Loss of Atoh1 Promotes Tumor Formation in Two Colorectal Cancer Mouse Models

In the mouse, Atoh1 is a master regulator of secretory cell fate commitment in the intestinal epithelium [2,11]. To investigate whether Atoh1 plays a role in intestinal tumors, we analyzed the function of the mammalian ato homolog, Atoh1, in colon tumorigenesis. We assessed the tumor susceptibility of mice with an intestine-specific deletion of Atoh1 (Atoh1^Δintestine) [3] in two different established mouse models of CRC.

We first treated Atoh1^Δintestine mice with azoxymethane (AOM), a chemical carcinogen that preferentially induces colon tumors. We found a significant enhancement in polyp formation in the large intestines of Atoh1^Δintestine compared to wild-type littermate mice, characterized by increased incidence (33% [4/12] vs. 100% [8/8], p < 0.005; Figure 1A), multiplicity (0.75 vs. 10.3 polyps/colon, p < 0.0008; Figure 1C), and size (2.1 vs. 4.1 mm/polyp, p < 0.0004; Figure 1E) when examined under the dissecting microscope. Histological analysis of colons from AOM-treated Atoh1^Δintestine mice confirmed these findings and showed a range of histological phenotypes, from severely hyperplastic mucosa with embedded multifocal adenomas, to large, highly dysplastic adenomas, with one animal having invasive adenocarcinoma (Figure 2A, 2A′, and 2A′′). In contrast, histological examination of AOM-treated Atoh1^wt littermates showed none with adenomatous changes; with all polyp-like structures shown to be gut-associated lymphoid tissue (GALT; Figures 2B and S1). These data support the notion that loss of Atoh1 can be an initiating event in mammalian cancer formation.

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Figure 1. Loss of Atoh1 Enhances Tumor Formation in the Mouse Colon

(A) shows the incidence of polyps in AOM-treated mice.

(B) shows the incidence of polyps in the Atoh1^wt and Atoh1^Δintestine mice in an APC^min background.

In (A) and (B), the two-tailed Fisher exact test was used.

(C) Bar graph shows the average number of macroscopically visible polyps (>1 mm) in the colons of AOM-treated Atoh1^wt (WT; n = 9 polyps in 12 mice, white bars) and Atoh1^Δintestine (n = 82 polyps in 8 mice, black bars).

(D) The average number of polyps in the colons of APC^min (n = 27 polyps in 27 mice) and APC^min; Atoh1^Δintestine (n = 123 polyps in 10 mice).

(E and F) The average maximum diameter of each polyp is shown as a bar graph for Atoh1^wt or Atoh1^Δintestine mice. (E) Comparison of polyp size from AOM-treated Atoh1^wt and Atoh1^Δintestine mice; and (F) from APC^min and APC^min; Atoh1^Δintestine mice.

For (C–F), the two-tailed Student t-test was used to measure significance. Error bars indicate the standard error of the mean. Double asterisks (**) indicate p <0.01; triple asterisks (***) indicate p <0.001.

https://doi.org/10.1371/journal.pbio.1000039.g001

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Figure 2. Increased Proliferation Contributes to Tumorigenesis in Atoh1^Δintestine Colon

(A) Hematoxylin and eosin staining of a representative adenoma in AOM-treated Atoh1^Δintestine colon (5× magnification). The section is in the rectal region and is characterized by cystic structures with severe crypt hyperplasia with multifocal embedded adenomas and severe lymphocyte infiltration of the submucosa. (L, lumen; M, muscle). (A)′ and (A)′′ show higher magnification (20×) of two areas of the adenoma, indicating the aberrant branched crypt structures that are imbedded in the submucosa.

(B) The colons of AOM-treated Atoh1^wt mice showed large GALT (highlighted by arrows) that were macroscopically counted as polyps. Magnification is 5×. Note the relative size of the GALT compared to adjacent normal-appearing crypts.

(C) Histological analysis of colon in APC^min.

(D) Histological analysis in APC^min; Atoh1^Δintestine mice show highly dysplastic adenomas with cystic structures and lymphocyte infiltration.

(E) Proliferation was measured by determining the percentage of BrdU-positive epithelial cells in AOM-treated Atoh1^wt and Atoh1^Δintestine normal-appearing colonic crypts. The white bar represents crypts from Atoh1^wt mice; the gray bar represents nondeleted Atoh1^wt crypts, and the black bar Atoh1-null crypts in Atoh1^Δintestine mice.

(F) Apoptosis was measured by determining the percentage of cleaved caspase-3 (c-Caspase 3)-positive epithelial cells in AOM-treated Atoh1^wt and Atoh1^Δintestine normal-appearing colonic crypts. Bar shading indicates crypt genotype as in (E).

For each graph in (E and F), the two-tailed Student t-test was used to measure significance: a single asterisk (*) indicates p < 0.05; double asterisks (**) indicate p < 0.01. Error bars indicate the standard error of the mean.

https://doi.org/10.1371/journal.pbio.1000039.g002

In previous work, we found that ablation of Atoh1 leads to an increase of proliferation but has no effect on apoptosis [3]. We therefore examined proliferation and apoptosis in the “preneoplastic” normal-appearing epithelium within the colons of AOM-treated Atoh1^wt and Atoh1^Δintestine mice. As in the Drosophila model, we find more proliferation of epithelial cells in Atoh1^Δintestine crypts compared to nonrecombined wild-type crypts within the same animals, or compared to crypts from Atoh1^wt littermates (Figures 2E and S2D–S2F) but no obvious difference in apoptosis (Figures 2F and S2G–S2I).

We extended our tumor analysis with an independent genetic mouse model for CRC by crossing Atoh1^Δintestine mice to APC^min mice (APC: ENSMUSG00000005871), in which Wnt signaling is constitutively activated [12,13]. APC^min mice develop spontaneous adenomas primarily in the small intestine, with only occasional polyps in the colon [14]. Interestingly, Wnt signaling is thought to interact negatively with ato during sense organ formation in the developing Drosophila epithelium [15]. Comparing the large intestines of APC^min; Atoh1^wt mice to APC^min; Atoh1^Δintestine littermates at 16 wk, we find significantly more polyps in the colons of double-mutant mice, characterized by increased incidence (52% [14/27] vs. 100% [10/10], p <0.007; Figure 1B), multiplicity (1.1 vs. 12.3 polyps/colon, p <0.0001; Figure 1D), and size (3.3 vs. 4.3 mm/colon, p <0.0002; Figure 1F). Histological examination confirmed these polyps as adenomas in both APC^min; Atoh1^wt and APC^min; Atoh1^Δintestine mice, with two polyps in APC^min; Atoh1^Δintestine mice having progressed to invasive adenocarcinoma (Figure 2C and 2D). The polyps in the APC^min; Atoh1^Δintestine mice originated from the Atoh1 mutant crypts as indicated by the absence of goblet cells in the polyps (Figure S3A and S3B).

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Figure 3. Loss of ATOH1 Expression and Genomic Deletions in CRC Patient Samples

(A) ATOH1 expression normalized to GADPH and control colon samples: ATOH1 expression is lower in tumor compared to control samples. Additionally, adenocarcinomas have significantly lower ATOH1 expression than adenomas. Box plot indicating 25–75 percentiles, central line indicates the median, red cross indicates the mean. Error bars indicate next data point. Outliers are represented as dots. Double asterisks (**) indicate p < 0.01 (t-test).

(B) Deletions in the ATOH1 locus. Upper dashed line indicates the upper limit of single-deletion detection set at 0.70 of the ratio of ATOH1 locus to control locus. White and gray bars indicate different primer sets.

(C) Endogenous ATOH1 expression in different MCC14.2-derived cell lines and human keratinocytes (HK) in response to Atoh1 expression.

(D) Detection of methylation at the ATOH1 locus using pull-down assay of methylated DNA. Bands in “M-CpG pulldown” indicate positive for ATOH1 methylation. “Input” shows DNA input before the pull-down. Samples were processed blindly. Internal methylated and unmethylated controls are shown in the last four lanes.

(E) Inhibition of methyltransferase activity with 5-aza-deoxycytosine for 7 d leads to an increase of ATOH1 expression in the Ht29 cell line. Error bars indicate the standard deviation.

(F) Graph representing observations on the genomic and mRNA levels. First column: percentage of patients showing deletions or duplications (black: deletions, gray: two copies, and white: duplications). Second column: percentage of patients showing methylation using the pull-down of methylated DNA assay (black: methylated, and white: not methylated). Third column: mRNA expression of CRC samples versus control colon (black: low expression, grey: normal expression, and white: high expression).

https://doi.org/10.1371/journal.pbio.1000039.g003

In summary, thus far, loss-of-function analysis of Atoh1 in two mouse models of colon tumorigenesis supports a role for Atoh1 as a key switch in tumor formation and progression. This role appears to be mediated by increased cell proliferation in the absence of Atoh1. Importantly, the mouse tumors lack the secretory cell types that depend on Atoh1 for their formation. Thus, the role of Atoh1 in tumor formation is likely linked to its function as a master regulator of differentiation.

Loss of ATOH1 Expression in Primary Human CRC and MCC Tumors

Given the remarkable evolutionary conservation of ato/Atoh1 function in cancer development, we decided to analyze whether loss of the human ortholog, ATOH1, might be selected for during malignant transformation in human cancer. To this end, we investigated the role of ATOH1 in CRC and MCC. In vitro overexpression experiments in CRC cell lines suggest that ATOH1 gain of function decreases the population growth potential of CRC cells [10]. Similarly, expression analysis in MCC cell lines suggests an inverse correlation between ATOH1 levels and the population growth of MCC cells (Figure S4A–S4C) [16]. These observations hint at a potential role for ATOH1 in these cancers.

To test this possibility in vivo in primary human tumors, we began by asking whether the expression of ATOH1 is down-regulated in primary tumor samples from 42 CRC and four MCC patients. As MCC is a very rare cancer, only four samples were available for analysis. Seventy percent of the CRC samples show a significant decrease of ATOH1 mRNA expression compared to tissue-matched colon samples from normal controls as analyzed by quantitative reverse-transcriptase PCR (RT-qPCR), suggesting that loss of ATOH1 expression is a highly common feature of CRC oncogenesis (Figures 3F and S4). Furthermore, ATOH1 mRNA levels are significantly lower in adenocarcinoma samples compared to adenomas (t-test: p = 0.017), indicating progressive loss of ATOH1 expression levels with increasing tumor severity (Figure 3A). Similarly, in MCC, the two samples with lower ATOH1 expression levels were derived from the two patients showing metastases (Figure S5).

CRC and MCC Patients Show Deletions in the ATOH1 Locus

One mechanism to explain loss of gene expression during oncogenesis is the deletion of the locus, which we tested for using quantitative PCR on genomic DNA in all 46 patient samples, one MCC cell line (MCC14.2), and one CRC cell line (Ht29). At least one deletion in the locus was detected in 57% of the samples (Figure 3B and 3F). A second primer set yielded similar results (49% deletion rate, Figure 3B). One allele was also deleted in both Ht29 and MCC14.2. Comparative genomic hybridization (CGH) array experiments [17] on three samples did not show a deviation of the clones flanking the ATOH1 locus (Figures S5 and S6), indicating that the deletions observed in patients are likely to be ATOH1-specific microdeletions. When only one copy is deleted, complete loss of gene function could be achieved by point mutations in the remaining allele. Sequencing of the ATOH1 open reading frame of 24 samples, however, did not reveal any mutations in any of the samples (unpublished data). Together with the loss of ATOH1 mRNA expression in patients, these results suggest that an epigenetic silencing mechanism may be involved—a possibility we tested in further detail.

The ATOH1 Locus Is Methylated in CRC and MCC Patients

To address a putative epigenetic transcriptional silencing mechanism, we began by asking whether transcription could, in principle, be initiated from the ATOH1 locus in cancer cells. Drosophila ato and mouse Atoh1 are both known to be autoregulatory [18,19]. This provides the opportunity to test whether transcription could be activated from the ATOH1 locus by ATOH1 itself in normal versus cancer cells. We took advantage of expressing the mouse ortholog in human cells to distinguish the expression of endogenous ATOH1 from that of Atoh1. For the tests in human cancer cell lines, we used the MCC14.2 cell line, which is derived from MCC and has strongly reduced expression of ATOH1. Surprisingly, Atoh1 overexpression failed to activate endogenous human ATOH1 expression in the MCC14.2 cell line (Figure 3C), despite the virtually complete conservation of the two proteins. This may be due to a key difference between the two proteins, or to a possible disruption of the autoregulatory loop during oncogenesis. To test whether Atoh1 can activate ATOH1 expression in normal noncancerous skin cells, we transduced normal human primary keratinocytes with an Atoh1 expression vector and tested the expression of endogenous ATOH1 40 h after lentiviral transduction. In contrast to the cancer cell lines, endogenous ATOH1 is up-regulated 38-fold by Atoh1 (Figure 3C). This indicates that the expressability of the ATOH1 locus is inhibited during oncogenesis.

Genomic database searches show that all known ATOH1 orthologs, from human to Drosophila, reside in a CpG island covering at least the promoter and the transcription start site, suggesting that CpG methylation may be a mechanism of ATOH1 loss of function. We therefore assayed all patient samples and both cell lines for ATOH1 methylation using three independent assays: pull-down of methylated DNA, methylation-sensitive restriction digest, and methylation-specific PCR for bisulfite DNA modification. We found that up to 81% of the patients, as well as both cell lines, show methylation of the ATOH1 locus in both the coding sequence and the promoter sequences directly upstream of the ATG. In contrast, none of the control samples show ATOH1 methylation (Figures 3D, 3F, and S7A–S7C). This suggests that methylation is likely causal to the transcriptional silencing of the locus during oncogenesis as this is accompanied by lower ATOH1 expression. In a few cases, however, the correlation was not observed. In two cases (samples 27 and 30), this may be due to the presence of a putative duplication, which is yet to be methylated. Alternatively, but not exclusively, it may be due to heterogeneity of the degree of methylation in different cells comprising the tumor samples analyzed. To provide evidence that methylation silences the ATOH1 locus, we inhibited DNA methyltransferases (Dnmts) with 5-azadeoxycytidine in the Ht29 cell line. This results in an approximately 8-fold increase in ATOH1 expression (Figure 3E). Thus, the ATOH1 locus is methylated in cancer patients and derived cell lines, and this methylation can be reversed, resulting in the transcriptional reactivation of the locus. The autoregulatory function of ATOH1 combined with the methylation of its regulatory sequences might hint to the involvement of ATOH1 in the silencing of its own promoter. To gain some insight into whether this may indeed be the case, we asked whether Atoh1 can physically interact with Dnmt proteins. We tested the binding of Atoh1 to Dnmt1 (ENSG00000130816), Dnmt3a (ENSG00000119772), and Dnmt3b (ENSG00000088305) using GST pull-down assays. We find that Atoh1 binds to all three Dnmts. This is further supported by the ability of GST-Atoh1 to pull down DNA methyltransferase activity from nuclear extracts, similar to the positive control (EED: embryonic ectoderm development, ENSG00000074266) [20]. Focusing on Dnmt1, the maintenance DNA methyltransferase, we confirmed this observation using coimmunoprecipitation experiments in 293T cells transfected with HA-tagged Atoh1. Lastly, mapping experiments identified several regions of Dnmt1 as mediating the association with Atoh1 (Figure S8A–S8D).

In summary, deletions and epigenetic silencing via methylation combine to cause loss-of-function mutations of the human ATOH1 locus in primary human cancers. Together with loss-of-function evidence in five independent cancer models in human cells, Drosophila, and mouse, these data strongly support a role for the Ato/ATOH1 transcription factors as key modulators of oncogenic transformation.

ATOH1 Functions via JNK-Dependent Inhibition of the Cell Cycle and Induction of Apoptosis

To better understand the role that ATOH1 plays in cancer, we sought to determine the molecular mechanism by which it acts to suppress the formation and progression of tumors. Gain- and loss-of-function analyses point to Atoh1-dependent regulation of proliferation in mouse colon tumors (Figures 2E and S3D–S3F). Analysis of the role of Drosophila ato in fly retinal tumors [7] shows that this function is mediated by the JNK signaling pathway. We hypothesized that the Ato-JNK-p21 pathway may mediate the tumorigenic phenotype in our mouse models of CRC as well. If loss of ATOH1 were indeed an initiating event in tumor formation, we reasoned that the effects of Atoh1 function would have to be detectable in preneoplastic tissue as this is where oncogenesis begins. We therefore first examined the expression levels of Atoh1, Cdkn1a (p21, ENSG00000124762), Cdkn1b (p27, ENSG00000111276), and Cdkn1c (p57, ENSG00000129757) and the JNK target cJun (ENSG00000177606) in colon crypts isolated from Atoh1^wt and Atoh1^Δintestine mice (Figure 4A). We observe a 6.8-fold reduction in Atoh1 expression and an approximately 2.8-fold reduction in expression of all three Cdkn1 isoforms in colon crypts from Atoh1^Δintestine mice. We also observe a similar reduction in cJun mRNA levels. We further examined the Ato-JNK-Cdkn1 pathway by immunoblotting proteins from colon polyps and normal-appearing colon tissue from APC^min/+; Atoh1^wt and APC^min/+; Atoh1^Δintestine mice. Colon polyps show higher levels of cJun and p21^waf protein compared to normal colonic tissue. Importantly, however, we find a reduction in cJun levels in APC^min/+; Atoh1^Δintestine polyps compared to APC^min/+; Atoh1^wt polyps, consistent with the mRNA analysis showing less cJun (Figure 4B). Importantly, we find a reduction in p27 protein and a trend toward less p21 in Atoh1-mutant colon tissues and polyps, consistent with our analysis of mRNA levels (Figure 4B). We also find a specific reduction in pJNK1 levels (Figure 4C) in APC^min/+; Atoh1^wt versus APC^min/+; Atoh1^Δintestine tissues, in agreement with earlier reports that JNK1 (MAPK8: ENSG00000107643) may have anti-oncogenic activity in the mouse intestine [21], whereas JNK2 (MAPK9: ENSG00000050748) deficiency enhances tumorigenesis in other epithelia [22]. Finally, we examined the pattern of JNK activation in the colon of Atoh1^wt and Atoh1^Δintestine mice, and observe identical numbers of pJNK1/2-positive cells in Atoh1-null compared to control crypts (Figure S9). Together with western blot analysis demonstrating selective reduction in pJNK1, but not pJNK2, our data suggest that the level of pJNK1 in individual crypt cells is decreased upon loss of Atoh1. These data are consistent with a preneoplastic function of Atoh1 in regulating proliferation, by a JNK-dependent induction of cell cycle inhibitors.

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Figure 4. JNK Pathway Influenced by Atoh1

(A) Gene expression analysis in colon crypts from Atoh1^Δintestine and Atoh1^wt mice. Quantitative RT-PCR was used to assess gene expression in isolated colon crypts. The ratio of gene expression is shown, in which negative numbers indicate reduced expression in Atoh1^Δintestine crypts. t-tests determined p-values shown for each gene.

(B) Western analysis of p27^kip, p21^waf1, and c-JUN in APC^min; Atoh1^wt, and APC^min; Atoh1^Δintestine colonic tissues and polyps. Representative colonic tissue and polyp lysates of APC^min; Atoh1^wt, and APC^min; Atoh1^Δintestine were used for western analysis of p27^kip, p21^waf1, and c-JUN. Actin was used a loading control. p21^waf1 was significantly up-regulated in polyps compared to nonneoplastic colon tissue. c-Jun protein levels were significantly reduced in colonic polyps upon Atoh1 loss. p27 is down-regulated in preneoplastic colonic tissue upon loss of Atoh1. Quantifications are shown in Figure S12A–S12C.

(C) Representative tissues were used for Western analysis of phosphorylated JNK1 and JNK2. Total JNK1 and JNK2 are shown as control. Actin loading control is shown below. pJNK1, but not pJNK2, was significantly reduced in colon tissue from Atoh1^Δintestine compared to Atoh1^wt mice. Quantifications are shown in Figure S12D1–S12D2′.

https://doi.org/10.1371/journal.pbio.1000039.g004

Next we asked whether the molecular mechanism of ATOH1 function in human cancer is similar to the mouse. We took advantage of several established MCC cell lines [23,24] with either high ATOH1 expression (MCC1 and MCC6, derived from less aggressive tumors) or low ATOH1 expression (MCC13, MCC14.2, and MCC26, derived from highly aggressive metastatic tumors; Figure S4A and S4C). We find that the growth rate of cell lines, measured as their doubling time, correlates inversely with levels of ATOH1 expression (Figure S4B). To determine whether the reduction in ATOH1 levels might be causal to decreased doubling time, we restored ATOH1 function by creating stable cell lines expressing Atoh1 using lentiviral vectors (Figure 5A and 5B). Stable Atoh1-expressing cell lines (MCC14.2-Atoh1.1a, MCC14.2-Atoh1.1b, MCC14.2-Atoh1.2a, and MCC14.2-Atoh1.2b) have a significantly slower population doubling time compared to control cell lines (MCC14.2 and MCC14.2-GFP; p <0.0001; Figure 5C). To assess whether this increase in doubling time reflects decreased malignancy, we tested these cell lines for growth in soft agar. Cell lines with high levels of Atoh1 expression display a marked decrease in growth in soft agar compared to control cell lines (Figure 5D, p < 0.001). The change in population doubling time could be due to a slower cell cycle, an increased apoptotic rate, or both. Although the distribution of cells in the cell cycle appears unaltered (Figure S10A), the speed of the cell cycle is 25% lower in Atoh1-expressing cells, as assayed by BrdU pulse-chase experiments (Figure 5E, p <0.01).

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Figure 5. ATOH1 Suppresses Growth by Interfering with the Cell Cycle

(A) Constructs used for creating lentiviral and transfection vectors.

(B) Western blot analysis for Atoh1 and chromogranin on untransduced MCC14.2 cells (lane 1), GFP-transduced MCC14.2 cells (lane 2), and two independently derived MCC14.2 cell lines transduced with Atoh1-IRES-eGFP (lane 3: MCC14.2-Atoh1.1a, and lane 4: MCC14.2-Atoh1.2a). Quantifications in Figure S12E and S12F.

(C) Doubling time in hours for MCC14.2 (lane 1), MCC14.2-GFP (lane 2), and four lines independently transduced with Atoh1-IRES-eGFP (lanes 3–6: MCC14.2-Atoh1.1a, MCC14.2-Atoh1.1b, MCC14.2-Atoh1.2a, and MCC14.2-Atoh1.2b, respectively).

(D) Assay for growth in soft agar. Colonies per view with a 10× lens, lane 1: MCC14.2, lane 2: MCC14.2-GFP, lane 3: MCC14.2-Atoh1.1a, and lane 4: MCC14.2-Atoh1.2a. Error bars indicate the 25th and 75th percentiles.

The triple asterisks (***) in (C and D) indicate a significant difference from MCC14.2 (t-test: p < 0.001).

(E) Percentage of cells positively labeled for BrdU and past S-phase in a BrdU pulse-chase experiment; double asterisks (**) indicate p < 0.01 (t-test).

https://doi.org/10.1371/journal.pbio.1000039.g005

In addition to slower proliferation, we find a specific and strong increase in apoptotic cell death, as measured by annexin-V and cleaved caspase-3 (ENST00000308394), in MCC (MCC14.2 series) and CRC (Ht29) cell lines transduced or transfected with Atoh1 (Figures 6A, 6B, and S10B). This increase in cell death is mediated by the intrinsic apoptosis pathway, as suggested by enhanced caspase-9 (ENST00000333868) cleavage (Figure 6A and 6B).

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Figure 6. ATOH1 Leads to Activation of Apoptosis and Expression of p21^waf1

(A) Western blot analysis for p21^waf1, cleaved caspase-3, cleaved caspase-9, and phosphorylated JNK of lysates of MCC14.2 cells, MCC14.2-GFP, and two MCC14.2 cell lines transduced with Atoh1-IRES-eGFP (MCC14.2-Atoh1.1a and MCC14.2-Atoh1.2a). The corresponding actin loading controls are shown under each blot. Quantifications are shown in Figure S12G–S12J.

(B) Western blot analysis for p21^waf1, cleaved caspase-3, cleaved caspase-9, and p-JNK of lysates of Ht29 cell line transfected with pCLIG-eGFP (left lane) or pCLIG-Atoh1-IRES-eGFP (right lane); actin loading controls are shown under the respective blots. Quantifications are shown in Figure S12K–S12N.

(C) The molecular changes are specific to functional ato: western blot of lysates of Ht29 cells transfected with CMV-ato, CMV-ato¹, or empty vector. Actin loading controls are shown below the respective blots. Quantifications are shown in Figure S12O–S12Q.

(D) Graph expressing ratio of cell numbers of MCC14.2-derived cell lines transfected with pMSCV-ATOH1^ERD-IRES-eGFP versus cell number of cell lines transfected with pMSCV-IRES-eGFP (vector). Statistical analysis was done using t-test under different conditions compared to MCC14.2. Single asterisk (*) indicates p < 0.05; double asterisks (**) indicate p < 0.01.

(E) MCC1 cells transfected with dominant-negative ATOH1^ERD fusion (lanes 1 and 2) and with empty vector control (lanes 3 and 4). Western blot analysis for p21^waf1, phosphorylated JNK, and cleaved caspase-3 on lysates of MCC1 cell line. The actin loading control of each blot is shown below. Quantifications are shown in Figure S12R–S12T.

https://doi.org/10.1371/journal.pbio.1000039.g006

Data from mouse models indicate an involvement of JNK-mediated regulation of cell proliferation for the tumor suppressor effect of Atoh1. To assess whether the same mechanisms are operating in human cancer, we tested the expression levels of the caspase-3 and caspase-9, p21^waf1, and p-JNK in MCC and CRC cells lines. We note a clear and specific up-regulation of p21^waf1 and p-JNK levels upon Atoh1 expression (Figures 6A, 6B, and S10C–S10H). Similarly, transfection of wild-type Drosophila Ato results in the up-regulation of cleaved caspase-3, p21^waf1, and p-JNK (Figure 6C), in contrast to a null-mutant form of the protein that completely fails to bind DNA [25], indicating the conservation and specificity of the Ato/Atoh1 effect.

Next, we tested the expression of these proteins in a loss-of-function setting. We generated and expressed the transcriptional repressor form of ATOH1 by fusing it to the engrailed repressor domain (ATOH1^ERD) [26], which specifically inhibits the Atoh1-induced effects on MCC cell lines (Figure 6D). In the MCC1 cell line, which shows higher endogenous ATOH1 expression compared to MCC14.2 (Figure S4C), expression of ATOH1^ERD leads to a down-regulation of p21^waf1 and p-JNK as well as inhibition of caspase-3 cleavage (Figure 6E).

In summary, gain- and loss-of-function studies in mouse colon cancer models and human cancer cells support a conserved antitumor function for ATOH1 mediated by JNK and p21.

ATOH1 Activates RTK Expression in Human Cancer Cells

How does ATOH1 expression lead to JNK activation in cancer cells? ATOH1 and its orthologs are known to exert their developmental functions by modulating Notch; Atonal orthologs inhibit Notch signaling to induce differentiation during development, whereas Notch inhibits ato expression by its target gene Hes1 (ENSG00000114315) [27–29]. Notch signaling has also been described to be upstream of JNK [30]. However, three lines of evidence suggest that the Atoh1-mediated effects are not Notch-dependent. First, Atoh1 expression levels do not influence expression of HES1, a Notch signaling target gene (Figure S11A). Second, we do not detect cleaved intracellular Notch in the MCC14.2 cell line, which expresses low levels of ATOH1 (Figure S11B). Finally, blocking of Notch activation by selective inhibition of γ-secretase [31] has no effect on the growth of MCC cells (Figure S11C). Other known upstream activators of JNK are RTKs [32]. As Atoh1 is a transcription factor, we checked mRNA expression of all 90 human RTKs upon Atoh1 expression in both the MCC14.2 and the Ht29 cell line compared to control cells. We observed a significant and specific Atoh1-dependent up-regulation of Neurotrophic tyrosine kinase receptor type 1 (NTRK1: ENSG00000198400), a hallmark for differentiation in Merkel cells [33,34] in MCC14.2, and of FGF receptors in Ht29 (Figure S11D). The increase in NTRK1 expression in MCC14.2 seen on the mRNA level was confirmed using RT-qPCR (Figure S10E). We also observed elevated NTRK1 levels in the endogenously ATOH1-expressing MCC1 cells (Figure S10E). This was accompanied by higher protein levels of both the NTRK1 receptor and one of its ligands, Neurotrophin-3 (NT3: ENSG00000185652; Figure S10E–S10H), also a marker for Merkel cell differentiation. Therefore, Ato gain of function results in tumor type–specific elevation of RTK levels.

ATOH1 Functions by RTK-Mediated JNK Activation

To test whether the RTKs may be functionally linked with ATOH1, we incubated the various MCC cell lines with K252a, a narrow-specificity RTK inhibitor (NTRK, FGFR, PDGFR, and IGFR) [35–37]. This results in a dose-dependent decrease in doubling time of the MCC14.2-derived, Atoh1-expressing cell lines, indicating that the Atoh1-induced change in doubling time is RTK-dependent (Figure 7A). In addition, the induction of apoptosis by Atoh1 in the MCC14.2 and the Ht29 cell line is blocked by RTK inhibition (Figure 7B and 7C). These effects are accompanied by RTK inhibitor–dependent decrease in the expression of p21^waf1 and p-JNK (Figure 7D), suggesting that both p21^waf1 and p-JNK are downstream effectors of Atoh1-induced RTK signaling. Next, we asked whether p-JNK is required for p21^waf1 up-regulation and apoptosis by treating MCC cells with SAPK Inhibitor II, a specific JNK inhibitor [38]. We observed a decrease in the Atoh1-induced expression of p21^waf1 and cleaved caspase-3 (Figure 7E). Similarly, transfection of dominant-negative c-jun (TAM67) [39] leads to a significant increase in cell number compared to mock-transfected cells (t-test: p = 0.04), specifically in the MCC14.2-Atoh1.2a, which has high Atoh1 expression levels (Figure 7F).

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Figure 7. Molecular Mechanism for ATOH1 Function

(A–D) show that ATOH1′s anti-oncogenic function acts through receptor tyrosine kinases. (A) Doubling times in hours of MCC14.2 and MCC14.2 transduced with eGFP (lane 2) or Atoh1-IRES-eGFP (lanes 3 and 4) with increasing concentrations of K252a. (B) AnnexinV-positive signals normalized to transfected MCC14.2 cells (GFP). Cells transduced with eGFP (lanes 1 and 2) and Atoh1-IRES-eGFP (lanes 3 and 4) with 0.33 μM K252a (lanes 2 and 4) or DMSO as a control (lanes 1 and 3). The double asterisks (**) indicate significant difference from GFP transfected without K252a (t-test: p < 0.05). (C) AnnexinV-positive signals normalized to transfected Ht29 cells (GFP). Cells transfected with eGFP (lanes 1 and 2) and Atoh1-IRES-eGFP (lanes 3 and 4) with 0.33 μM K252a (lanes 2 and 4) or DMSO as control (lanes 1 and 3). The double asterisks (**) indicate significant difference from GFP transfected without K252a (t-test: p < 0.05). (D) Inhibition of RTKs inhibits p21^waf1 expression and JNK phosphorylation. Western blot analysis for p21^waf1 and pJNK of MCC14.2 (lanes 1 and 5), MCC14.2 transduced with eGFP (lanes 2 and 6), and two MCC14.2-derived cell lines transduced with Atoh1-IRES-eGFP (lanes 3, 4, 7, and 8) with 0.3 μM K252a (lanes 1–4) and with DMSO as a control (lanes 5–8). Actin loading controls are shown below each blot. Quantifications are shown in Figure S12U and S12V.

(E) JNK regulates apoptosis and p21^waf1 expression. Western blot analysis for p21^waf1 and cleaved caspase-3 of MCC14.2 (lanes 1 and 5), MCC14.2 transduced with eGFP (lanes 2 and 6), and two MCC14.2-derived cell lines transduced with Atoh1-IRES-eGFP (lanes 3, 4, 7, and 8) with 1 μM JNK inhibitor (lanes 1–4) and with DMSO as a control (lanes 5–8). Actin loading control is shown below the blots. Phospho-JNK (P-JNK) blot shows the effect of the JNK inhibitor on JNK phosphorylation status Quantifications are shown in Figure S12W–S12Y.

(F) Relative cell numbers (the number of Tam67-transfected cells divided by the number of mock-transfected cells) for MCC14.2 and MCC14.2 transduced with eGFP (lane 2) or Atoh1-IRES-eGFP (lanes 3 and 4) with dominant-negative c-jun (TAM67).

https://doi.org/10.1371/journal.pbio.1000039.g007

In summary, taken together, evidence from mutation analysis in human patients, as well as gain- and loss-of-function analysis in mouse and human cells, support a model (Figure 8) in which ATOH1 modulates JNK activity, possibly via co-option of context-specific RTK signaling, to induce apoptosis and up-regulate p21^waf1 expression, keeping tumor growth in check. Loss-of-function mutations in ATOH1 prevent JNK-mediated apoptosis and p21-mediated cell cycle arrest, leading to enhanced tumor progression.

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Figure 8. Schematic Representation of the Potential Mechanism of ATOH1′s Function as a Tumor Suppressor

In preneoplastic tissue, Atoh1 keeps malignant transformation in check in a JNK-dependent mechanism by the induction of apoptosis and the inhibition of cell cycle progression. When Atoh1 is lost due to deletion or methylation, these brakes on oncogenesis fail, and malignant transformation can progress.

https://doi.org/10.1371/journal.pbio.1000039.g008

Cloning of ATOH1^ERD expression construct.

The ATOH1 open reading frame was PCR amplified and fused in frame with the engrailed repressor domain after XhoI restriction digest. This fusion product was then blunt ligated in the MSCV-IRES-GFP vector (Clontech), giving rise to the MSCV-ATOH1^ERD-IRES-GFP construct.

Mouse models and treatments.

Atoh1^Δintestine mice are a conditional deletion of Atoh1 and are described in more detail elsewhere [3,45]. These mice were generated using the loxP/cre system in which Cre-mediated deletion of Atoh1 is mosaic and is restricted to the distal ileum and large intestine (80%–90% deletion). APC^min mice were purchased from The Jackson Laboratory and mated with Atoh1^Δintestine mice to generate APC^min; Atoh1^Δintestine mice. Eight-week-old male Atoh1^wt and Atoh1^Δintestine littermates were injected intraperitoneally with AOM (Midwest Research Institute of the National Cancer Institute's chemical carcinogen repository) at 10 mg/kg body weight. AOM was injected weekly for a total of six injections, and the mice were sacrificed 20 wk after the first AOM injection. Two hours prior to sacrifice, mice were injected with 50 mg/kg BrdU. The large intestine was isolated and flushed with PBS and then fixed in 10% buffered formalin at room temperature for 16–24 h. Colons were placed in 70% ethanol before macroscopic analysis of polyps, and then embedded into paraffin blocks. Similarly, 16–19-wk-old APC^min and APC^min; Atoh1^Δintestine colons were isolated and fixed as for the AOM-treated colons. For macroscopic analysis, the position and diameter (millimeters) of each polyp were recorded using a dissecting microscope. The protocol for use of animals was approved by the Cincinnati Children's Hospital Institutional Animal Care and Use Committee.

Immunohistochemistry and microscopic analysis.

The colons were imbedded in paraffin blocks and sectioned for hematoxylin and eosin staining by the University of Cincinnati pathology core facility. Tumor phenotypes were determined as in Boivin et al. (2003) [46]. Paraffin-embedded colons of AOM-treated mice were sectioned at 5 μm and used for BrdU and cleaved caspase-3 staining. Mouse anti-BrdU antibodies were obtained from the Developmental Studies Hybridoma Bank maintained by the Department of Biological Sciences of the University of Iowa. The sections were de-paraffinized, rehydrated, and antigen retrieval was performed in citric acid buffer (pH = 6) using a microwave. Endogenous peroxidase activity was blocked with hydrogen peroxide/methanol solution, and Avidin/Biotin block was performed according to the manufacturer's recommendations (Vector Laboratories). Endogenous immunoglobulins block and primary and secondary antibody incubations were performed using the M.O.M. kit following the manufacturer's recommendations (Vector Laboratories). Anti-BrdU antibody was incubated for 16 h at 4 °C in a humidified chamber. Color was developed using the DAB peroxidase substrate kit (Vector Laboratories) followed with hematoxylin staining and dehydration of the tissues. Nuclei from well-oriented crypts in Atoh1^wt and Atoh1^Δintestine colons were counted at 40× magnification, followed by counts of BrdU-positive cells. The percent of BrdU-positive Atoh1^wt or Atoh1-null cells was determined for each animal (at least 1,000 cells per genotype were counted for each animal). Student two-tailed t-test was performed to measure significance. Cleaved caspase-3 staining and counting were performed similarly to BrdU staining and analysis. Polyclonal rabbit anti–cleaved caspase-3 antibodies (1:100) were obtained from Cell Signaling Technology. The Rabbit IgG VECTASTAIN ABC kit was used according to the manufacturer's recommendations for blocking, antibody incubations, and signal amplification (Vector Laboratories).

Crypt preps.

Crypts from wild-type and Atoh1^Δintestine mice were isolated using a modification of the Evans method [47,48]. Mice were sacrificed and the colons removed and flushed with ice-cold PBS, opened flat, and then placed in cold PBS containing protease and phosphatase inhibitors. After a brief vortexing, the colon was cut into four pieces and placed into a 15-ml tube containing shaking solution (1.5 mM KCL, 96 mM NaCl, 27 mM Na Citrate, 8 mM KH₂PO₄, 5.6 mM Na₂HPO₄, 15 mM EDTA, and 1 mM dithiothreitol supplemented with protease and phosphatase inhibitors). The tubes were agitated at 4 °C on a vortexer holding the tubes at 180° until the solution appeared cloudy (5–8 min). The colon pieces were transferred into new tubes with shaking buffer and agitated until most of the crypts were released (enrichment was assessed by phase contrast microscopy at different time points). The solution containing the crypts was filtered through a 100-μm cell strainer to isolate the crypts, and sorbitol (2% final concentration) was slowly added and mixed immediately. The solution was centrifuged at 160g for 8 min at 4 °C. The supernatant (containing single cells) was removed, and the pellet (containing purified crypts) was snap frozen in liquid nitrogen, to be used for protein and RNA analyses.

Quantitative reverse-transcriptase polymerase chain reaction.

RNA was purified from colon crypts isolated from wild-type and Atoh1^Δintestine mice using Trizol (Invitrogen) according to the manufacturer's recommendations. Trizol-purified RNA (100 μg) was subjected to DNase digestion and further purification (RNeasy Mini; Qiagen); 2 μg of total RNA was reverse transcribed (Superscript III, Invitrogen), and cDNA equivalent to 100 ng of RNA used for SYBR Green–based real-time PCR using an Mx3005 (Stratagene). For each gene assessed, colon crypt RNA from nine wild-type and eight Atoh1^Δintestine mice was compared using the standard curve method of relative quantification. All gene expression levels were normalized to the expression of GAPDH. t-Tests measured significant differences between the average normalized expression levels in wild-type versus Atoh1^Δintestine mice.

Whole-tissue/polyp preparations.

Cecal and colonic tissues from wild-type and Atoh1^Δintestine mice were homogenized in lysis buffer (1× Phosphate Buffered Saline, 1% NP-40, 0.5% sodium deoxycholate, 0.1% SDS, 0.7 mM EDTA supplemented with protease and phosphatase inhibitors), sonicated, and the lysates used for quantitation and western blots. Similarly, lysates of colonic tissues and polyps from APC^min, and APC^min; Atoh1^Δintestine double-mutant mice were used for whole lysate preparation and western blots. The following antibodies were used for western blot analysis: p21^waf1 mouse monoclonal antibody (1:500, cat #556431; BD Pharmingen), actin mouse IgM antibodies (1:100, JLA20; Developmental Studies Hybridoma Bank), mouse monoclonal p27^kip1 antibody (1:250, cat #610241; BD Transduction Laboratories), goat anti-p57 polyclonal antibodies (1:100, cat # sc-1039; Santa Cruz Biotechnology), rabbit polyclonal c-Jun antibodies (1:1000, cat # sc-1694; Santa Cruz Biotechnology), and pJNK antibody (1:1000, cat #559309; EMD-Calbiochem).

pJNK IHC.

Formalin-fixed tissues from AOM-treated mice were used for phosphorylated-JNK (pJNK) immunohistochemistry. Rabbit polyclonal pJNK antibody was used (1:100, cat #559309; EMD-Calbiochem). The sections were de-paraffinized, hydrated, and the antigen retrieval was performed in citric acid buffer using a microwave. Endogenous peroxidase activity was blocked with hydrogen peroxide/methanol solution, a short remobilization step was included (0.2% Triton X 100 in PBS), and Avidin/Biotin block was performed according to the manufacturer's recommendations (Vector Laboratories). The Rabbit IgG VECTASTAIN ABC kit was used according to the manufacturer's recommendations for blocking and antibody incubations (Vector Laboratories). Primary antibody was incubated at 4 °C for 14–16 h; color was developed using the DAB peroxidase substrate kit (Vector Laboratories) followed by hematoxylin staining and coverslipping. Cells of well-oriented crypts in wild-type and Atoh1^Δintestine colons were counted at 40× magnifications, followed by counts of pJNK-positive cells. The total number of cells and pJNK-positive cell numbers were added for each animal, and the average numbers were compared across crypts and animals. A Student t-test was performed to measure significance.

Cell culture.

MCC cell lines were cultured in RPMI medium supplemented with 15% FCS (Perbio). The Ht29 cell lines (obtained from Deutsche Sammlung von Mikroorganismen und Zellkulturen [DSMZ]) were cultured in McCoy medium supplemented with 10% FCS. Primary human keratinocytes were isolated and pooled from foreskins of three different donors (less than 6 y). Fourth passage cells were used in the experiments. The procedure has been approved by the ethical committee of the University of Leuven. Experiments performed adhered to the Declaration of Helsinki Principles. Keratinocytes were seeded in serum-free and growth factor–containing medium (Keratinocyte-SFM; Invitrogen), which contains several growth factors (5 μg/ml insulin, 74 ng/ml hydrocortisone, 6.7 ng/ml triiodo-l-thyronine, 50 μg/ml bovine pituitary extract, and 5 ng/ml human recombinant EGF). All culture experiments were done at 37 °C in 5% CO₂.

Doubling time.

A fixed amount of cells was seeded in standard culture conditions, and the number of cells was counted after 3 to 5 d. The doubling times (T^2X) were calculated using the formula T^2X = LN(2)/((LN(n_Δt)-LN(n₀))/Δt) with Δt, time in culture, n₀, number of seeded cells, and n_Δt the number of cells after Δt

Lentiviral vectors.

The lentiviral vector HIV-CMV-Atoh1-IRES-GFP was constructed by cloning a PCR fragment of Atoh1-IRES into the HIV-CMV-GFP vector [49], which was used as control vector. The HIV-CMV-GFP vector was first restricted with XbaI and Age1, and ligated to the Atoh1-IRES fragment, which was spanned between Xba1 and Age1. The Atoh1 and GFP were expressed from a bicistronic vector under the control of the CMV promoter. These lentiviral vectors were produced as described in [50,51]. Stable cell lines were created by transducing the cells in normal medium, supplemented with 8 nM polybrene and a multiplicity of infection between 100 and 150). Four stable cell lines were made with the lentiviral vectors expressing Atoh1 and eGFP from a bicistronic Atoh1-IRES-eGFP construct (MCC14.2-Atoh1.1a, MCC14.2-Atoh1.1b, MCC14.2-Atoh1.2a, and MCC14.2-Atoh1.2b), and one with eGFP alone (MCC14.2-GFP) as a negative control, under the control of a CMV promoter (Figure 5A). We failed to created stable cell lines from the Ht29. Cells were heavily selected against, and FACS isolation did not yield surviving cells.

Colony formation in soft agar.

A total of 2,500 cells/ml were resuspended in 0.6% agarose (Invitrogen) in culture medium. A 2-ml layer of 0.35% agarose in culture medium was added on top of the 2-ml 0.6% layer. After 1 wk, 2 ml of 0.35% agarose in culture medium was added. Cells were cultured for 2 wk in standard conditions, and the number of colonies was analyzed under an inverted microscope with a 10× magnification. The experiment was done in triplicate.

Cell cycle distribution and progression.

Cells were grown up to 40% confluency under standard conditions and were trypsinized. Cells were fixed and permeabilized using 70% ice-cold ethanol for 2 h. Cells were washed in PBS, and cells were stained for 30 min using 0.1% (v/v) Triton X-100 (Sigma) in PBS and 0.2 mg/ml DNase-free Rnase A (Sigma) and 20 μg/ml PI (Sigma). Cells were analyzed on a FACSCalibur cytometer (BectonDickinson). To analyze the cell cycle progression, cells were pulsed for 20 min with 10 μM BrdU (Sigma). Medium was then substituted with normal medium for 6 h. Cells were collected and fixed in 70% ice-cold ethanol. DNA was denatured using 2 M HCl for 20 min. Cells were stained using anti-BrdU and IgG-alexa555 and analyzed in a FACSCalibur (BD Bioscience).

Apoptosis detection.

Annexin-V staining was performed using Annexin-V-biotin (Roche) and streptavidin-PercP (BD biosciences) as described by the manufacturer. Cells were imaged with the Leica TCS Sp2 confocal microscope, and images were analyzed and quantified using LCS software.

Western blotting.

Lysates were obtained by scraping cells in medium and centrifuging the medium. The pellet was resuspended in lysis buffer containing PBS with 1 mM EDTA, 1 mM EGTA, 50 mM NaF, 1% TritonX, 5 mM Na₃VO₄, 20 μM PAO, and Complete Protease Inhibitor. The crude extract was separated by SDS-PAGE in a 4%–12% NuPAGE novex bis-tris gel (Invitrogen) and electroblotted onto Hybond-ECL membrane (Amersham). Antibodies were diluted in the appropriate concentrations in 5% BSA in TBS-tween20. The antibodies used are beta-actin (clone AC-15; Sigma), anti–cleaved caspase-3 (Asp-175; Cell Signaling Technology), polyclonal antibody to caspase-9(active) (ALEXIS), chromogranin A+B (Abcam), cyclinA1 (Santa Cruz Biotechnology), pJNK (Thr183/Tyr185; BioSource), p27 (BD Bioscience), p21^waf1 (DCS60; Cell Signaling Technology), PCNA (clone PC10; Sigma), TRK (Santa Cruz Biotechnology), Atoh1 (1/50; Developmental Studies Hybridoma Bank), NT3 (Santa Cruz Biotechnology), and Cleaved Notch (Val1744; Cell Signaling Technology). All western blot analyses are quantified in Figure S11.

Tyrosine kinase inhibitor, SAPK inhibitor II, and γ-secretase inhibitor.

K252a diluted in DMSO, from VWR, and DMSO as control were used. Final concentrations ranged from 0 μM to 0.5 μM; when no concentration is mentioned, 0.33 μM was used. SAPK inhibitor II, from Calbiochem, was used at a concentration of 10 μM. We used inhibitor X as a γ-secretase inhibitor (Calbiochem). Concentrations used are mentioned in the figure legend.

RT-PCR and qPCR.

mRNA was amplified using the Superscript II One-Step RT-PCR system with Platinum Taq-100 reactions (Invitrogen) and the ATOH1 primers CAGCCAGTGCAGGAGGAAAA and GAAAATTCCCCGTCGCTTCT, and the Hes1 primers GGACATTCTGGAAATGACAGTGAA and AGCGCAGCCGTCATCTG, according to the manufacturer's protocol. Quantitative RT-PCR was performed on a ABI prism 7000 (Applied Biosystems). The primers were designed using Primer Express (Applied Biosystems). The primers used are ATOH1: CAGCCAGTGCAGGAGGAAAA and GAAAATTCCCCGTCGCTTCT; Atoh1: GCTGTGCAAGCTGAAGGG and TCTTGTCGTTGTTGAAGG. Primers to check copy number of the ATOH1 locus: ATOH1 locus set 1: CCCCGGGAGCAATCTTG and GGGACCGAGGCGAAGTT; control locus set 1: TCTGGGACCTGAGCTAATGGA and GGCCATAATTAGGACCATGAAAGA; and ATOH1 locus set 2: GCCAGTGCAGGAGGAAAACA and GAAAATTCCCCGTCGCTTCT. Control locus set 2: GGGTTCAGCCTCAACTTGTATCC and CCCACCACCTGGCATCTCT.

The 90 kinase RT-PCR assay.

RNA was diluted to 300 μg/μl and treated with TURBO DNA-free (Ambion) following the manufacturer's protocol. cDNA was synthesized using random primers and SuperscriptII reverse transcription (Invitrogen) using the manufacturer's protocol. cDNA was amplified using gene-specific primers (concentration 5 μM) for the different human tyrosine kinases.

Methylation detection.

Methylation of the DNA was detected using ApaI enzyme. DNA (2 μg) was dissolved in 50 μl of the appropriate buffer and 10 U of ApaI, and incubated at 30 °C overnight. A mirror condition was done in which the ApaI was exchanged by glycerol as a control for unspecific degradation. The resulting DNA was analyzed using two primer sets: one set spanning the restriction site (AATAAGACGTTGCAGAAGAG and TCGCAGAGCAAAAATTAAAGGGTGC) and another set next to the restriction site (CCCCGGGAGCATCTTGCAGCCA and TCGCAGAGCAAAAATTAAAGGGTGC).

Pull-down of methylated DNA fragments (restricted using EcoRI) was performed using the Methylcollector kit (Active Motif), and bisulfite modification of DNA was done using the EZ DNA Methylation-Gold kit (Zymo Research) according to the manufacturers' protocols.

Array CGH.

We carried out array CGH using Code Linked Slides (AP Biotech) containing the 3,527 BAC clones from the Wellcome Trust Sanger Institute 1 Mb Clone Set, a gift from N. P. Carter (The Wellcome Trust Sanger Institute). Array CGH was performed as described [17].

Sequencing.

The ATOH1 open reading frame was amplified with PCR using AATAAGACGTTGCAGAAGAG and TCGCAGAGCAAAAATTAAAGGGTGC and AmpliTaq Gold DNA polymerase and the GeneAmp PCR System 2400 (Applied Biosystems). The PCR products were purified and sequenced in both directions on the ABI Prism BigDye (Terminator Cycle Sequencing Kit version 1.1) on an ABI PRISM 3100 Genetic Analyser (Applied Biosystems).