and Tfap2a may interact synergistically at one or more regulatory elements, or Mitfa may regulate tfap2a expression in melanocytes.

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Fig 5. tfap2a and mitfa display a genetic interaction in zebrafish.

(A-I) Compared to wildtype embryos at 72 hpf (A), melanocytes in mitfa^z25/z25 mutant embryos (B) are dark but punctate, and melanocytes in mitfa^w2/z25 trans-heterozygous embryos (C) are pale, punctate, and reduced in number. While heterozygous mutation of tfap2a in the wildtype background (tfap2a^+/-) (D) does not result in a melanocyte phenotype, mitfa^z25/z25;tfap2a^+/- mutants (E) and mitfa^w2/z25;tfap2a^+/- mutants (F) display a loss of melanocytes in the ventral stripe (brackets). Similarly, the well-characterized phenotype of fewer, paler melanocytes in tfap2a^-/- null mutants (G) is more pronounced in both the mitfa^z25/z25 mutant background (H) and the mitfa^w2/z25 mutant background (I). (J, K) Counts of ventral stripe melanocytes (see brackets in A-I) showed no difference between wildtype and tfap2a^+/- embryos (J, n = 10+), while tfap2a^+/- embryos in both mitfa mutant backgrounds had significantly fewer melanocytes in the ventral stripe (K, n = 4–8). One-way ANOVA: *p<0.05, **p<0.01, ****p<0.0001.

https://doi.org/10.1371/journal.pgen.1006636.g005

Forced expression of tfap2a does not rescue melanocytes in mitfa mutant zebrafish

Genetic interaction between tfap2a and mitfa supports the idea that the factors encoded by these genes regulate shared targets in melanocytes, possibly within single or converging pathways. Previously, we found that elevating expression of mitfa (under the sox10 promoter) partially compensates for the reduction of Tfap2 in tfap2a/tfap2e-depleted zebrafish embryos [30]. Thus, we next investigated whether the reciprocal experiment, artificially elevating expression of tfap2a, could rescue melanocyte development in the absence or reduction of Mitfa. To test this, we engineered the tfap2a cDNA to encode 6 Myc epitopes at the carboxy terminus. To confirm that the epitope-tagged Tfap2a was still functional, we injected tfap2a-Myc mRNA into embryos depleted of tfap2a and tfap2c and observed differentiated melanocytes, which are otherwise completely absent from such animals [33]. We then fused tfap2a-Myc cDNA downstream of the mitfa promoter, which is active in melanocytes (mitfa promoter described in [96]), and injected the mitfa:tfap2a-Myc plasmid into embryos derived from an incross of mitfa^w2/z25 adults. At 48 hpf, embryos were sorted into three groups based on the mitfa mutant phenotype, fixed, and subsequently processed for anti-Myc immunoreactivity. Ten plasmid-injected embryos and ten uninjected control embryos were documented in each group. We detected brightly-labeled cells, including several melanocytes, in both mitfa^z25/z25 (Fig 6A and 6B) and mitfa^w2/z25 (Fig 6C and 6D) plasmid-injected embryos, whereas uninjected controls had no labeled cells. However, the anti-Myc immunoreactive melanocytes were phenotypically indistinguishable from neighboring unlabeled melanocytes by morphology, dendricity, or pigmentation (Fig 6B and 6D, arrows). The mitfa^w2/w2 plasmid-injected embryos had similar numbers of anti-Myc immunoreactive cells to the other genotypes, but none showed any hint of pigmentation (Fig 6E and 6F). We did observe a highly dendritic but wholly unmelanized cell, which is likely to be a xanthophore (Fig 6F, arrow). In summary, elevating tfap2a expression under the mitfa promoter was not sufficient to rescue melanocytes in mitfa^w2/w2 null mutants, nor did it improve the quality or pigmentation of melanocytes in mitfa^z25/z25 and mitfa^w2/z25 mutants. We conclude that, despite regulating many of the same targets, Tfap2a is unable to replace Mitfa in the melanocyte lineage, at least at the dose of over-expression tested here.

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Fig 6. Forced expression of tfap2a does not rescue melanocytes in mitfa mutant zebrafish.

(A-D) Nuclei of melanocytes expressing mosaic Tg(mitfa:tfap2a-Myc) are brightly labeled after anti-Myc immunostaining. At 48 hpf, labeled melanocytes in mitfa^z25/z25 mutant embryos (A, B) and mitfa^w2/z25 mutant embryos (C, D) display no apparent improvement in pigmentation or dendricity compared to adjacent unlabeled cells (B and D, white arrowheads). (E, F) In mitfa^w2/w2 mutants, all labeled cells are unpigmented and resemble other cell types including xanthophores (F, white arrowhead).

https://doi.org/10.1371/journal.pgen.1006636.g006

Cell culture

Microarrays and qRT-PCR

ChIP-seq and ChIP-qPCR

ChIP-seq data analysis

The ChIP-seq Tool Set in Galaxy was used for all peak overlap analyses [107]. For comparison of human ChIP-seq peaks to gene expression, we used a published RNA-seq expression profile of human penis foreskin melanocytes from the Roadmap Epigenomics Project (GEO accession number, GSM958174) [64]. Motif enrichment analysis was carried out using the MEME-ChIP suite [108], including CentriMo [60], MEME-ChIP [69], and AME [109] tools. For gene set enrichment analysis, we used the Genomic Regions Enrichment of Annotations Tool (GREAT), with the association rule basal plus extension, proximal: 5 kb upstream, 1 kb downstream, plus distal: up to 100 kb [72]. ChIP-seq read density clustering analysis and quantitative comparisons were performed using k-means linear enrichment cluster function in seqMINER with the following parameters: window size = -5K to +5K, read extension = 200bp, seed = 12 [110] (http://bips.u-strasbg.fr/). The Panther Classification System was used for GO term enrichment analysis on gene lists [76,77].

RNA-seq in mouse melan-a cells

RNA sequencing (RNA-seq) libraries were prepared with the TrueSeq stranded mRNA kit (Illumina) and sequenced on the Illumina HiSeq 2000 platform. The ten 5’-most bases were trimmed from all of the raw RNA-seq reads. Reads were aligned to the mouse reference genome sequence (mm9) using the STAR alignment software (v. 2.3.0e). RNA-seq reads derived from rRNAs were removed using the split_bam.py script available in RSeQC (v. 2.3.7), using genomic locations of known rDNAs that were downloaded from UCSC. Counts for RNA-seq reads mapping to Ensembl-annotated transcripts (release 67) were calculated using the htseq-count software (v. 0.5.3p3). These raw RNA-seq counts were used for differential gene expression analysis that was performed using DESeq2 (v. 1.10.1).

Promoter deletion analysis

The following versions of the TRPM1 promoter sequence (748 bp) were obtained as double stranded gBlocks gene fragments from IDT: intact sequence, ΔAP2A with mutations in four TFAP2A binding sites, and ΔE1 with a mutation in the main MITF binding site. The consensus TFAP2A binding site GCCNNNGG was disrupted by changing the two underlined bases to T, whereas the E1 MITF site was changed as previously published by [86]. Fragments were cloned into a Tol2-cfos-FFluc vector via Gibson assembly and confirmed with Sanger sequencing.

M21 melanoma cells were grown to 70–90% confluency in a 24-well culture plate. In each well, the reporter plasmid (1μg) and a β-galactosidase plasmid (100ng) were transfected using Lipofectamine 3000 (Life Technologies). Approximately 48 hours after transfection, luciferase assays were conducted using the Dual-Light System from Applied Biosystems, and 20/20n Luminometer (Turner Biosystems, Sunnyvale, CA) according to the manufacturer protocols. Briefly, cells were washed with cold PBS and incubated in 40 μL lysis solution on a shaker for 15 minutes. Cell lysates were transferred to tubes and centrifuged at 12000 rpm for 2 minutes at 4°C, and supernatant was transferred to a clean tube. 10 μL of cell lysate was added to 25 μL of Buffer A and placed in the luminometer, where the injector adds 100 μL Buffer B/Galacton-Plus substrate and reads the signal after 1 second. Samples were incubated in the dark for 30 minutes before injection of 100 μL Accelerator-II and measurement of the β-gal signal after 1 second. For each version of the TRPM1 promoter element, transfection was carried out in triplicate. Firefly luciferase reads from each sample were normalized to the respective β-gal reads, and an average signal was calculated within groups.

Mouse genetics and immunostaining

Experiments utilizing mice in this study were carried out in strict accordance with the recommendations in the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health. The protocol was approved by the Institutional Animal Care and Use Committee of the University of Colorado Denver. Noon on the day a copulatory plug was present was denoted as embryonic day 0.5. Mice used in this study included males that were heterozygous for a Tfap2a-null allele [113], heterozygous for a newly generated Tfap2b-null allele (EVO/TW, in preparation), and hemizygous for the Wnt1-Cre transgene [89] (Tfap2a^null/wt;Tfap2b^null/wt;Wnt1-Cre). These males were crossed with females that were homozygous for both a Tfap2a-conditional [24] and Tfap2b-conditional (EVO/TW, in preparation) allele, resulting in a 1:8 frequency of generating single or double conditional mutants, as well as various other genotype combinations. Mice were maintained on an outbred Black Swiss background. Of note, by virtue of this breeding scheme, the single conditional mutants would always be conditionally heterozygous for the alternate paralog. For r26r [90] and ‘tomato’ [114] experiments, the female was also homozygous for the reporter allele. Yolk sacs or tail clips were used for genotyping. DNA for PCR was extracted using DirectPCR Lysis Reagent (Viagen Biotech. Inc) plus 10 ug/ml Proteinase K (Roche) followed by heat inactivation at 85°C for 45 min. Samples were then used directly for PCR-based genotyping using allele specific primers (available upon request) at a concentration of 200 nM using the Qiagen DNA polymerase kit, including the optional Q Buffer solution (Qiagen).

With the exception of whole-mount β-galactosidase (β-gal) staining, procedures used for mouse embryo analysis (in situ hybridization, ɑ-neurofilament immunostaining, and immunofluorescence) have all been previously described [115]. Following euthanasia, embryos were collected at the indicated time-points in DEPC-PBS and subsequently processed. Briefly, for in situ hybridization, after embryo collection, trunks were removed, fixed overnight in 4% paraformaldehyde (PFA), and stained using the indicated riboprobe (Pmel, Dct, Sox10). For immunostaining, embryos were processed using an ɑ-neurofilament primary antibody [116] (IgG clone 2H3, obtained from the Developmental Studies Hybridoma Bank–University of Iowa), followed by colorimetric staining using a standard secondary antibody and DAB-detection. For β-gal immunofluorescence, collected embryos underwent a short fixation in 0.25% glutaraldehyde, were taken through a series of sucrose/O.C.T. solutions (Tissue-Tek O.C.T. Compound, Electron Microscopy Sciences), until being embedded in 100% O.C.T. and frozen on dry-ice. Cryosections were then cut at ~12μM on a Leica CM 1900 cryostat (Leica Biosystems Inc.) using the hindlimb as a delimiting rostral-caudal boundary between samples. Following sectioning, slides were washed in PBS, 2 x 10 min, blocked in 3% BSA (in PBS) 1 hr at room temperature, incubated over-night with a rabbit polyclonal anti-β-gal antibody (Product 55976, MP Biomedicals, LLC) diluted 1:200 in block solution. Subsequently, sections were washed 2 x 10 min in PBS, followed by a 1 hr incubation in goat-anti-rabbit Alexa Fluor 488 (Thermo Fisher) and counterstained with DRAQ5 (Abcam), washed again 2 x 10 min in PBS, and then cover-slipped. Processed samples were imaged on a Leica TCS SP5 II confocal microscope and representative images taken. Finally, for β-gal staining, embryos were collected at appropriate time points and fixed ~1hr at room temperature with 0.25% glutaraldehyde in PBS. Subsequently, embryos were washed 3 x 30 minutes in a ‘lacZ rinse buffer’ (0.2M sodium phosphate, 2mM magnesium chloride, 0.02% NP40, and 0.01% sodium deoxycholate), and then incubated overnight in a ‘lacZ staining solution’ (lacZ rinse buffer plus 5mM potassium ferricyanide, 5mM potassium ferrocyanide, and 1 mg/ml X-gal) at 37°C. Following adequate staining, embryos were post-fixed in 4% PFA overnight, moved to PBS, and subsequently imaged.

Zebrafish genetics and immunostaining

Zebrafish alleles used in this study include tfap2a^low [lockjaw allele, 28], mitfa^w2 [94], and mitfa^z25 [95]. Lines were maintained as mitfa trans-heterozygous and tfap2a heterozygous (mitfa^w2/z25;tfap2a^+/-) or mitfa wildtype and tfap2a heterozygous (tfap2a^+/-). For genetic interaction experiments, mitfa^w2/z25;tfap2a^+/- animals were incrossed to generate tfap2a^+/+, tfap2a^+/-, and tfap2a^-/- genotypes in each of mitfa^z25/z25 and mitfa^w2/z25 backgrounds. To obtain control animals in the mitfa^+/+ background, mitfa^+/+;tfap2a^+/- animals were also incrossed. At 72 hpf, embryos were binned according to melanocyte phenotype, photographed, and genotyped via High Resolution Melt Analysis (HRMA) using Precision Melt Supermix (Bio-Rad) on a CFX96 Real-Time PCR Detection system (Bio-Rad) according to the default settings. Precision Melt Analysis Software (Bio-Rad) was used to analyze melt curves. HRMA primer sequences are as follows: tfap2a^low (forward: GTA GCT ATG TTT CGT GGT TA; reverse: ACA ATA AGC AGC TGC TTT AC), mitfa^z25 (forward: GCA GAA GTC AGA GCC CTG GC; reverse: ACG GAT CAT TTG ACT TGG GAA TTA AAG). Ventral stripe melanocytes were counted from lateral-view images taken at 72 hpf and statistical significance between groups was tested using a one-way ANOVA with multiple test correction.

For zebrafish rescue experiments, mitfa^w2/z25 trans-heterozygotes were incrossed to obtain mitfa^z25/z25, mitfa^w2/z25, and mitfa^w2/w2 genotypes, which were clearly distinguishable based on melanocyte phenotype at 48 hpf. 3-way Gateway cloning technology was used to generate a pDestTol2CG2 plasmid [117] containing 5’ entry mitfa promoter [96], middle entry tfap2a cDNA, and 3’ entry 6x Myc epitope tag followed by polyA, resulting in the presence of Myc epitope tags on the carboxy terminus of Tfap2a. This construct was then injected into the above cross along with Tol2 mRNA, both at a concentration of 30 ng/uL. Due to the presence of a cmlc2:GFP reporter in the vector, injected embryos were screened for successful plasmid integration based on expression of GFP in the heart at 28–30 hpf. GFP positive embryos and uninjected control embryos were fixed overnight in 4% paraformaldehyde at 48 hpf. After fixation, embryos were rinsed 3x in PBS, blocked in PBDT+2.5% goat serum for 1 hr at room temperature, and incubated in anti-Myc primary overnight at 4°C (9E10, obtained from the Developmental Studies Hybridoma Bank—University of Iowa, 1:100 diluted in block solution). The following day, embryos were rinsed 4x in PBS + 0.1% Triton X-100 (PBS-Tx) and incubated overnight at 4°C in Alexa Fluor goat-anti-mouse 488 secondary (Thermo Fisher) diluted in block solution. Embryos were then rinsed 4x15 minutes in PBS-Tx and mounted on slides. Ten individual embryos of each genotype with plasmid injection or uninjected control were viewed at 40x and photographed.

Immunostaining in cell lines

Cells were grown to approximately 75% confluency in 24-well plate on poly-lysine coated disks and fixed with 4% paraformaldehyde for 1 hr at 4°C. Following fixation, cells were rinsed 3x20 minutes with 1x PBS, permeabilized in 1x PBS + 0.3% Triton X-100 for 30 min at 37°C, rinsed 3x10 minutes in PBS-Tx, and blocked (1% BSA in PBS-Tx) overnight at 4°C. Cells were then incubated in anti-TFAP2A primary overnight at 4°C (3B5, Santa Cruz Biotechnology, Inc., 1:100, 250ug/mL stock diluted in block solution). The following day, cells were rinsed 3x20 minutes in PBS-Tx and incubated for 2 hr at room temperature in Alexa Fluor goat-anti-mouse 488 secondary (Thermo Fisher) diluted in block solution. Cells were then rinsed 4x15 minutes in PBS-Tx and disks mounted in Prolong Gold antifade reagent and DAPI SlowFade solution (Life Technologies). Slides were imaged on a Zeiss LSM 700 Flexible Confocal Microscope (Carl Zeiss Microscopy). Briefly, for quantification of immunofluorescent intensity approximately three 20x images were taken of each slide, and three slides of each cell line processed (including a no primary control). Each image included both the antibody of interest as well as DAPI staining to identify cell nuclei.

Ethics statement

All experiments were approved by the University of Iowa or University of Colorado Institutional Animal Care and Use Committee (IACUC). We abide by PHS Policy, USDA-Animal Welfare regulations, the Guide for the Care and Use of Laboratory Animals, University of Iowa policies and regulations and any state and local laws and regulations.