*Chang Y-T, Chaturvedi P, Schock EN and Brugmann SA (2016) Understanding Mechanisms of GLI-Mediated Transcription during Craniofacial Development and Disease Using the Ciliopathic Mutant, talpid2 . Front. Physiol. 7:468. doi: 10.3389/fphys.2016.00468* The primary cilium is a ubiquitous, microtubule-based organelle that cells utilize to transduce molecular signals. Ciliopathies are a group of diseases that are caused by a disruption in the structure or function of the primary cilium. Over 30% of all ciliopathies are primarily defined by their craniofacial phenotypes, which typically include midfacial defects, cleft lip/palate, micrognathia, aglossia, and craniosynostosis. The frequency and severity of craniofacial phenotypes in ciliopathies emphasizes the importance of the cilium during development of the craniofacial complex. Molecularly, many ciliopathic mutants, including the avian *talpid2* (*ta2* ), report pathologically high levels of full-length GLI3 (GLI3FL), which can go on to function as an activator (GLIA), and reduced production of truncated GLI3 (GLI3T), which can go on to function as a repressor (GLIR). These observations suggest that the craniofacial phenotypes of ciliary mutants like *ta2* are caused either by excessive activity of the GLIA or reduced activity of GLIR. To decipher between these two scenarios, we examined GLI3 occupation at the regulatory regions of target genes and subsequent target gene expression. Using *in silico* strategies we identified consensus GLI binding regions (GBRs) in the avian genome and confirmed GLI3 binding to the regulatory regions of its targets by chromatin immunoprecipitation (ChIP). In *ta2* mutants, there was a strikingly low number of GLI3 target genes that had significantly increased expression in facial prominences compared to the control embryo and GLI3 occupancy at GBRs associated with target genes was largely reduced. *In vitro* DNA binding assays, further supported ChIP results, indicated that the excessive GLI3FL generated in *ta2* mutants did not bind to GBRs. In light of these results, we explored the possibility of GLI co-regulator proteins playing a role in regulatory mechanism of GLI-mediated transcription. Taken together our studies suggest that craniofacial ciliopathic phenotypes are produced via reduced GLIT production, allowing for target gene transcription to be mediated by the combinatorial code of GLI co-regulators.

Keywords: primary cilia, craniofacial, talpid2 , c2cd3, ciliopathies, GLI

# INTRODUCTION

Primary cilia are ubiquitous organelles that serve as cellular hubs for transduction of numerous signaling pathways. Most notably, cilia have been identified as transducers of the Hedgehog (Hh) pathway. Identification of the primary cilium as a signaling hub for the Hh pathway came from seminal experiments reporting that anterograde and retrograde intraflagellar transport (IFT) proteins in the cilium are required for Sonic Hedgehog (SHH) signal propagation (Huangfu et al., 2003; Huangfu and Anderson, 2005). When SHH ligand is present, it binds to its receptor Patched (PTCH), thus allowing Smoothened (SMO) to localize and accumulate in the primary cilium (Corbit et al., 2005; Rohatgi et al., 2007). Activated, ciliary SMO, in concert with Kif7, then promotes the dissociation of GLI from Suppressor of Fused (SUFU) (Humke et al., 2010; Tukachinsky et al., 2010; Li et al., 2012) and the subsequent post-translational processing of GLI proteins necessary for their function as activators and repressors (Goetz and Anderson, 2010).

In vertebrates, there are three members of the GLI transcription factor family: GLI1, GLI2, and GLI3. GLI1 and GLI2 are considered transcriptional activators, whereas GLI3 mostly behaves as a repressor. However, there have been examples of GLI2 functioning as a repressor in the absence of GLI3, and GLI3 functioning as an activator in the absence of GLI2 (Mo et al., 1997; Theil et al., 1999; Tole et al., 2000; Bai and Joyner, 2001; Persson et al., 2002; Rallu et al., 2002; Buttitta et al., 2003; Motoyama et al., 2003; Bai et al., 2004; Lei et al., 2004; McDermott et al., 2005; Pan et al., 2009). Full-length GLI2 and GLI3 can be processed via phosphorylation and other post-translational modifications into the activator isoform (GLIA) or truncated into the repressor isoform (GLIR) (Wang et al., 2000; Pan et al., 2006). Inhibition of GLI processing prevents production of GLIA and GLIR isoforms. Thus, an essential role of primary cilia is to establish the ratio of GLIA to GLIR proteins (Haycraft et al., 2005; Liu et al., 2005), which in turn controls transcription of SHH target genes.

Three basic models have been proposed to depict the potential mechanism of how SHH target genes are activated by a gradient of GLI isoforms: (1) the ratio sensing model, (2) the threshold repression model and (3) the threshold activation model (Falkenstein and Vokes, 2014). Ratio sensing, as the name implies, is based on the ratio of GLIA and GLIR rather than concentration of either. The net balance of GLIA to GLIR then determines if, and the extent to which, a target is activated or repressed. The other two models suggest threshold-specific mechanisms. The threshold activation model suggests that SHH targets are activated when GLIA reaches a threshold-specific concentration. On the other hand, in the threshold repression model (de-repression), the activation of SHH target genes is dictated by the alleviation of repression via loss of GLIR.

The talpid2 (ta2 ) is a naturally occurring avian mutant that is best characterized by severe polydactyly and its oral-facial phenotype (Abbott et al., 1959, 1960; Dvorak and Fallon, 1991; Schneider et al., 1999). The face of affected ta2 embryos is characterized by a dysmorphic frontonasal prominence, facial clefting, hypoplastic maxillary prominences, incomplete fusion of the primary palate and hypoglossia (Chang et al., 2014). Our recent work determined that the ta2 mutation affected ciliogenesis via a 19 bp deletion in C2CD3 (Brugmann et al., 2010; Chang et al., 2014), a centriolar protein required for ciliogenesis (Hoover et al., 2008). Our studies also determined that the ta2 mutant genetically, biochemically and phenotypically phenocopied the human craniofacial ciliopathy, Oral-facialdigital syndrome 14 (OFD14) (Schock et al., 2015). ta2 embryos, similar to many other ciliopathies, have a significant increase in GLI3FL and a reduction in the amount GLIT (Chang et al., 2014). However, the mechanism by which this disruption in GLI isoform production affects expression of GLI targets in the developing craniofacial complex remains unknown.

Herein, we utilize a combination of several biochemical techniques to determine the impact loss of cilia has on GLI function. Specifically, we examine the expression of GLI target genes and occupation of GLI binding regions (GBRs) associated with those targets in the developing frontonasal, maxillary and mandibular prominences (FNP, MXP, and MNP, respectively) in order to uncover the mechanism by which GLI mediated transcription is being impacted in ta2 mutants. Understanding the full extent of molecular disruptions in ta2 mutants will hopefully guide future therapeutic strategies for craniofacial ciliopathies, a rapidly growing group of disorders that currently have little to no therapeutic treatment.

# MATERIALS AND METHODS

# Embryo Preparation

talpid2 (ta2 ) heterozygous carriers were mated, eggs were collected and shipped from the UC Davis Avian Sciences Department. Embryos were incubated at 37◦ C for approximately 5–7 days when embryos reached Hamburger Hamilton stage 25–31 (HH25-31).

# Quantitative RT-PCR of GLI Targets

FNPs, MXPs, and MNPs were harvested from day 5 chick embryos. mRNA was prepared with TRIzol reagent (Thermo Fisher Scientific), and then converted to complementary DNA through reverse transcription reaction (High-Capacity cDNA Reverse Transcription Kit, Applied BiosystemsTM ). Different amounts of cDNA (40, 20, 10 and 5 ng) was used for quantitative PCR to test PCR efficiency and a linear range of duplication (SsoAdvancedTM Universal SYBR R Green Supermix, BioRad). Expression of genes known to play a role in craniofacial development were examined with the following primer sets: ALX4 (105 bp) F: GTTACGGTAAGGAGAGCAGTTT, R:CTTTCACTCCAGCCTCCTTC, BMPR1A (100 bp) F: GTGCTGTCGGACTGATTTCT, R:TGCCATCCAACGAAT GCT, WIF1 (100 bp) F: CAACCTGTTTCAATGGAGGAAC, R: GGCTGATGGCATTTACTGATTT, OSR2 (140 pb) F: CCACTTCACCAAGTCCTACAA, R: TCTCTTTGGAATGGAT GTACCG. The statistical significance of the data was evaluated through two-tailed Student's t-test. p-values less than or equal to 0.05 (95% confidence level) were considered as statistically significant differences.

# Western Blotting of GLI2/3 Proteins

FNPs, MXPs, or MNPs were pooled and lysed in RIPA buffer containing protease inhibitors and phosphatase inhibitors (1 mM Phenylmethylsulfonyl fluoride, 1 mM NaVO4, 1X complete protease inhibitor cocktail, EDTA-free), and slightly sonicated with a microprobe to recover chromosome bound GLI2 and GLI3 proteins. BCA assay (Pierce) was used to measure protein concentration of cell lysates. Proteins were boiled with 1X Laemmli sample buffer and run on 6% SDS-PAGE for GLI2 and GLI3, or 12% SDS-PAGE for GAPDH, which later were wet-transferred to Polyvinylidene difluoride (PVDF) membrane. Anti-GLI2, anti-GLI3 (Polyclonal goat IgG 1:500, R&D systems) and anti-GAPDH (FL-335, polyclonal rabbit IgG 1:4000, Santa Cruz Biotechnology) were prepared in 1X TBST (0.1% Tween-20)/6% nonfat milk, as well as secondary antibodies (antigoat and anti-rabbit, 1:10,000). Proteins were detected by Electrochemiluminescence assay (Amersham ECL Prime, GE Healthcare Life Science).

# DNA Binding Affinity Assay

PATCHED 1 promoter oligonucleotides were designed according to the location of a Gli binding region (GBR) at −2549 from the TSS site (GGAAGAAGTGTCAGTGTAAG AGTCTCCACGTGGGTGGTCAAGGCCATGGCTGCCTCAC GG). 100 pmole of biotin-conjugated positive oligonucleotides and complementary oligonucleotides (Integrated DNA Technologies) were annealed in 1X TE/50 mM NaCl buffer in a PCR cycler and incubated with Dynabeads Streptavidin (M280, Invitrogen). FNPs, MXPs, or MNPs from day 5 control or ta2 embryos were pooled and processed as described for Western blotting. Cell lysates were incubated with oligonucleotidesbound Dynabeads at 4◦C for 2 h. Beads were washed with 1 ml RIPA buffer three times and processed for Western Blotting analysis.

# Chromatin Immunoprecipitation

FNPs, MXPs, or MNPs from day 5 control or ta2 embryos were harvested, pooled and crosslinked in 1% formaldehyde. Tissue was homogenized in RIPA buffer and sonicated (Bioruptor R Pico, Diagenode) at 5 cycles of 30 s on/45 s off. Sheared DNA was distributed around 0.3 kb to 1 kb on 1% agarose gel. Cell lysates were pre-cleaned with Dynabeads protein G (ThermoFisher Scientific) and quantified by BCA assay. Dynabeads for immunoprecipitation were blocked with 20µg/ml Glycogen, 20µg/ml BSA, 20µg/ml yeast RNA in RIPA buffer at 4◦C for an hour. GLI3 antibody (AF3690, R&D systems) and pre-blocked Dynabeads Protein G were incubated with 90% of cell lysates at 4◦C overnight. 10% of cell lysates were kept as Input. Beads were washed with IP wash buffer I (low salt; 50 mM HEPES-KOH pH 7.5, 150 mM NaCl, 1 mM EDTA, 0.1% sodium deoxycholate, 1% Triton X-100), IP wash buffer II (high salt; 50 mM HEPES-KOH pH 7.5, 500 mM NaCl, 1 mM EDTA, 0.1% sodium deoxycholate, 1% Triton X-100), IP Wash Buffer III (LiCl containing buffer; 10 mM Tris-Cl pH 8.0, 250 mM LiCl, 1 mM EDTA, 0.5% Sodium deoxycholate, 0.5% NP-40) and TE buffer (10 mM Tris-HCl pH 8.0, 1 mM EDTA). ChIP samples were reverse-crosslinked by boiling with 10% Chelex-100 (BioRad), and treated with 0.2 mg/ml Proteinase K at 55◦C for 30 min. Immunoprecipitated DNA samples were analyzed with quantitative real-time PCR (BioRad) with primers to GBR of target genes. Error bars in all figures represent standard error of the mean (S.E.M.) from five to seven independent experiments. The statistical significance of the data was evaluated through two-tailed Student's t-test. p-values less than or equal to 0.05 (95% confidence level) were considered as statistically significant differences.

# GBR Analysis

Sequences for GBRs from previous publications (Vokes et al., 2007, 2008) were used with a custom perl script to search for all the exact matches of the various possible sequences of the consensus GBRs in the chicken genome. Acquired positions of the motifs in the genome were run through a second custom perl script to search for genes that encompass these motif sites at a distance of 20 kb from either end. Potential GLI targets were confirmed using chromatin-immunoprecipitation (ChIP) assays.

# RESULTS

### The Avian Ciliopathic Mutant, talpid2 , has Craniofacial Anomalies Characteristic of a Ciliopathy

To understand the transcriptional networks affected in craniofacial ciliopathies we analyzed the talpid2 (ta2 ) mutant, a naturally occurring avian ciliopathic mutant that has been established as a model for the human craniofacial ciliopathy, Oral-facial-digital syndrome 14 (Chang et al., 2014; Schock et al., 2015). The ta2 craniofacial phenotype characterized by facial and palatal clefting, micrognathia, and hypoglossia, is fully evident at day 7 (**Figure 1**). Although our previous work has characterized this phenotype (Chang et al., 2014; Schock et al., 2015), to determine the transcriptional networks that contribute, we first needed to identify when phenotypic onset occurred. At day 7 the frontonasal prominence (FNP) is shorter and wider and frequently does not fuse to adjacent prominences (**Figures 1A–B'**). Two days earlier at day 5, the MXPs were medially rotated, the nasal pits were larger, thus preventing the proper juxtapositioning of the FNP with adjacent prominences (**Figures 1C–D'**). Palatal views showed increased patency of the naturally cleft avian secondary palate in ta2 embryos relative to controls at day 7 (**Figures 1E–F'**, dotted white lines). Two days earlier, at day 5, the dysmorphology and malposition of the MXPs is just becoming apparent (**Figures 1G–H'**). Dorsal views of the developing lower beak showed the MNP of ta2 embryos failed to fuse completely and had a hypoplastic tongue (hypoglossia, dotted black line) (**Figures 1I–J'**, white arrow). At day 5 there was little difference in mandibular growth between control and ta2 MNPs (**Figures 1K–L'**). From phenotypic evaluations, taken together with the fact that these anomalies were not readily identifiable at day 4, we determined that craniofacial anomalies were initiated at approximately day 5 of development. Thus, to determine the molecular networks responsible for these

phenotypes we carried out our analyses in the facial prominences of day 5 embryos.

# Loss of Cilia Results in Aberrant GLI Isoform Production

Our previous work determined that aberrant ciliogenesis in ta2 embryos disrupted the production of GLI proteins in such a manner that there was increased GLIFL production and decreased GLIT production (Chang et al., 2014). The GLIFL isoform typically goes on to function as an activator, whereas the GLIT isoform goes on the function as a repressor. The excessive production of GLIFL is an extraordinarily common molecular phenotype in ciliopathies, including those with craniofacial phenotypes (Huangfu and Anderson, 2005; Davey et al., 2006; Tran et al., 2008; Tabler et al., 2013). To carefully exam the differences of GLIFL and GLIT protein levels in control and ta2 mutants, we performed Western blot analysis of GLI2 and GLI3 in the three facial prominences affected in the ta2 at day 5. We detected a very low level of GLI2FL and a substantial amount of GLI2T isoforms in the control facial prominences (**Figure 2A**). The loss of cilia in ta2 embryos disrupted GLI processing and altered the production of GLI2 protein isoforms. In ta2 facial prominences, we detected dramatically increased levels of GLI2FL, and low levels of GLI2T, relative to control prominences. We next examined the production of GLI3 isoforms. Western blot analyses showed that, similar to GLI2FL, GLI3FL was increased in ta2 prominences relative to controls. Contrary to what was observed with the GLI2T, GLI3T was readily detectable in both control and ta2 facial prominences. Specifically, ta2 MNP had less GLI3T than control MNP. These data suggested that disrupted ciliogenesis affected the processing of GLI2 and GLI3 in distinct manners, yet the net result was an increase in GLIFL production. We did not observe a change in GLI1 protein levels between control and mutant embryos (Data not shown).

Since the GLIFL isoform typically goes on to act as an activator, these results suggest that craniofacial phenotypes in the ta2 mutant could be caused by increased GLI activator function leading to increased expression of GLI target genes. To test this hypothesis, we utilized an in silico approach to look for potential GLI target genes within the avian genome. Using previously published sequences of GBRs (Vokes et al., 2007, 2008) we scanned the chicken genome for possible GLI targets. GBR positions were run through a custom perl script to search for genes that encompass these motif sites at a distance of 100 kb from the 5′ or 3′ end of the gene. From this list we identified several genes known to play a role in craniofacial development (**Supplemental Table 1**). Confirmation of our in silico approach was carried out on selected genes using ChIP assays (**Supplemental Figures 1 A–D**). Through these analyses, we selected four GLI3 target genes known to be expressed during, and have a role in, craniofacial development: ALX4 (Beverdam et al., 2001; Mavrogiannis et al., 2001), BMPR1A (Li et al., 2011, 2013; Saito et al., 2012), OSR2 (Lan et al., 2001), and WIF1 (Hsieh et al., 1999; Darnell et al., 2007). PTCH1, a known GLI target, was used as a positive control (**Supplemental Figure 1E**) and IgG IP as an antibody background control. The data were all normalized to IgG IP percentage, and the genes with relative enrichment >1 were considered positive for GLI3 binding. To determine if increased production of GLI3FL in ta2 facial prominences correlated with increased expression of these target genes, we performed quantitative RT-PCR (qPCR) with mRNA from facial prominences of day 5 control and ta2 embryos (**Figures 2B–E**). There was not an increase in ALX4 expression in any of the facial prominences (**Figure 2B**); however, we detected a significant decrease in ALX4 expression in the MXP. BMPR1A expression was also not significantly increased in any of the developing prominences (**Figure 2C**). No significant changes in OSR2 expression were detected in the FNP or MNP; however, a significant increase in expression was observed in the MXP (**Figure 2D**). Finally, WIF1 expression was not changed in the FNP, yet was significantly increased in the MXP and significantly decreased in the MNP (**Figure 2E**). Taken together, these data do not support the idea that increased production of GLI3FL directly and uniformly results in increased expression of GLI targets throughout the facial prominences. Additionally, these data suggest that each facial prominence interprets aberrant GLI production in a unique manner.

# Excess Production of GLIFL Does Not Correlate with an Increase of GLIFL Occupancy at GBRs

For GLIFL to function as an activator, it has to occupy the regulatory regions of GLI targets. We wondered if lack of uniform increases of target gene expression in ta2 embryos was due to failure of GLIFL to recognize and occupy GBRs of target genes. To test this hypothesis, we performed an in vitro DNA binding assay using the PATCHED 1 (PTCH1) promoter. We synthesized a 60 base pair biotin-labeled oligonucleotide of the PTCH1 promoter containing an endogenous GLI binding motif found at position −2549 proximally upstream of the transcription start site (TSS). Through high affinity of Biotin-Streptavidin interaction, we were able to evaluate the DNA binding ability of GLI3 isoforms by Western blot. Pre-incubation of nonlabeled PTCH1 oligonucleotides depleted GLI3 protein signals, which confirmed the specificity of the GLI3-GBR interaction. Under the same exposure, the affinity based pull-down assay showed that GLI3T predominantly bound to the biotin-labeled oligonucleotides in both control and ta2 mutants. Interestingly, despite the high level of GLI3FL production in ta2 embryos, GLI3FL failed to bind to the biotin-labeled oligonucleotides (**Figures 3A,B**). On the contrary, comparison between control and ta2 embryos indicated that the amount of GLI3T binding to the oligonucleotides correlates with the protein concentration (**Figures 3A,B**). Taken together these data suggest that despite increased GLI3FL production, target gene expression is not increased via increased GLI activator function because GLI3FL does not occupy the GBRs of target genes. In addition, the predominant binding of the GLI3T repressor at GBRs supports the idea of the threshold repression model, in which GLI targets underwent de-repression due to the removal of GLIT repressor.

FIGURE 2 | Excessive GLIFL production in ta2 embryo does not correlate with increased gene activation of GLI targets. (A) Western blot of GLI2 and GLI3 proteins from the frontonasal prominence (FNP), maxillary prominence (MXP) and mandibular prominence (MNP) of day 5 control and *ta2* embryos. GAPDH was used as a loading control. (B–E) mRNA-qPCR analyses of *ALX4*, *BMPR1A*, *OSR2*, *WIF1* from FNP, MXP, and MNP of day 5 control and *ta2* embryos. The data was normalized by the individual facial prominences of control and *ta2* embryos. The asterisks indicate statistically significant differences and are assigned as followed: \**P* < 0.05, \*\**P* < 0.01. Error bars are based on the standard error of the means (S.E.M.). *n* = 4.

mandibular prominence (MNP) in day 5 control and *ta2* embryos. (B) Western blot of GLI3 proteins from the same lysates as (A). GAPDH as loading control of cell lysates.

# GLI3 Occupancy at GBRs within the Regulatory Regions of Target Genes Is Altered in ta2 Mutants

In vitro DNA binding affinity assays suggested that there was not an increase of ectopic GLI3FL at GBRs of target genes in ta2 mutants. To determine if GLI3 binding to GBRs was altered in vivo in ta2 mutants compared to control embryos, we next performed ChIP-qPCR (**Figure 4**). In ta2 mutants, GLI3 enrichment at GBRs associated with the craniofacial genes ALX4 and OSR2 was reduced in all facial prominences, yet only significantly in the MXP and MNP (**Figures 4A,B**). GLI3 enrichment at the GBRs associated with BMPR1A and WIF1, was also overwhelmingly reduced in facial prominences, yet only significantly in the ta2 FNP and MXP (**Figures 4C,D**). GLI3 binding in the MNP of WIF1 was below detectable levels in controls, and thus could not be evaluated. These data suggested that GLI3 occupancy at the GBR of target genes is decreased. Our in vitro data suggested that GLI3FL did not bind at target gene GBRs (**Figure 3**). Thus, taken together these data indicate that the observed reduction of GLI3 binding is indicative of reduced GLI3T binding at the regulatory regions of target genes.

Reduced GLI3 binding at GBRs of GLI target genes appeared to be a general trend in ta2 mutants, as several other target genes expressed in the craniofacial complex also exhibited reduced enrichment of GLI3 at associated GBRs (**Supplemental Figure 2**). Thus, these data suggested that altered expression of GLI targets could be caused by aberrant GLI3T binding in ta2 mutants. Interestingly, there was not always a direct correlation with loss of GLI3T repressor binding and increased gene expression. Furthermore, different results among the three facial prominences examined pointed to context—and tissue-specific regulation of GLI targets (**Supplemental Figures 1A–D**). Several studies have proposed the possibility that GLI proteins work together with co-regulators to influence expression of targets genes (Brewster et al., 1998; Koyabu et al., 2001; Mizugishi et al., 2001; Lee et al., 2010; Peterson et al., 2012). To determine if target gene expression in the craniofacial complex was influenced by the action of GLI coregulators, we next examined the proximity of GBRs to motifs for potential GLI co-regulators.

# Motif Analyses Identified Sequences for Known GLI Co-Regulators Frequently Co-Localize with GBRs

GBRs have previously been identified in close proximity to binding motifs for other transcription factors, including members of the bHLH, SP, and Sox families (Vokes et al., 2008; Peterson et al., 2012; Aberger and Ruiz I Altaba, 2014). Furthermore, the co-occupancy and cooperativity of GLI with SOX transcription factors was previously shown to be essential for activating neural gene expression signatures (Peterson et al., 2012). To determine if expression of our identified GLI targets could be influenced by the presence or absence of GLI coregulators, we first examined the genomic sequence around the GBRs of our target genes for E-box, SP, and SOX binding motifs. (**Figure 5**, **Supplemental Figure 3**). We defined the area <1 kb upstream of the transcription start site (TSS) as the promoter region, <20 kb away from the TSS as proximal upstream or downstream, and <100 kb away from the TSS as distal upstream or downstream. We analyzed the sequence surrounding GBRs in our four selected GLI target genes for sequences predictive of E-box, SP, and SOX binding. All four of our identified GLI target genes contained at least one motif cluster containing a GBR, E-box, SP, and SOX binding site within 1 kb of each other (**Figures 5A–D**). Several other clusters containing three of the four motifs were also identified (**Figures 5A–C**; red box). The close proximity of these binding motifs suggested that expression of GLI targets in the developing craniofacial complex could be influenced by the cooperative function of GLI isoforms and co-regulator proteins.

# GLI Co-Regulators Have a Prominence Specific Expression Pattern That Changes When Primary Cilia Are Lost

Several transcription factors synergistically cooperate with GLI proteins to influence GLI target gene expression (Aberger and Ruiz I Altaba, 2014). Their co-occupancy at the promoter of GLI targets is required for the optimal activation/repressor. ChIPbased, high-throughput analyses uncovered several transcription factor motifs located close to GBRs in the cis-regulatory modules of GLI targets. Specifically, binding sequences for Sox (Peterson et al., 2012), bHLH (Lee et al., 2010), and SP proteins (Vokes et al., 2008) have been shown to exist in very close proximity

to GBRs. Our in silico analyses confirmed these sequences exist in near GBRs in four previously identified GLI-targets. We hypothesized that differential expression of these co-regulators could contribute to the differential gene expression of GLI targets in both control and ta2 mutant embryos. We first investigated the expression of genes that could bind to motifs found in close proximity to GBRs, specifically SOX8, SP3, and HAND2 (**Figures 6A–C**). SOX8 expression was significantly reduced in the FNP and MXP of ta2 mutants, yet was not significantly altered in the MNP (**Figure 6A**). SP3 expression was significantly increased in ta2 MNP, yet not changed in the FNP and MXP (**Figure 6B**). HAND2 expression was reduced in the MXP, but not significantly changed between control and ta2 FNPs and MNPs (**Figure 6C**). Thus, differential expression of GLI co-regulators could possibly contribute to altered target gene expression in ta2 mutants and explain why changes in GLIFL and GLIT isoforms do not uniformly or directly correlate with changes in target gene expression.

Our previous analyses indicate that GLI target gene expression changed in a prominence specific manner (Chang et al., 2014). We wondered if differential expression of potential GLI co-regulators could contribute to this prominence-specific expression changes. SOX8 was robustly expressed within the FNP, with levels significantly higher than those in the MXP or MNP (**Figure 6A'**). SP3 was more robustly expressed in the MXP and MNP, relative to the FNP (**Figure 6B'**). HAND2 was robustly and exclusively expressed in the MNP (**Figure 6C'**). Thus, each of these potential co-regulators has a prominences specific expression pattern that could differentially influence GLI target gene expression. Collectively, the proximity of binding motifs, coupled with the differential expression of these coregulators could possibly contribute to altered expression of GLI targets in ta2 embryos.

# DISCUSSION

Craniofacial ciliopathies are a rapidly growing group of disorders that severely impact craniofacial growth and development. Currently, there are little to no therapeutic options for these conditions. Although the molecular mechanism behind these disorders remains nebulous, many ciliopathies have aberrant production of GLIFL and GLIT isoforms.  Herein, we attempted to identify the mechanism by which aberrant GLI protein production impacts craniofacial development. To do so we used the avian ta2 model, which has recently been characterized as a bona fide model for the human craniofacial ciliopathy Orofacial-digital syndrome 14 (Schock et al., 2015). In silico and ChIP assays identified GLI target genes in the avian genome that play a role in craniofacial development. RT-qPCR of mRNA levels verified some significant

changes in the expression of these GLI targets in the developing facial prominences, yet there was not a clear, linear relationship between changes in GLI isoform production and target gene expression. Motif cluster analysis supported the hypothesis that GLI proteins work in concert with co-regulators. GBRs associated with GLI target genes were found to be situated within 1000 bp of binding motifs for several, previously identified GLI co-regulators. Differential expression within developing facial prominences of predicted co-regulators supported a hypothesis in which expression of GLI target genes is dependent upon the cooperative function of GLI isoforms and co-regulator proteins.

Together, these data provide a better understanding of the complex nature of GLI-mediated transcription that occurs in normal and ciliopathic craniofacial development.

# GLI Binding Regions Are Present throughout the Avian Genome in Genes That Affect Craniofacial Development

Work in other species has identified GBRs and examined the role of GLI proteins as transcription factors that affect gene expression in numerous signaling pathways (Vokes et al., 2007, 2008).

Three models currently exist to explain GLIFL/GLIT function (Falkenstein and Vokes, 2014). First the GLIFL::GLIT ratio sensing model suggest that the relative levels of GLIFL to GLIT is integrated and results in graded levels of transcription of targets. This model suggests that response to changes in ratio, rather than concentration, affect target gene expression. The threshold activation model suggests that, rather than ratio of GLIFL::GLIT, a threshold-specific concentration of GLIFL activates target gene expression (Oosterveen et al., 2012). The threshold repression model depends on the removal of the GLIT repressor from target genes, allowing for transcriptional activation by other transcription factors to initiate gene expression. From our results, mRNA expression of most target genes was not dramatically altered despite excessive GLIFL production in ta2 mutant, indicating that the ratio sensing model cannot explain our observation in ta2 mutant. Secondly, the accumulated GLI3FL in the nucleus was expected to bind and activate target gene in the threshold activation model; however, instead we found reduced occupancy of GLI3 at target genes, along with lack of detectable GLIFL binding to GBRs (**Figures 3**, **4**), suggesting that a dysfunctional GLIFL that fails to promote GLIA-mediated activation is produced under ciliopathic conditions. Thus, these data do not support the threshold activation model for GLIFL function. Conversely, our results showed that reduced GLIT production could lead to a de-repression of target genes and potentially a gene activation, when the required transcription factors are available. The threshold repression model seems to be more applicable to the observation owing to less GLI3 occupancy and the failure of GLIFL to GLIT conversion. A more in depth understanding of GLI mechanisms of action will require tools with the ability to definitively decipher between GLIFL and GLIT in vivo.

# Excessive GLIFL Production Does Not Equate to Increased GLIA Activity in Ciliary Mutants

A number of disorders identified as ciliopathies have craniofacial abnormalities including Oral-facial-digital syndrome, Joubert syndrome, Bardet-Biedl syndrome, Meckel-Gruber syndrome, Ellis-van Creveld syndrome (Zaghloul and Brugmann, 2011). The phenotypes for syndromes such as these, while not identical, do have several phenotypes indicative of aberrant SHH signaling including widening of the midface, cleft lip/palate, micrognathia, craniosynostosis and oral/dental anomalies (Zaghloul and Brugmann, 2011). Increased production of GLIFL isoforms has been observed in several ciliary mutants (Huangfu and Anderson, 2005; Liu et al., 2005; Humke et al., 2010), thus a common interpretation of these data is that ciliopathic phenotypes were due to either increased activator function or skewed GLIFL::GLIT ratio in favor of GLIFL. Despite these common interpretation regarding the molecular mechanism causing ciliopathic phenotypes, examination of subsequent levels of GLI processing, binding, and transcriptional function in ciliary mutants has not been performed.

Our studies are among the first to examine how and if GLI proteins function in the developing craniofacial complex of ciliary mutants; however, some questions remain regarding the mechanistic reasons as to why GLIFL appears not to function in ciliary mutants. Prior to processing, GLI proteins associate with Suppressor of Fused (SUFU), a conserved protein known to regulate the activity of GLI proteins via modulating GLI processing, stabilization and subcellular localization (Barnfield et al., 2005; Humke et al., 2010; Tukachinsky et al., 2010; Wang et al., 2010). In the presence of a SHH signal, the SUFU-GLIFL complex traffics through the cilium (Eggenschwiler and Anderson, 2007). Activated ciliary SMO then works through KIF7 to promote the dissociation of the inhibitory SUFU-GLIFL complex (Humke et al., 2010; Tukachinsky et al., 2010; Li et al., 2012). Free GLIFL is then processed into an activator and moves to the nucleus to activate downstream targets. In the absence of the SHH ligand, SMO is not translocated into the cilium and thus cannot antagonize SUFU. SUFU remains in complex with GLIFL, GLIFL is proteolytically processed into GLIT and the SUFU-GLIT complex moves to the nucleus where it recruits the Sap18-Sin3 co-repressor complex to repress GLI target genes (Ding et al., 1999; Kogerman et al., 1999; Cheng and Bishop, 2002; Paces-Fessy et al., 2004). Furthermore, our recent work with the murine ciliary mutant, Kif3fl/fl ;Wnt1-Cre shows increased SUFU production and nuclear localization, as well as enhanced association of SUFU with GLI3 (Chang et al., in press). We hypothesize a similar mechanism is at play in ta2 mutants. Specifically, excessive GLIFL produced in ta2 mutants cannot dissociate from SUFU because the complex cannot undergo ciliary trafficking. We further hypothesize that lack of dissociation prevents GLIFL from occupying GBRs and directly activating target gene transcription. Our future studies will address the association of GLIFL and SUFU in ta2 mutants and determine if their maintained association contributes to ciliopathic phenotypes.

# GLI-Binding to Target Genes Occurs in a Prominence Specific Manner

Frequently the craniofacial complex is thought of as a singular organ system; however, the prominences that make up the face develop independently prior to their fusion. There is evidence to support that these prominences have distinct molecular profiles and develop as separate developmental fields (Brugmann et al., 2007). Our ChIP-qPCR results detected differential GLI binding to target genes in facial prominences of control embryos (**Figures 2B–D**). Further, the expression of co-regulators also followed a prominence specific pattern in both control and ta2 embryos (**Figures 6A'–C'**). These data supported the hypothesis that during normal development, as well as when cilia are lost, each facial prominence uses a unique mechanism to transduce a SHH-dependent GLI signal. Based on our examination of coregulators, we hypothesize that there is a combinatorial code of GLI isoforms and co-regulators that work together to precisely regulate target gene expression. Thus, when cilia are lost and GLI production is altered, target gene expression is dictated by how the combinatorial code of remaining GLI isoforms and co-regulators function together.

In sum, our data suggest that the increased or decreased production of GLI isoforms alone is not sufficient to explain how target gene expression will be altered. To understand the molecular mechanisms responsible for ciliopathic, GLImediated phenotypes, future studies will have to account for tissue specificity, the presence or absence of co-regulators and the mode of GLI function (activation or de-repression) to begin to address this process.

# AUTHOR CONTRIBUTIONS

YC designed and tested all primers, and performed and interpreted all ChIP and qPCR experiments, PC performed in silico GBR analysis, ES harvested and photographed embryos, SB conceived the project, analyzed data and wrote the manuscript with input from all authors.

# FUNDING

This work was funded by the National Institutes of Health, National Institute of Dental and Craniofacial Research (NIDCR) [R00-DE01985 and [R01-DE023804] to SB] and by C.T.O. and Research in Progress (RIP) funds from the Cincinnati Children's Research Foundation to SB.

# ACKNOWLEDGMENTS

We thank the UC Davis Avian Facility and Jackie Pisenti for husbandry of the talpid2 line and providing talpid2 eggs for this study.

# SUPPLEMENTARY MATERIAL

The Supplementary Material for this article can be found online at: http://journal.frontiersin.org/article/10.3389/fphys. 2016.00468

Supplementary Figure 1 | GLI3 binding to GLI targets is confirmed by ChIP. ChIP-qPCR analyses of GLI3 precipitated with GLI binding regions of *ALX4* (A), *BMPR1A* (B), *OSR2* (C), *WIF1* (D), and *PTCH1* (E) in facial prominences of control embryo. Error bars are based on the standard error of the means (S.E.M.). *n* = 3.

# REFERENCES

Supplementary Figure 2 | ChIP-qPCR analyses of GLI3 precipitated with GLI binding regions of PTCH1 (A), TFAP2A (B), CTNNB1 (C), WNT2B (D), DKK1 (E), FGFR (F), BMP2 (G), BMP4 (H), BMP7 (I), and EPHA4 (J) in control and ta2 facial prominences. Error bars are based on the standard error of the means (S.E.M.). *n* = 3.

Supplementary Figure 3 | The schematic of the clusters of transcription factor motifs in GBRs of GLI targets. The *in silico* analyses of GLI (blue circle), E-box (magenta triangle), SP site (red rectangle), and SOX site (yellow circle) in the GBRs of GLI targets. We defined 5′ -untranslated region (UTR), gene and 3′ -UTR as intragenic region, <1 kb upstream of transcription start site (TSS) is promoter region, 1–20 kb away from TSS as proximal regulatory region, and <100 kb away from TSS as distal regulatory region. The numbers labeled below the symbols indicate the positions of the motifs according to the distance away from TSS sites. The position at the upstream of TSS site is assigned a negative symbol. The primer sets used for ChIP-quantitative PCRs are labeled above the specific GLI binding motif (red line). Clusters of four motifs are highlighted in a black box, clusters of three motifs are highlighted in a red box.

### Supplementary Table 1 | Identified Gli target genes.

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2016 Chang, Chaturvedi, Schock and Brugmann. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Mesenchymal Remodeling during Palatal Shelf Elevation Revealed by Extracellular Matrix and F-Actin Expression Patterns

Matthias Chiquet 1 \*, Susan Blumer 1 , Manuela Angelini 1 , Thimios A. Mitsiadis 2 and Christos Katsaros 1

1 Department of Orthodontics and Dentofacial Orthopedics, Medical Faculty, School of Dental Medicine, University of Bern, Bern, Switzerland, 2 Orofacial Development and Regeneration, Center for Dental Medicine, Institute for Oral Biology, University of Zurich, Zurich, Switzerland

During formation of the secondary palate in mammalian embryos, two vertically oriented palatal shelves rapidly elevate into a horizontal position above the tongue, meet at the midline, and fuse to form a single entity. Previous observations suggested that elevation occurs by a simple 90◦ rotation of the palatal shelves. More recent findings showed that the presumptive midline epithelial cells are not located at the tips of palatal shelves before elevation, but mostly toward their medial/lingual part. This implied extensive tissue remodeling during shelf elevation. Nevertheless, it is still not known how the shelf mesenchyme reorganizes during this process, and what mechanism drives it. To address this question, we mapped the distinct and restricted expression domains of certain extracellular matrix components within the developing palatal shelves. This procedure allowed to monitor movements of entire mesenchymal regions relative to each other during shelf elevation. Consistent with previous notions, our results confirm a flipping movement of the palatal shelves anteriorly, whereas extensive mesenchymal reorganization is observed more posteriorly. There, the entire lingual portion of the vertical shelves moves close to the midline after elevation, whereas the mesenchyme at the original tip of the shelves ends up ventrolaterally. Moreover, we observed that the mesenchymal cells of elevating palatal shelves substantially align their actin cytoskeleton, their extracellular matrix, and their nuclei in a ventral to medial direction. This indicates that, like in other morphogenetic processes, actin-dependent cell contractility is a major driving force for mesenchymal tissue remodeling during palatogenesis.

Keywords: palate morphogenesis, palatal shelf elevation, extracellular matrix, actin, tissue remodeling, mouse embryo

# INTRODUCTION

The secondary palate, which separates the oral cavity from the nasal cavity, is a specific feature of mammals (Ferguson, 1988). Cleft lip/palate is the most frequent craniofacial malformation in humans (Jugessur et al., 2009a; Mossey et al., 2009; Dixon et al., 2011). Formation of the secondary palate during embryogenesis is a complex process involving extensive tissue growth and remodeling (Gritli-Linde, 2008; Jugessur et al., 2009b; Bush and Jiang, 2012). Palate development

### Edited by:

Giovanna Orsini, Marche Polytechnic University, Italy

### Reviewed by:

Gianpaolo Papaccio, Seconda Università degli Studi di Napoli, Italy Catherine Chaussain, Paris Descartes University, France

\*Correspondence: Matthias Chiquet<EMAIL_ADDRESS>
### Specialty section:

This article was submitted to Craniofacial Biology, a section of the journal Frontiers in Physiology

Received: 11 July 2016 Accepted: 23 August 2016 Published: 07 September 2016

### Citation:

Chiquet M, Blumer S, Angelini M, Mitsiadis TA and Katsaros C (2016) Mesenchymal Remodeling during Palatal Shelf Elevation Revealed by Extracellular Matrix and F-Actin Expression Patterns. Front. Physiol. 7:392. doi: 10.3389/fphys.2016.00392

**142**

starts from the palatal shelves, extensions of the maxillary processes that are located left and right from the tongue. The palatal shelves initially have a vertical orientation, but around the 6th week of gestation in humans, or at embryonic day 14.5 (E14.5) in mice, they elevate rapidly into a horizontal position above the tongue. The shelves meet medially, and fuse by disappearance of the midline epithelium and merging of the mesenchyme (Gritli-Linde, 2007; Bush and Jiang, 2012). While much information exists concerning the mechanism of palatal shelf fusion (Kaartinen et al., 1995; Proetzel et al., 1995) (Jin et al., 2014), the process of shelf elevation still remains elusive, and knowledge about its cellular and molecular basis is scarce (Bush and Jiang, 2012).

Palatal shelves are known to be divided into distinct regions along their anterior-posterior axis. Their frontal and medial parts will give rise to the hard palate, their posterior part to the soft palate (Yu and Ornitz, 2011; Smith et al., 2012). It was debated whether palatal shelves elevate and fuse first from the front or from the back (reviewed in Brinkley and Vickerman, 1979). In vitro, their medial parts elevate first (Brinkley and Vickerman, 1979), whereas in the embryo, tongue movements strongly affect shelf elevation (Iseki et al., 2007; Kouskoura et al., 2016). This causes a remarkable variability in the time course of elevation in vivo between siblings, and even between the two shelves of one embryo (Luke, 1984; Iseki et al., 2007; Yu and Ornitz, 2011). A recent histomorphometric analysis concludes that in the embryo, elevation usually starts at the medial to posterior part of the palatal shelves, which then zipper in both directions (Yu and Ornitz, 2011).

The above findings imply that all parts of the palatal shelves have an intrinsic capability to elevate. However, it has been debated how this reshaping occurs, and several reports indicated regional differences in tissue reorganization during shelf elevation. Histological observations suggested that anteriorly, the palatal shelves perform a simple flipping movement (Coleman, 1965). However, other reports indicated that the medial and posterior parts of the shelves remodel extensively during elevation, such that their lingual side protrudes whereas the ventral tip regresses (Brinkley and Vickerman, 1979; Chou et al., 2004). A recent study used the expression of the Mmp13 gene to determine the origin of midline epithelial cells during palatal shelf elevation. Its results supported a flipping movement of the shelves in the front and extensive remodeling posteriorly (Jin et al., 2010), which was confirmed by careful histomorphometric analysis (Yu and Ornitz, 2011).

Although there is general agreement that extensive remodeling of the palatal shelves occurs during elevation, it is not known how different regions of the palatal mesenchyme move relative to each other. Moreover, the forces that drive tissue remodeling in the elevating shelves are still unknown. Distinct patterns of cell proliferation (Sasaki et al., 2004) and density (Brinkley and Bookstein, 1986) are unlikely to contribute substantially to the remodeling process, since it occurs in less than 1 day in mouse embryos (Yu and Ornitz, 2011). The finding of hyaluronidase-sensitive material in the "hinge" region of palatal shelves led to the hypothesis that rapid accumulation of hyaluronic acid or other glycosaminoglycans might provide the driving force for shelf elevation, by generating osmotic swelling pressure (Brinkley and Morris-Wiman, 1987). Alternatively, this process might be powered by coordinated cytoskeletal contractions of both epithelium and mesenchyme, similar to what is observed in many other morphogenetic tissue rearrangements (Wozniak and Chen, 2009). Although an early report suggested a role for actin-based contractility in palatal shelf elevation (Lessard et al., 1974), to our knowledge this idea has never been followed up.

Therefore, in the present study we address two important questions concerning the process of palatal shelf elevation. Firstly, is it possible to detect rearrangements of different parts of the palatal mesenchyme relative to each other during the elevation process? Secondly, can we find any evidence for actinbased contractility in the remodeling of palatal mesenchyme during elevation? To address these points, we took advantage of the fact that certain extracellular matrix (ECM) components have a restricted expression in defined regions of the palatal mesenchyme, which can be followed during shelf elevation. Furthermore, we stained sections through elevating palatal shelves for F-actin, and detected elongated cell nuclei that aligned with the actin network. Our results confirm and refine previous findings of extensive remodeling of the palatal mesenchyme at medial and posterior levels, and provide the first evidence for tensile stress acting within this tissue during the process of shelf elevation.

# MATERIALS AND METHODS

# Animals, Embryonic Tissues, and Cryosectioning

C57BL/6 wild-type mouse embryos were obtained from J.-F. Spetz at the Friedrich-Miescher Institute for Biomedical Research in Basel, Switzerland, or from in house breeding at the central animal facilities, Department of Clinical Research, University of Bern. After mating, appearance of a vaginal plug was considered embryonic day 0.5 (E0.5). Pregnant females were sacrificed at the desired stage (E13.5–E14.5), embryos were removed from the uterus and decapitated. Animal experiments were approved by the Cantonal Veterinary Offices of Basel, Zurich and Bern, Switzerland. The embryo heads were washed in ice-cold PBS, fixed in 4% paraformaldehyde in phosphate buffered saline (PBS; 150 mM NaCl, 20 mM Na-phosphate, pH 7.4) overnight, soaked for 24 h in 30% sucrose in PBS, embedded in Tissue Tek (O.C.T. compound; Sakura Finetek Europe B.V., Zoeterwoude, Netherlands), and frozen on an aluminum block cooled to −80◦C. Frozen heads were stored at −80◦C before sectioning. Serial frontal sections (12 µm thick) were prepared on a Cryocut E cryomicrotome (Reichert-Jung, Leica Microsystems, Heerbrugg, Switzerland). Sections were dried at 37◦C for 1–5 min, and stored at −80◦C for further use.

# Gene-Specific RNA Probes and In situ Hybridization

Total RNA was isolated from E14.5 C57BL/6 wildtype mouse embryos using an RNAeasy Mini Kit (Qiagen, Hombrechtikon, Switzerland), and reverse transcribed to cDNA using Moloney murine leukemia virus reverse transcriptase (Promega, Dübendorf, Switzerland). Gene-specific primers (Microsynth, Balgrach, Switzerland) were designed using free NCBI software (http://www.ncbi.nlm.nih.gov/tools/primer-blast/index.cgi?

LINK\_LOC=BlastHome). Primers were fitted with BamH1 (forward primers) or HindIII (reverse primers) restriction sites at their 5′ ends, respectively. With these primers (**Table 1**), specific products were amplified from E14.5 mouse cDNA by PCR using Go Taq polymerase (Promega), cut with respective restriction enzymes, and cloned into pBluescript SK+ plasmid (Stratagene/Agilent, Santa Clara, USA). In the case of mouse tenascin-C (Tnc) and tenascin-W (gene name Tnn), full-length cDNA clones were obtained from R. Chiquet-Ehrismann (Friedrich Miescher Institute for Biomedical Research, Basel, Switzerland), and restriction fragments of suitable size (**Table 1**) were cloned into pBluescript SK+. Plasmids were linearized by cutting upstream or downstream of the insert, respectively, and digoxygenin-labeled anti-sense or sense RNA probes were synthesized (Koch et al., 1995) with a labeling kit from Roche Diagnostics using T3 or T7 RNA polymerase (Promega). In situ hybridization was done with the labeled probes as published in detail before (Flück et al., 2000). After incubation with alkaline phosphatase conjugated anti-digoxygenin antibody (Roche Diagnostics), a color substrate solution was used to which 10% polyvinyl alcohol (MW 31′ 000-50′ 000; Sigma-Aldrich, Buchs, Switzerland) was added (Shen, 2001). After development, the sections were counterstained with Nuclear Fast Red (Sigma).

# Fluorescence Labeling of Tissue Sections

Monoclonal antibody mTn12 against mouse tenascin-C (Aufderheide and Ekblom, 1988) was obtained from

### TABLE 1 | Gene-specific RNA probes for in situ hybridization.

### MOUSE PERIOSTIN (Postn: NM\_015784.3)

Forward primer: 5′CCGGATCCGCAGCCGCCATCACCTCTGAC 3′ (nucleotide: 196-216)

Reverse primer: 5′ CCAAGCTTGCTTGTCTTGCCGCAGCTTGT 3′ (nucleotide: 964-944)

PCR product/probe length (without restriction sites): 769 bp

### MOUSE TENASCIN-C (Tnc: D\_90343.1)

Restriction fragment: Sac1 (nucleotide 5862)–EcoR1 (nucleotide 6813) Probe length: 950 bp

### MOUSE TENASCIN-W (Tnn: AJ\_580920.1)

Restriction fragment: HindIII (nucleotide 1190)–BamH1 (nucleotide 1440) Probe length: 250 bp

### MOUSE SMOC-2 (Smoc2: NM\_022315.2)

Forward: 5 ′ CCGGATCCCAGGTGCCCCGGTTCAA 3′ (nucleotide: 661-679) Reverse: 5 ′ CCAAGCTTCCAGGGTGTGGCTGGGGT 3′ (nucleotide: 1253-1235) PCR product/probe length (without restriction sites): 593 bp

### MOUSE TRANSFORMING GROWTH FACTOR-β3 (Tgfb3: NM 009368.3)

Forward primer: 5′ CCGGATCCCAACCCCAGCTCCAAGCG 3′ (nucleotide: 1578-1597)

Reverse primer: 5′ CCAAGCTTCCAGGTTGCGGAAGCAGT 3′ (nucleotide: 2046-2026)

PCR product/probe length (without restriction sites): 469 bp

R. Chiquet-Ehrismann (Friedrich Miescher Institute for Biomedical Research, Basel, Switzerland). Cryosections from paraformaldehyde-fixed mouse embryo heads were blocked with 3% bovine serum albumin in phosphate-buffered saline (BSA/PBS) and incubated with anti-tenascin antibody (1:100 in BSA/PBS) for 1 h at room temperature. After washing with PBS containing 0.2% Tween-20, sections were incubated with Alexa488-labeled secondary antibody (Invitrogen; 1:1000 in BSA/PBS). For labeling the F-actin network on the same section, 1 µg/ml rhodamine-phalloidin (Sigma) was added to the secondary antibody solution. After washing again with PBS/Tween, sections were embedded in phosphate-buffered glycerol (pH 7.4) containing 1 µg/ml DAPI stain (Sigma) to label nuclei.

# Microscopy

Slides processed for in situ hybridization were viewed under bright field optics with 4x or 10x objectives on an Olympus BX-51 microscope. Slides triple-labeled with fluorescent dyes were observed on the same microscope by epifluorescence optics, using 10x and 20x fluorescence objectives and the following Olympus filter/mirror series: U-MWIBA3 for Alexa488; U-MWIGA3 for rhodamine; U-MNUA2 for DAPI. Digital images were recorded using a ProgRes CT3 CMOS camera and ProgRes Capture Pro software (Jenoptik, Jena, Germany). Slides from one experiment were photographed at identical camera settings, and resulting images were processed identically.

# Image Analysis

DAPI-labeled nuclei on tissue sections were color-coded depending on their aspect ratio (ratio of x to y axis) in the following way. Images were imported into ImageJ, binarized, and the Watershed tool was used to separate touching/overlapping nuclei. Particles (i.e., DAPI-stained nuclei) were fitted with ellipses, and their aspect ratio was analyzed. A ROI (region of interest) color coder plugin with the "Phase" LUT (look up table) was then used to color-code all nuclei with an aspect ratio of >2.5 and above in red, 2.0–2.5 in pink, and <2.0 in blue.

# RESULTS

# Periostin as a Mesenchymal Marker for Palatal Shelves during Elevation

To generate a fate map of palatal mesenchyme and epithelium during the process of shelf elevation, we screened the expression patterns of more than 20 ECM components by in situ hybridization on frontal sections of wild-type C56/BalbC mouse embryo heads during the relevant stages, i.e., E13.5–E14.5. Of these genes, the expression of eight was highly enriched in either all or just a specific region of the palatal shelves compared to the surrounding maxillary tissue: Periostin, tenascin-C, tenascin-W, collagen types II, IX, XI, and XIV, and SMOC-2.

Periostin (gene name Postn) is an ECM component belonging to the "matricellular" proteins (Murphy-Ullrich and Sage, 2014) and known to be involved in morphogenesis and wound healing (Walker et al., 2016). Periostin protein was demonstrated in mouse palatal shelves by immunofluorescence (Oka et al., Chiquet et al. Remodeling of Elevating Palatal Shelves

2012). Here, we used in situ hybridization to reveal the expression pattern of Postn mRNA in palatal shelves of wild-type mouse embryos immediately before (E13.5) and after (E14.5) elevation. Serial frontal cryosections through palatal shelves were hybridized with Postn-specific RNA probe at three levels: anterior (future anterior hard palate), medial (future posterior hard palate), and posterior (future soft palate). Anteriorly, Postn mRNA was expressed in a broad layer of mesenchyme on both the buccal (ventrolateral) and lingual (medial) aspect of E13.5 palatal shelves (**Figure 1A**). After shelf elevation (E14.5), the layer of Postn expression was located ventrolaterally in the shelves, and filled their medial tips (**Figure 1B**). At the medial level, a strong Postn signal covered the entire mesenchymal area of vertical palatal shelves at E13.5 (**Figure 1C**). After elevation at this level, Postn expression was observed in a broad area around the midline, from where it extended into a ventral stripe facing the oral cavity (**Figure 1D**). In contrast and remarkably, the dorsolateral regions of the elevated shelves were free of Postn signal (**Figure 1D**). A similar distribution was seen at the posterior level both before and after elevation, except that the signal was weaker in the central mesenchyme region of the shelves (**Figures 1E,F**).

# Tenascin-C Expression in the Medial Palatal Mesenchyme Both before an after Shelf Elevation

Tenascin-C (gene name Tnc) is another "matricellular" ECM protein with highly regulated patterns of expression in vertebrate morphogenesis and pathologies (Chiquet-Ehrismann and Chiquet, 2003). Like periostin, tenascin-C protein has been detected in the developing secondary palate before by immunofluorescence (Ferguson, 1988). At the anterior level at E13.5, the Tnc mRNA expression pattern resembled that of periostin: In vertical shelves, the signal was found in the sub-epithelial mesenchyme both on the lingual (medial) and the buccal (ventrolateral) aspect (**Figure 2A**). After elevation, Tnc mRNA was concentrated at the tips of the shelves and extended laterally into the subepithelial mesenchyme both dorsally and ventrally (**Figure 2B**). A different distribution was observed at the medial level, i.e., in the future posterior hard palate. There, Tnc expression covered the entire lingual half of the vertical shelves at E13.5, whereas the buccal half was devoid of signal except for a thin stripe underneath the epithelium (**Figure 2C**). In elevated shelves (E14.5) at this level, Tnc mRNA was observed in a broad mesenchymal region that extended from dorsal to ventral around the midline, and in a thin stripe underneath the ventral epithelium (**Figure 2D**). A similar distribution was seen for Tnc at the posterior level (future soft palate), although the signal was weaker (**Figures 2E,F**).

# Dorsomedial Tenascin-W Expression in Palatal Shelves Both before and after Shelf Elevation

Tenascin-W (gene name Tnn) is the latest member of this family of ECM proteins (Scherberich et al., 2004). During embryogenesis, its expression partially overlaps with that of tenascin-C but is even more restricted. Notably, tenascin-W was shown to be involved in osteogenesis (Martina et al., 2010) and is a marker for osteogenic areas in embryos (Scherberich et al., 2004). In anterior palatal shelves at E13.5, we found Tnn to be expressed exclusively in the dorsomedial quadrant of the mesenchyme, whereas the tips of the vertical shelves were devoid of signal (**Figure 3A**). At E14.5, the Tnn signal was restricted to the dorsal mesenchyme around the midline of the elevated shelves (**Figure 3B**). A similar distribution was found at the medial level both at E13.5 and E14.5, except that in the elevated shelves the area of Tnn expression was spread out laterally and filled the entire dorsal half of the shelf mesenchyme (**Figures 3C,D**). More posteriorly, the Tnn signal was present in addition around the developing os palatinum (**Figures 3E,F**).

# SMOC-2 Expression at Palatal Shelf Tips before, but Not at the Midline after Elevation

SMOC-2 (SPARC related modular calcium binding 2; gene name Smoc2) belongs to the SPARC/Osteonectin family of matricellular proteins (Vannahme et al., 2003) and is involved in keratinocyte migration (Maier et al., 2008) and angiogenesis (Rocnik et al., 2006). In mouse embryos, Smoc2 is prominently expressed in perichondrial mesenchyme (**Figure 4A**). Surprisingly, Smoc2 mRNA is also detected in defined areas of the oral epithelium. At a medial to posterior level before elevation (E13.5), the Smoc2 signal outlines the buccal surface of the palatal shelves and extends into the epithelium at their tips, but finishes abruptly at the lower end of their lingual aspect (**Figure 4A**). At E14.5, Smoc2 mRNA is present in the entire ventral epithelium of the elevated shelves, but stops shortly before the midline (**Figure 4B**). A similar expression pattern in palatal epithelium before and after elevation was found for type XIV collagen, also named undulin, a minor fibril-associated collagen (Ricard-Blum, 2011) (not shown). For comparison, sections from a similar level were hybridized with a Tgfβ3 probe. This growth factor is secreted by midline epithelial cells and required for palatal shelf fusion (Kaartinen et al., 1995). At E13.5, Tgfβ3 mRNA extended from the shelf tip about halfway up the lingual epithelium (**Figure 4C**), and at E14.5 was found at the midline (**Figure 4D**). Thus, Smoc2 and Tgfb3 exhibit a reciprocal expression pattern in the palatal epithelium, except for a short overlap at the shelf tip before, or close to the ventral midline after elevation, respectively.

# Mesenchymal and Epithelial Rearrangements during Palatal Shelf Elevation at the Medial Level

Since the mRNAs of the chosen ECM genes were enriched in distinct areas of the palatal shelves before and after elevation, this allowed the mapping of their combined expression patterns (**Figure 5**; compare with **Figures 1**–**4**). At the medial level, the combined results implied that elevation is not due to a simple rotation of the palatal shelves from the vertical into the horizontal plane. If this were the case, the Tnc-expressing lingual half of the vertical shelves should end up dorsally in the elevated secondary palate. Instead, the Tnc-expressing

FIGURE 1 | Periostin expression as a marker for palatal shelves during elevation. In situ hybridization for Postn mRNA on frontal sections of palatal shelves from wild-type mouse embryos at the anterior (A,B), medial (C,D), and posterior (E,F) level. Serial sections were obtained from an E13.5 (A,C,E) and an E14.5 (B,D,F) embryo, respectively. For description, see text. Abbreviations: palatal shelf (p), tongue (t). Bar, 400 µm.

area had moved to a broad zone at the midline, whereas the lateral parts of the elevated shelves were devoid of Tnc signal. The Postn signal, which filled the entire shelves at E13.5, was drawn out after elevation into a broad tringle that filled the entire mesenchyme around the midline, and tapered on the ventrolateral aspect of the shelves (cf. **Figure 1D**). Mesenchyme around the nasopharyngeal fold that forms at E.14.5 lacked expression of Postn (**Figure 5**), suggesting that maxillary tissue originally outside the palatal shelf proper is drawn into the elevated shelves at E14.5. Thus, at this level the palatal shelf bulges out medially whereas the original tip retracts during elevation. This involves massive reorganization of the palatal mesenchyme: The lingual mesenchyme of the E13.5 shelf, rather than that at the distal tip, forms the tissue around the midline at E14.5. The mesenchyme at the original tip ends up more laterally after elevation, whereas the originally buccal mesenchyme distorts and

FIGURE 2 | Tenascin-C expression primarily in the lingual/medial half of palatal shelves both before and after shelf elevation. In situ hybridization for Tnc mRNA on frontal sections of palatal shelves from wild-type mouse embryos at the anterior (A,B), medial (C,D), and posterior (E,F) level. Serial sections were obtained from an E13.5 (A,C,E) and an E14.5 (B,D,F) embryo, respectively. For description, see text. Abbreviations: palatal shelf (p), tongue (t). Bar, 400 µm.

spreads out along the ventrolateral aspect of the elevated shelves. Remodeling of the mesenchyme is paralleled by reorganization of the palatal epithelium. Smoc2 mRNA was detected in shelf tip epithelium before, but not in midline epithelium after elevation. Conversely, Tgfβ3 was expressed primarily in lingual epithelium of shelves before, and in midline epithelium after elevation (**Figures 4**, **5**).

# Anterior to Posterior Differences in ECM Rearrangement during Shelf Elevation

In wild-type Balb/C litters collected at E14.5, it is quite frequent to find embryos in which either one or both of the palatal shelves have not elevated yet (Luke, 1984; Yu and Ornitz, 2011). These mice can be compared to their siblings with already elevated shelves, and one representative example is shown in

**Figure 6**. The images show frontal sections at three levels through the palatal shelves of two different E14.5 embryos from the same litter, stained by fluorescence for actin (red) and tenascin-C (green). **Figure 6A** shows a still vertical palatal shelf of one E14.5 embryo at the anterior level, and it is noteworthy that a prominent epithelial invagination has formed on its buccal side. Staining for tenascin-C protein largely fills the shelf mesenchyme, except for its central area. At the same level in the sibling (**Figure 6B**), its elevated shelf occupies the space between nasal process and tongue, and the epithelial invagination has disappeared. Tenascin-C is present in the medial and ventrolateral parts of the shelf. Since tenascin-C positive regions largely correspond to each other before and after elevation, these observations agree with a flipping movement of the anterior palatal shelves. On the other hand, comparison of shelves from the same two embryos also confirms that

extensive mesenchymal reorganization takes place at the medial and posterior level during elevation. At the medial level, a small protrusion has formed on the dorsolingual aspect of the still vertical shelf (**Figure 6C**). Tenascin-C is found primarily in the lingual half of the shelf mesenchyme, with only a narrow ribbon underneath the buccal epithelium. In case of the elevated shelf at the same level (**Figure 6D**), the entire tenascin-C containing mesenchymal shelf ECM between the original tip and the dorsolingual protrusion appears compressed between nasal process and tongue. Remarkably, the palatal artery appears circular in cross-section by actin staining before elevation, but is flattened in the vertical direction after elevation (c.f. **Figures 6C,D**). At the posterior level, the dorsolingual protrusion is very prominent on the not yet elevated shelf, forming a second tip (**Figure 6E**). Tenascin-C protein is enriched in the mesenchyme between the two shelf tips, rather than at the tips themselves. At this level, it is most obvious that the entire tenascin-C containing mesenchyme has moved to the midline after elevation (**Figure 6F**). Rather than a flipping movement as seen at the front, this again implies a slug-like reshaping of the medial/posterior palatal shelf, with a deformation and medial movement of the lingual mesenchyme (including the palatine artery) and a retraction of the original shelf tip.

# Orientation of Actin Fibers and Nuclei in Palatal Mesenchyme during Elevation

Fluorescently labeled phalloidin is a specific marker for actin stress fibers of cells in culture (Small et al., 1999), and can also be used to label the cytoskeletal F-actin network of embryonic tissues (**Figures 7A,B**). As a major component of the contractile apparatus, actin stress fibers are known to align with the major direction of strain within a cell (Burridge and Wittchen, 2013). In **Figure 7A**, a higher magnification of a E14.5 palatal shelf at the medial level just before elevation (c.f. **Figure 6C**) is shown after phalloidin labeling. Note that the F-actin network of mesenchymal cells is aligned within this shelf, pointing toward the original ventral tip and the newly forming lingual protrusion, and forming an arc between them. In already elevated shelves of the same stage, the cellular actin network appears less organized, and no preferred direction is apparent in the shelf proper (**Figure 7B**). We also determined, on the same sections, the deformation of DAPI-labeled cell nuclei in the palatal mesenchyme. Using ImageJ, the ellipticity

level before (E13.5; above) and after (E14.5; below) elevation. The approximate areas of mesenchymal expression of periostin, tenascin-C, tenascin-W, and epithelial expression of Smoc2 and Tgfβ3 are indicated. Note that tenascin-C, tenascin-W, and Tgfβ3 expressing areas all end up at the midline after elevation, although their center of expression does not localize to the shelf tip before elevation. Conversely, Smoc2 mRNA is epithelially expressed at the shelf tip before, but excluded from midline epithelium after elevation.

of stained nuclei was measured, and nuclei with an aspect ratio of >2.5 were labeled in red. As can be seen in **Figure 7C**, elongated (red) nuclei are enriched in, and align with, the arclike F-actin network that is visible between the ventral and the lingual tips of this shelf before elevation. In contrast, elongated nuclei appear sparse and have no preferred direction in the elevated shelf (**Figure 7D**); they are more frequent in lateral maxillary mesenchyme. Note further that antibody to tenascin-C labels an ECM meshwork that lies parallel to the actin arc and the elongated nuclei in elevating shelves (**Figure 7E**). After elevation, the tenascin-C positive ECM has moved toward the midline as seen before, and the ECM fibrils appear randomly arranged (**Figure 7F**). These observations suggest that tensile stress builds up in the palatal shelves before elevation, which is released after the shelves have reached the horizontal position.

# DISCUSSION

# Remodeling of Palatal Shelf Mesenchyme during Elevation Revealed by ECM Expression Patterns

In the present study, we took advantage of the differential expression patterns of certain ECM genes to monitor tissue rearrangements during palatal shelf elevation. After screening over 20 ECM genes by in situ hybridization before and after elevation, we identified a few that exhibited expression patterns restricted to specific areas of the shelf mesenchyme or epithelium. Genes for ECM components are strongly expressed, and the turnover of their mRNAs is relatively slow (Eckes et al., 1996). We therefore argue that the observed changes in ECM expression patterns reflect morphogenetic rearrangements of palatal shelf tissue, rather than alterations in the rate of gene transcription. In the case of tenascin-C, in situ hybridization results were further confirmed by antibody stainings. This makes it even more unlikely that the observed dynamic changes in tenascin-C patterns during shelf elevation are due to rapid alterations in its synthesis.

Our results confirm the findings of Jin et al. (2010) concerning palatal shelf remodeling during elevation (see below). In addition, they extend earlier studies by providing more detailed information on the reshaping and medial translocation of the lingual shelf mesenchyme during this morphogenetic process. A further observation is noteworthy: During elevation, epithelial folds form dorsolaterally from the palatal shelves; these folds mark the lateral edges of the nasal cavity. Interestingly, the mesenchyme surrounding these folds express little or no periostin, which before elevation is a marker for the entire shelf proper. This suggests that at the epithelial folds, some originally nasal mesenchyme might be pulled into the elevating shelves. We have shown before that the epithelial folds are characterized by high expression of Mmp2 mRNA and gelatinase activity, which presumably is needed there to avoid closure of the nasal cavity (Gkantidis et al., 2012). From our results, these specialized structures might originate from tissue outside the original shelf proper.

The mechanism of shelf elevation during secondary palate formation in mammals has been disputed (Brinkley and Vickerman, 1979; Jin et al., 2010; Bush and Jiang, 2012). Early reports favored a simple flipping movement of the vertical palatal shelves into a horizontal position (Coleman, 1965), but it was soon postulated that posteriorly the palatal shelves reorganize differently. From histological observations, it appeared that their lingual/medial aspect bulged out whereas their original ventral tip retracted (Brinkley and Vickerman, 1979). This process requires extensive remodeling of the entire shelf, but the underlying mechanism remained obscure. Recently, Jin et al. (2010) found that matrix metalloproteinase Mmp13, a specific marker for midline epithelial cells, was expressed in still vertical E14.5 shelves posteriorly not at their tip but on their lingual/medial aspect, whereas anteriorly the Mmp13 signal extended into the shelf tip. The authors concluded that only anteriorly, the original tip epithelium will form the midline of elevated palatal shelves, whereas in middle and posterior regions, the midline epithelial cells are derived from the lingual epithelium of vertical shelves (Jin et al., 2010). These findings were in agreement with the earlier hypothesis that upon elevation, shelves undergo a flipping motion anteriorly, and a reshaping in middle and posterior regions.

(Jin et al., 2010) also showed that the mesenchymal transcription factor Goosecoid was expressed around the midline in E14.5 embryos, but in the lingual rather than the tip mesenchyme of non-elevated shelves of the same stage. This provided the first indication for large scale reorganization of

palatal shelves at the anterior (A,B), medial (C,D), and posterior (E,F) level were double-stained with rhodamine-phalloidin for actin (red) and antibody to tenascin-C (green), and viewed in a fluorescence microscope. Sections were obtained from two embryos of the same E14.5 litter; one in which the shelves had just started to elevate (A,C,E), and one in which they had already risen above the tongue (B,D,F). Abbreviations: palatal shelf (p), tongue (t), nasal septum (n). The palatine artery is marked with an arrow in (C,D), and shown enlarged in the inserts. Bar, 400 µm.

pre-existing mesenchyme during shelf elevation, rather than for differential cell growth as it has been suggested (Brinkley and Bookstein, 1986; Sasaki et al., 2004). In any case, the rapid and often asynchronous time course of shelf elevation (Yu and Ornitz, 2011), as well as our present results, strongly argue against differential cell growth as a mechanism for palatal shelf remodeling.

# Actin-Based Cellular Contractility Combined with Differential ECM Stiffness as a Possible Mechanism for Shelf Remodeling during Elevation

Cells attach to the ECM via integrins, which in turn are linked to the intracellular actin cytoskeleton. Embryonic cells and their

shelves were triple-stained with phalloidin for actin (A,B), DAPI for nuclei (C,D), and antibody to tenascin-C (E,F), respectively, and viewed in a fluorescence microscope. On the left (A,C,E), a shelf just before elevation is shown, and on the right (B,D,F) an already elevated shelf from an embryo of the same litter. The images (C,D) were processed by ImageJ, such that all elongated nuclei with an aspect ratio above 2.5 are false-colored in red, those with 2.0–2.5 in pink. Roundish nuclei (aspect ratio <2.0) are blue. Note that before elevation (C), many elongated (red/pink) nuclei are present between the original tip and the medial protrusion of the elevating shelf (arrowheads in A,C,E), and that their direction parallels that of the actin (A) and ECM (E) meshwork. The palatine artery is marked with an arrow in (A,B). Abbreviations: palatal shelf (p), tongue (t), nasal septum (n). Bar, 200 µm.

ECM thereby form a mechanical continuum, in which actingenerated tensile stresses are transmitted from one cell to the next (Wozniak and Chen, 2009). Stresses align the intracellular F-actin, and also reorient and organize extracellular ECM fibers (Burridge and Wittchen, 2013). Moreover, cell nuclei are mechanically connected to the actin cytoskeleton and deformed by tensile stresses that pull on the cell (Brosig et al., 2010). Thus, preferred orientations of the interconnected actin-integrin-ECM meshwork, together with elongated cell nuclei, reflect vectors of tensile stress that act within embryonic mesenchyme. The fungal toxin phalloidin specifically binds to cellular F-actin (Small et al., 1999). Tenascin-C is known to decorate ECM fibrils in embryonic tissues (Chiquet-Ehrismann and Chiquet, 2003). We used these two tools, together with measurements of the ellipticity of cell nuclei, to determine whether tensile stresses might occur in palatal shelves during their elevation. Our results clearly indicate that in the medial and posterior parts of the elevating shelf, the mesenchyme is tensed between the original tip and the newly forming lingual/medial protrusion. It is more than likely that such tensile stress is generated by F-actin dependent contraction of the shelf mesenchymal cells themselves, and that it is transmitted onto the ECM in which they are embedded.

Despite of several hypotheses that have been put forward over the years, the forces driving palatal shelf elevation were essentially unknown. Walker and Fraser (1956) first postulated an "internal shelf force." A popular hypothesis, still cited in recent reviews (Gritli-Linde, 2007), postulated that a cushion of hyaluronic acid in the "hinge" region of the palatal shelves provides a swelling force by generating osmotic pressure, which pushes up the shelf (Brinkley and Morris-Wiman, 1987). This idea was based on the presence of hyaluronidase-sensitive material in the respective region of the palatal shelf (Brinkley and Morris-Wiman, 1987), and on the fact that a glycosaminoglycan synthesis inhibitor caused cleft palate in mice (Brinkley and Vickerman, 1982). However, the main effect of the inhibitor was to cause a substantial shrinkage in shelf size compared to controls (Brinkley and Vickerman, 1982). Therefore, the inhibitor was likely to cause its damage long before shelf elevation occurred. In essence, there is no direct evidence that hyaluronic acid or other glycosaminoglycans are involved in generating the forces for shelf elevation.

Actin-based cellular contractility is well recognized for driving diverse morphogenetic movements during embryogenesis of both vertebrate and non-vertebrate species (Wozniak and Chen, 2009). Compared to alternatives such as differential cell growth or accumulation of ECM, this mechanism has a large advantage for the involved cells. Namely, they can control it rapidly and precisely in time and space, by activating the intracellular RhoA/ROCK signaling pathway that triggers actomyosin contraction (Burridge and Wittchen, 2013). Surprisingly, although actin contractility has been suggested as a mechanism for palatal shelf elevation early on (Lessard et al., 1974), this idea has apparently not been followed up in more recent years, but is now strongly supported by our current findings.

An actin-based mechanism for palatal shelf reorganization by no means implies that ECM itself has no function in the

# REFERENCES

process. ECM sustains and counteracts forces generated by cellular contractility, and its local stiffness determines to which extent embedded cells are able to deform the tissue (Shawky and Davidson, 2015). Thus, if palatal shelf elevation is driven by cellular actin contraction, differences in ECM composition and stiffness within the shelf might have substantial effects on reshaping of the tissue. Moreover, specific ECM components can modulate cell adhesion, and thereby affect cell contractility. For example, tenascin-C inhibits RhoA/ROCK signaling (and thus actomyosin contraction) by interfering with cell adhesion to fibronectin (Midwood and Schwarzbauer, 2002). It can therefore be speculated that cell contraction is diminished in tenascin-C rich regions within the palatal mesenchyme. Future studies involving a careful mapping of F-actin dynamics, as well as of ECM composition and stiffness, within elevating palatal shelves will be required to fully understand this complex morphogenetic process.

# AUTHOR CONTRIBUTIONS

MC, SB, TM, and CK contributed to the conception and design of the work; MC, SB, and MA were responsible for data acquisition, analysis, and interpretation; MC drafted the manuscript; SB, MA, TM, and CK critically revised the manuscript; MC, SB, MA, TM, and CK approved the manuscript; MC, SB, MA, TM, and CK agreed to be accountable for all aspects of the work.

# FUNDING

MC, TM, and CK were supported by grant 31003A\_146825 from the Swiss National Science foundation.

# ACKNOWLEDGMENTS

We are grateful to François Spetz and Dr. Ruth Chiquet-Ehrismann (Friedrich Miescher Institute for Biomedical Research, Basel, Switzerland), Dr. Daniel Graf (University of Alberta, Edmonton, Canada), and Younes El Fersioui (Department of Orthodontics and Dentofacial Orthopedics, University of Bern, Switzerland) for providing us with tissue and reagents for this study. We thank Sabrina Ruggiero for technical help. MC would like to express his profound gratitude to Ruth Chiquet-Ehrismann, close collaborator, friend and partner for more than 30 years, for her unfailing support throughout his career. Ruth sadly and unexpectedly died on September 4, 2015.

Brinkley, L. L., and Vickerman, M. M. (1979). Elevation of lesioned palatal shelves in vitro. J. Embryol. Exp. Morphol. 54, 229–240.

and migration. Exp. Cell Res. 314, 2477–2487. doi: 10.1016/j.yexcr.2008. 05.020

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2016 Chiquet, Blumer, Angelini, Mitsiadis and Katsaros. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# MORN5 Expression during Craniofacial Development and Its Interaction with the BMP and TGFβ Pathways

Petra Cela1, 2 , Marek Hampl 1, 2, Katherine K. Fu3 , Michaela Kunova Bosakova4 , Pavel Krejci 4, 5, Joy M. Richman3 and Marcela Buchtova1, 2 \*

1 Institute of Animal Physiology and Genetics, v.v.i., Academy of Sciences of the Czech Republic, Brno, Czech Republic, 2 Department of Animal Physiology and Immunology, Institute of Experimental Biology, Masaryk University, Brno, Czech Republic, 3 Life Sciences Institute, University of British Columbia, Vancouver, BC, Canada, 4 Department of Biology, Faculty of Medicine, Masaryk University, Brno, Czech Republic, 5 International Clinical Research Center, St. Anne's University Hospital, Brno, Czech Republic

### Edited by:

Thimios Mitsiadis, University of Zurich, Switzerland

### Reviewed by:

Amel Gritli-Linde, University of Gothenburg, Sweden Javier Catón, CEU San Pablo University, Spain David Clouthier, University of Colorado Anschutz Medical Campus, USA

> \*Correspondence: Marcela Buchtova<EMAIL_ADDRESS>
### Specialty section:

This article was submitted to Craniofacial Biology, a section of the journal Frontiers in Physiology

Received: 05 June 2016 Accepted: 17 August 2016 Published: 31 August 2016

### Citation:

Cela P, Hampl M, Fu KK, Kunova Bosakova M, Krejci P, Richman JM and Buchtova M (2016) MORN5 Expression during Craniofacial Development and Its Interaction with the BMP and TGFβ Pathways. Front. Physiol. 7:378. doi: 10.3389/fphys.2016.00378 MORN5 (MORN repeat containing 5) is encoded by a locus positioned on chromosome 17 in the chicken genome. The MORN motif is found in multiple copies in several proteins including junctophilins or phosphatidylinositol phosphate kinase family and the MORN proteins themselves are found across the animal and plant kingdoms. MORN5 protein has a characteristic punctate pattern in the cytoplasm in immunofluorescence imaging. Previously, MORN5 was found among differentially expressed genes in a microarray profiling experiment of the chicken embryo head. Here, we provided in situ hybridization to analyse, in detail, the MORN5 expression in chick craniofacial structures. The expression of MORN5 was first observed at stage HH17-18 (E2.5). MORN5 expression gradually appeared on either side of the primitive oral cavity, within the maxillary region. At stage HH20 (E3), prominent expression was localized in the mandibular prominences lateral to the midline. From stage HH20 up to HH29 (E6), there was strong expression in restricted regions of the maxillary and mandibular prominences. The frontonasal mass (in the midline of the face) expressed MORN5, starting at HH27 (E5). The expression was concentrated in the corners or globular processes, which will ultimately fuse with the cranial edges of the maxillary prominences. MORN5 expression was maintained in the fusion zone up to stage HH29. In sections MORN5 expression was localized preferentially in the mesenchyme. Previously, we examined signals that regulate MORN5 expression in the face based on a previous microarray study. Here, we validated the array results with in situ hybridization and QPCR. MORN5 was downregulated 24 h after Noggin and/or RA treatment. We also determined that BMP pathway genes are downstream of MORN5 following siRNA knockdown. Based on these results, we conclude that MORN5 is both regulated by and required for BMP signaling. The restricted expression of MORN5 in the lip fusion zone shown here supports the human genetic data in which MORN5 variants were associated with increased risk of non-syndromic cleft lip with or without cleft palate.

Keywords: cleft lip, maxillary prominence, mandibular prominence, frontonasal mass, BMP

# INTRODUCTION

The vertebrate face is formed very early in development from the paired maxillary and mandibular prominences and the single frontonasal mass surrounding the oral cavity. These facial prominences arise during early embryogenesis from interactions between neural crest derived mesenchyme and head ectoderm. The frontonasal mass grows out, contacts and fuses together with the maxillary prominences to form the upper jaw. The midline facial skeleton consisting of the nasal septum, prenasal cartilage and premaxilla are all derived from the frontonasal mass (Richman and Lee, 2003). Craniofacial development is complex process coordinated by a network of transcription factors and signaling molecules (Murray and Schutte, 2004; Chai and Maxson, 2006; Jiang et al., 2006; Brunskill et al., 2014; Kurosaka, 2015; Marcucio et al., 2015; Nimmagadda et al., 2015). Disruption of this tightly controlled cascade can result in clefts where the facial prominences fail to meet and fuse (Leslie and Marazita, 2013).

Cleft lip and/or cleft palate are the most common craniofacial birth defects in humans (Setó-Salvia and Stanier, 2014; Watkins et al., 2014). The majority of clefts appear as isolated or non-syndromic clefts, because they occur in isolation from other developmental abnormalities. The causes of clefting are thought to be multifactorial, including an interaction between genes and the environmental factors (Schutte and Murray, 1999; Dixon et al., 2011; Leslie and Marazita, 2013; Setó-Salvia and Stanier, 2014; Watkins et al., 2014). Identification of genes contributing to clefts formation is important not only for our understanding of facial development, but also for improved prevention and treatment of affected individuals. The chicken embryo is a valuable experimental model to study the signals that control lip fusion. The avian primary palate closely resembles the primary palate in mammals (Abramyan et al., 2015). Moreover, the face can be accessed directly in the living embryo through a window in the shell. The disruption of FGF (Szabo-Rogers et al., 2008), BMP (Ashique et al., 2002), SHH (Hu et al., 2015), and WNT signaling (Geetha-Loganathan et al., 2014) causes a cleft lip in chickens that resembles that of humans.

Previously, a microarray study was performed to profile gene expression in individual chicken facial prominences in stage 18 embryos (Buchtová et al., 2010). From the list of genes that were significantly more highly expressed in the maxillary prominence, we selected MORN5 (also known as C9orf113, C9orf18 or FLJ46909) for further studies because it was described as a cleft susceptibility gene (Letra et al., 2010). Microarray analysis revealed 24 times higher expression of MORN5 in the maxillary prominence compared to expression in the frontonasal mass at stage 18, while mandibular prominence showed 10 times higher expression than the frontonasal mass (Buchtová et al., 2010).

Members of the MORN family were named for the presence of multiple MORN motifs (Membrane Occupation and Recognition Nexus). There are five paralogous genes in the MORN family (MORN1-5). Limited functional information is available for a subset of MORN genes. MORN1 has been identified in the parasite Toxoplasma gondii and other Apicomplexan protists where it plays role during cell division (Ferguson et al., 2008; Lorestani et al., 2010). Human MORN2 was found to facilitate phagocytosis-mediated restriction of some bacteria in macrophages (Abnave et al., 2014). Expression of MORN3 was detected in mouse testis, where it regulates spermatogenesis (Zhang et al., 2015). Finally, MORN4 promotes axonal degeneration in mouse sensory axons (Bhattacharya et al., 2012).

In chicken, the MORN5 gene is located on the forward strand of chromosome 17. On the reverse strand, NDUFA8 and LHX6 genes are nearby to the MORN5 gene. The size of the MORN5 gene is 13.5 kb and there are 6 exons (only 5 exons are coding) with four splice variants. The MORN5 gene encodes a protein of 172 amino acids, which contains a histone H3 K4-specific methyltransferase SET7/9 N-terminal domain (SSF82185) and three MORN motifs (**Figure 1**).

As the gene expression pattern or possible function of MORN5 during development had not been investigated in any animal model, we aim to analyzed chicken MORN5 expression in embryos and its integration into signaling pathways.

# MATERIALS AND METHODS

# Embryonic Material

Fertilized chicken eggs (ISA brown) were obtained from the farm Integra (Žabcice, Czech Republic). Eggs were incubated in a ˇ humidified forced air incubator at 37.8◦C. Embryos were staged and morphological characteristic were described according to Hamburger and Hamilton (1951). All procedures were conducted following a protocol approved by the Laboratory Animal Science Committee of the Institute of Animal Physiology and Genetics (Libechov, Czech Republic). ˇ

# Section In situ Hybridization (ISH)

Chicken MORN5 was obtained as chicken EST clone CHEST ID 543 F09 (Biovalley, France), where the probe sequence was cloned into pBluescript II KS+ vector. The entire region containing the probe sequence flanked by T3 and T7 RNA polymerase sites was amplified using M13 primers (forward primer: 5′ - GTA AAA CGA CGG CCA G-3′ , reverse primer: 5′ -CAG GAA ACA GCT ATG AC-3′ ). Then, the amplicon was isolated via gel purification (QIAquick Gell Extraction Kit, Qiagen, Germany) and this linearized DNA fragment was used in RNA polymerase reactions. DIG labeled antisense riboprobe was synthesized with T3 RNA polymerase (antisense) or with T7 polymerase (sense controls).

Embryos were fixed in 4% paraformaldehyde (PFA), processed through ethanol and xylene into paraffin, and sectioned for ISH. Hybridization was performed with RNA probe at 60◦C overnight as described previously (Holland et al., 1996). Anti-digoxigenin sheep antibody conjugated with alkaline phosphatase (1:2000, Roche, USA) was applied overnight at 4◦C. Visualization was achieved by incubation with substrates for alkaline phosphatase (BM Purple AP, Roche, Germany) for several days. Slides were then counterstained with eosin. ISH was carried out on at least three embryos for each stage.

# Embryo Manipulations

Embryos were treated with beads soaked in All-trans retinoic acid (RA), Noggin protein, Tris or Dimethyl Sulfoxide (DMSO) as described (Lee et al., 2001). Since DMSO was the solvent for RA, we used DMSO bead as a control for RA treatment and Tris as a control for Noggin treatment. AG1-X2 beads (Bio-Rad Laboratories, Hercules, Canada) of 100µm in diameter were soaked in RA (cat. No. R2625 Sigma) at a concentration of 1 mg/ml for 1 h as previously described (Lee et al., 2001; Nimmagadda et al., 2015). Noggin proteins were soaked into Affigel blue beads (Bio-Rad Laboratories, Hercules, Canada) of 200µm diameter for a minimum of 1 h at a concentration of 1 mg/ml (cat. No. 1967-NG, R&D Systems, Minneapolis, USA). Control beads were soaked with DMSO or Tris. Two beads were implanted into the maxillary region on the right side of chicken embryo at stage HH15. For ISH and QPCR, samples were collected 24 h post-bead implantation, embedded into paraffin and processed for ISH.

# Immunofluorescence on Slides

Embryos were collected at stage HH24 for MORN5 protein detection. Chicken duodenum was used as a control according to the manufacturer's instruction. Samples were fixed in 4% PFA and processed into paraffin. Following deparaffinization and rehydration, antigen retrieval was carried out using citrate buffer for 1 min at 97◦C. Polyclonal antibody to MORN5 (1:50, cat. No. NBP1-91230, Novus Biologicals, USA) was applied overnight at 4 ◦C. The secondary anti-rabbit antibody (1:200, Alexa Fluor 594, cat. No. A-21207) was applied for 30 min at RT. Sections were washed in PBS and coverslipped with Prolong Gold anti-fade reagent containing DAPI (cat. No. P36935, Invitrogen, USA).

# Quantitative RT-PCR

Gene expression of MORN5 was analyzed on tissues isolated from normal chicken prominences at stage 15, 18, 20, and 26. Moreover, Noggin or RA treated maxillary prominences were dissected 24 h following bead implantation at stage 15. Prominences were pooled from at least 15 embryos to produce one sample and 4 biological replicates were analyzed. Total RNA was extracted using the Mini RNeasy Kit (Qiagen, Germany) according to the manufacturer's instructions. The total RNA concentration and purity of each sample were assessed by spectrophotometry using a NanoDrop1000 (Thermo Scientific, Waltham, USA). First-strand cDNA was synthesized using the SuperScript Vilo cDNA synthesis Kit (cat. No. 11754050, Thermo Fisher, USA). The qPCR was performed in 10µl final reaction volumes containing the one-step master mix (no AmpErase UNG, cat. No. 4324018, Applied Biosystems, Carlsbad, USA) mixed with MORN5 (TaqMan Assays, Assay ID: AJKAKYV, context sequence: TTCCTGAGAAATGCAGAC GATGAGG, FAM-MGB, Applied Biosystems, Austin, USA) on LightCycler R 96 (Roche, Manheim, Germany) with preheating at 95◦C/10 min, followed by 40 cycles of 95◦C/15 s and 60◦C/1 min. Gene expression levels were calculated using 11CT method with normalization against the HPRT1 level (TaqMan Assays, Assay ID: Gg033338900\_m1, context sequence: TTGAATCATATC TGTGTGATCAGTG, FAM-MGB, Applied Biosystems, Austin, USA), which was used as the housekeeping gene. Means of 3 technical replicates were generated for each of 3 biological replicates and these values were used for statistical analysis. All procedures were repeated in at least three independent experiments.

# Transfection with MORN5 Plasmids in Cell Cultures

The expression vector containing C-terminally FLAG-tagged human MORN5 was obtained from OriGene (Rockville, MD). HEK293T cells were obtained from ATCC (Manassas, VA) and propagated in DMEM media (Sigma-Aldrich, St. Louis, MO) with 10% fetal bovine serum, 1% Pen/Strep and 1% lglutamin (Invitrogen, Carlsbad, CA). Cells were transfected using FUGENE6 reagent according to manufacturer's protocol (Promega).

HEK293T cells grown on glass coverslips were fixed with 4% PFA (RT/15 min), permeabilized with 0.1%Triton-X100 in PBS (RT/5 min), and incubated with the following antibodies at 4◦C overnight: MORN5 (1:100, cat. No. NBP1-91230, Novus Biologicals), FLAG (1:200, cat. No. F1804, Sigma-Aldrich). The secondary antibody AlexaFluor 488 (1:500; cat. No. A21206, Life Technologies) or AlexaFluor 594 (1:500; cat. No. A21203, Life Technologies) were used. Coverslips were mounted into DAPIcontaining Mowiol. Images were taken on an LSM700 laser scanning microscope with acquisition done using ZEN Black 2012 software (Zeiss, Jenna, Germany).

# siRNA Targeting gMORN5 in Chicken Embryos

Silencer Select custom designed siRNA (gMORN5, cat. No. 4399666, Ambion, Austin, USA) was mixed with FUGENE 6 (Roche, Mannheim, Germany), and then was injected into the maxillary prominence of chicken embryos. Negative siRNA (Silencer select negative control No.1 siRNA, cat. No. 4390843, Ambion, Austin, USA) was used as a control. The first injection of siRNA was performed at stage HH20 and the second one after 24 h about stage HH24. One day later after the second injection, embryos had reached stage HH28 and maxillary prominences were dissected for RNA isolation. Tissues were dissected from 5 embryos to form one sample and three biological samples were used for treated embryos (MORN5 siRNA) as well as for control (Silencer select negative control No.1 siRNA) embryos.

# PCR Arrays

Total RNA was extracted from siRNA treated maxillary prominences using the Mini RNeasy Kit (Qiagen, Germany) according to the manufacturer's instructions. First-strand cDNA was synthesized using the SuperScript Vilo cDNA synthesis Kit (cat. No. 11754050, Thermo Fisher, USA). Downregulation of MORN5 expression after injection was first confirmed using qPCR before further processing for PCR Array analysis.

Custom made Chick-bone plates (KRD, Czech Republic) were used for analysis of BMP pathway genes. The PCR arrays were performed in 12µl final reaction volumes containing SYBR Premix Ex Taq II (cat. No RR0821A, Takara, Japan) on LightCycler R 96 (Roche, Manheim, Germany) with preheating at 95◦C/30 min, followed by 45 cycles of 95◦C/5 s, 60◦C/20 s and 72◦C/15 s. Data were statistically evaluated by 11CT method with normalization against HPRT1 levels. In each PCR array plate, there were three technical replicates for 24 genes, and 2 technical replicates for an additional 13 genes.

# Statistical Analysis

All results were expressed as means ± standard deviations (SD) of three samples for each treatment and were compared by unpaired two-tailed Student's t-test for qPCR and PCR Array. Differences were considered to be significant at p < 0.05.

# RESULTS

# Spatiotemporal Gene Expression Pattern of MORN5 in Facial Prominences

First, we analyzed spatiotemporal expression pattern of MORN5 in individual prominences of chicken face. Facial prominences begin to form during early embryonic development. In situ hybridization showed no expression in chicken face at Hamburger-Hamilton (HH) stage 15 (50–55 h of incubation, **Figures 2A–C**) which is shortly after neural crest cells have entered the face. Later at stage HH17 (52-64 h of incubation, **Figures 2D–F**), MORN5 expression appeared in the caudal part

FIGURE 2 | Gene expression of MORN5 in early chicken face. (A–C) Frontal sections of chicken face at stage HH15. (D–F) Frontal sections of chicken face at stage 17. (D,E) In the ventral part of maxillary and mandibular prominences, there was no expression. (F) MORN5 expression gradually appeared dorsally in caudal part of maxillary region. (G–I) Frontal sections of chicken face at stage HH18. MORN5 expression was strong in maxilla and also in central part of each mandibular prominence (G). MdP, mandibular prominence; MxP, maxillary prominence. Scale bars = 100µm.

of the presumptive maxillary mesenchyme close to the maxillomandibular cleft. At stage HH18, the bulge of the maxillary prominence contained high levels of MORN5 transcripts (65– 69 h of incubation, **Figures 2G–I**). Expression was also detected in the dorsal (oral side) part of the mandibular prominences close to the maxillo-mandibular cleft (**Figures 2G,H**). At stage HH20 (70–72 h of incubation), there continued to be restricted expression in caudal and medial domains within the maxillary prominences (**Figures 3A–C**). In the mandibular prominences, there was expression in the cranial mesenchyme on either side of the midline groove (**Figures 3A–O**) with the exceptions of mesenchymal condensations of Meckel's cartilage (**Figures 3B,C**). At stage HH24 (4 days of incubation), maxillary prominence enlarged and strong MORN5 expression was present throughout the mesenchyme (**Figures 3E–G**). Mandibular expression was similar to stage HH20 (**Figures 2E–G**). Thus,

FIGURE 3 | Gene expression of MORN5 in later stages of chicken embryo. (A–C) ISH analysis in frontal sections of chicken head at stage HH20. There was strong expression in the maxillary prominence. The expression appeared in the cranial part in each mandibular prominence and it continued in dorsal direction. (B) No expression was observed in mesenchymal condensations and close to the fusion region of mandibular prominences. (E–G) MORN5 expression in frontal section of chicken head at stage HH24. MORN5 expression was strong in the mesenchyme of maxillary prominences. MORN5 expression was localized in the dorsal part of mandibular prominence, but not in mesenchymal condensation and close to the midline. (I–K) Frontal sections of chicken head at stage HH27 showed prominent expression in the maxillary prominence. There was weak expression in the globular process. Prominent expression was observed in rostral part of maxillary prominence and also in the mandibular prominence with the exception of midline. (M–O) Frontal sections of chicken head at stage 29 where beak is evident. MORN5 expression was localized in rostral part of maxilla and in fusion region. In the mandibular prominence, there was strong expression but not in the midline. (D,H,L,P) ISH analysis using sense probe. FNM, frontonasal mass; GP, globular process; mc, mesenchymal condensation; MdP, mandibular prominence; MxP, maxillary prominence. Scale bars = 100µm.

MORN5 is expressed in a restricted pattern in neural crestderived mesenchyme but not in epithelium. Sense probe did not show signal in the maxillary prominence (**Figures 3D,H,L,P**).

# MORN5 Expression in the Lip Fusion Zone at Later Stages

The next critical phase of facial morphogenesis is the fusion of the lip. Between stage HH27–29, the cranial-medial edges of the maxillary prominences meet the lateral corners of medial nasal prominences (globular processes) and fuse (Abramyan et al., 2015). At stage HH27 (5 days of incubation), MORN5 expression was observed for the first time in the corners of the frontonasal mass (globular processes, **Figures 3I,J**). Expression in the maxillary prominences was high in the rostral-medial corner just where fusion with the globular processes will take place. There continued to be expression in the mandibular prominences similar to stage 24 (**Figures 3I–K**). At stage HH29 (6 days of incubation), MORN5 expression was located in the region of lip fusion (**Figures 3M,N**) as well as in the mandible. This is the first stage where expression of MORN5 in Meckel's cartilage was detected (**Figure 3M**). Further confirmation of the restricted expression in the lip fusion zone is shown in other embryos cut in the frontal (**Figures 4A–C**) or transverse plane at stage HH29 (**Figures 4A–F**). Note that mesenchymal bridging has taken place by stage HH29, unifying the domains of expression of MORN5 in the globular processes and maxillary prominences (**Figures 4B,C**).

To quantify the relative levels of expression between the stages of development, we performed QPCR for evaluation of MORN5 expression level in each prominence at four different stages (HH15, 18, 20, 26). Since stage HH15, we did not observe any expression of MORN5 by ISH, this level of expression was chosen as the reference value for 11Ct analysis for individual prominences. In the maxillary prominence, MORN5 expression gradually increased during development with the peak level seen at stage HH20 (**Figure 5A**). In the mandibular prominence, we observed significantly increased expression at stage HH20 and 26 compared to stage HH15 embryos (**Figure 5B**). In the frontonasal mass, MORN5 expression is very low except of the globular processes we were surprised to see a statistically significant increase in expression of stage HH20 embryos (**Figure 5C**). In the section of in situ experiments, we could not detect MORN5 at stage HH20 (data not shown) therefore sensitivity of QPCR is greater than in situ hybridization. By stage HH27, there is expression of MORN5 in the in situ experiments; however, QPCR data did not pick up a significant expression level in stage HH26 embryos (**Figure 5C**). Some of the variability may be due to the dissection process and whether the globular process was included in all the samples. We did not compare expression levels between the facial prominences due to the experimental design.

# MORN5 Protein Expression in the Face

To correlate MORN5 protein distribution with MORN5 gene expression, we performed immunofluorescence staining. MORN5 protein was localized in developing chicken face at stage

in the area where edges of the maxillary prominences grow together with medial nasal prominence.  (E) Region of fusion had prominent MORN5 expression. (F) More caudal section (other sample). Scale bars = 100µm.

HH24, with the most prominent expression in individual cells in the maxillary and mandibular prominences (**Figures 6A–H**). Thus, only a subset of cells expressing MORN5 RNA expresses the protein. In positive control (adult chicken intestine), there was expected signal in Goblet cells, in the apical parts of enterocytes and in fibroblasts of the lamina propria (**Figures 6I–L**).

The specificity of the MORN5 antibody was also confirmed in HEK293T cells transfected with a MORN5-FLAG plasmid (**Figures 7A–C**). The staining of MORN5 and FLAG antibodies overlapped (**Figures 7A–C**). Similar to tissue section data, exogenous MORN5 protein was found in the cytoplasm in a punctate pattern (**Figures 7D,E**).

# Downregulation of MORN5 after Noggin a Retinoic Acid Treatment

Our study uncovered high levels of MORN5 expression in normal chicken embryos, however a previous study from

overexpression in comparison to stage HH15. (B) In the mandibular prominence, we observed significant expression at stage HH20 and 26. (C) Very low MORN5 expression was detected in the frontonasal mass at stage HH15 and 18, but at stage HH20 was MORN5 expression significantly increased.  Beads soaked in the bone morphogenetic protein antagonist, Noggin and retinoic acid (RA) synergistically induced transformation of the maxillary prominence into the frontonasal mass (Lee et al., 2001). The tissues from embryos induced to form this duplicated beak were profiled using microarrays. A significant downregulation of MORN5 expression was observed in all the treatment groups compared to controls treated with DMSO-Tris beads (-3.77 fold Noggin-DMSO treatment, -3.68-fold Noggin-RA, -2-fold after RA-Tris treatment) (Nimmagadda et al., 2015). We wanted to follow up this findings since it appeared that the RA and BMP pathways were upstream regulators of MORN5 and that possibly MORN5 was one of a set of genes mediating the beak duplication phenotype. First, we validated the array results using QPCR on maxillary tissues collected from treated and control embryos. We found a significant downregulation of MORN5 after Nogin-RA and Noggin-DMSO treatment compared to Tris-DMSO controls (**Figure 8A**). Next, we asked whether there were any spatial differences in MORN5 expression induced by the bead implants using in situ hybridization. Control embryos implanted with beads soaked in DMSO-Tris showed strong expression in the maxillary region and maxillo-mandibular cleft (**Figure 8B**). In contrast, no expression was observed in the maxillary prominence of Noggin-RA or Noggin-DMSO treated embryos. Interestingly, there was residual expression of MORN5 observed in embryos treated with RA-Tris located just under the epithelium of maxilla-mandibular cleft (**Figure 8B**).

# Downregulation of MORN5 by siRNA Altered Gene Expression of BMP and TGFβ Pathways Members

We had discovered that BMP activity was required for MORN5 expression but next wanted to investigate the genes that might be downstream of MORN5. As the first group of potential targets, we studied genes that are known to be in the BMP pathway. MORN5 expression in the maxillary prominence was downregulated to 75% of control levels following transfection with siRNA (2 rounds of transfection: at stage 20 and 24; **Figure 9A**). We used a PCR array that included 34 genes specific for the BMP pathway with HPRT1 acting as the reference control gene (Table S1).

Eight genes showed a statistically significant increase in their expression caused by partial MORN5 silencing (**Figure 9B**). These included ENG (Endoglin), Gdf2 (Growth differentiation factor 2, also BMP9), PLAU (plasminogen activator, urokinase), FST (Follistatin), Runx1 (Runt-related transcription factor 1), ID1 (Inhibitor of DNA binding 1), TGFβR2 (Transforming growth factor beta receptor 2) and TGFβ3 (**Figure 9B**). The most striking increase was seen with GDF2 (increased 3.5-fold). Statistically significant downregulation was observed only in the case of BMP5 (**Figure 9B**). It is interesting that MORN5 normally repressesID1, a transcription factor that positively regulates BMP signaling. Although levels of ID1 were increased, which should imply higher BMP signaling, there is also decreased expression of the BMP5 ligand. It is likely that cytoplasmic MORN5 indirectly regulates the expression of these genes and that further work is needed to determine the intermediate mediators of BMP and TGFβ signaling affected by MORN5.

# DISCUSSION

Here, we found spatially and temporally restricted expression of MORN5 in the face area during embryonic development suggesting its role in patterning of the maxillary prominences. Moreover, there was expression in the globular processes of frontonasal mass just before their fusion with the maxillary

the mandibular prominence, there is very low signal in the mesenchyme. MORN5 expression showed dispersed pattern of distribution. (A,C,E,G) DAPI staining. (I–L) Chicken intestine was used as a positive control with strong positivity in Goblet cells (gc), in the apical parts of enterocytes and also spotted expression in fibroblasts (fb). MdP, mandibular prominence; MxP, maxillary prominence.

prominences. Previously, a human genetics study found that MORN5 was associated with non-syndromic cleft lip with or without cleft palate (NSCLP) (Letra et al., 2010). We are the first to document expression of MORN5 in the relevant parts of the face undergoing lip fusion. In addition, there is strong expression on the medial sides of the maxillary prominences, the sites where palatal shelves will arise. While the chicken has a naturally cleft palate, it is interesting that MORN5 is expressed in the intermediate stages of palatal shelf formation. Based on previous microarray studies carried out on the chicken face, MORN5 came up twice, once as a maxillary enriched gene (Buchtová et al., 2010) and second as a differentially expressed gene following Noggin and RA bead implants (Nimmagadda et al., 2015). Taken together, the human genetic and chicken data suggest that MORN5 is an important maxillary patterning and possibly lip fusion gene. Thus, it would be worthwhile targeting MORN5 using mouse models and to include this gene in human NSLCP studies.

Complex signaling interactions coordinate the outgrowth of facial prominences to form the adult face. Some of factors have been previously identified by whole genome expression screens or by candidate gene mapping. The BMP signaling pathway regulates many cellular processes of craniofacial development and it is necessary for mesenchymal outgrowth of facial prominences. The expression of BMPs in chicken face was found at the time prior and during lip fusion (Ashique et al., 2002). BMP4 transcripts were previously detected in the epithelium of the globular processes of frontonasal mass and MORN5 expression underlays the same area however entirely in the mesenchyme. Also in the maxillary and mandibular prominences, epithelial BMP4 expression was described in parallel areas to mesenchymal MORN5. Furthermore, BMP2 and BMP7 were previously detected in the mesenchyme of both facial prominences. Maxillary and mandibular prominences express also several downstream target of BMP signaling. MSX1 is strongly expressed in the maxillary prominence but in slightly different pattern than MORN5 in the mandible (Shigetani et al., 2000; Fuchs et al., 2010). There are additional transcription factors that appear to overlap with MORN5 specifically in the frontonasal mass globular processes and maxillary prominences, TBX22, DLX5, and MSX2 (Higashihori et al., 2010). These transcription factors may regulate expression

**162**

antibodies. Cells expressing transgenic MORN5 were also MORN5-positive pointing to the specificity of the antibody. (D,E) In addition, a cytosolic spotted pattern of MORN5 expression was present in non-transfected cells. Scale bars = 100µm.

of MORN5. Interestingly, in the microarray study on beak duplicated embryos, TBX22 was upregulated following Noggin-RA treatment and TBX22 acts as a transcriptional repressor. It will be necessary to analyze whether MORN5 is a target of TBX22 which is a known clefting gene in humans (Kantaputra et al., 2011). It is interesting to note that the gene LHX6 which is located 3′ to MORN5 on the opposite strand was reported to be highly expressed in the chicken face (Washbourne and Cox, 2006). Expression of LHX6 begins in the maxillo-mandibular cleft at stage 18 similar to MORN5 (Washbourne and Cox, 2006). There is also striking similarity of expression of LHX6 in the globular processes and medial maxillary prominences at stage 27. This suggests that the two genes may share some common enhancers that drive expression in particular regions of the face.

MORN5 knockdown revealed indirect roles for this gene in controlling the expression of BMP and TGFB signaling pathways. Since we have shown MORN5 is a cytoplasmic protein it is unlikely that it is directly involved in regulating gene transcription. Further biochemical studies are needed to determine the exact function of MORN5 in the cell. Nevertheless, the RNA changes we observed suggest a subset of genes are dependent on MORN5 for their expression. We did not study the TGFB pathway in our bead implantation studies; therefore, the PCR array data extended our original findings on MORN5

FIGURE 8 | QPCR analysis and ISH of MORN5 expression after bead implantation. (A) QPCR analysis showed 1.68 times downregulation after RA-Tris, 3.03 after Noggin-DMSO and 2.59 after Noggin-RA treatment in comparison to control DMSO-Tris. (B) Control embryos implanted with beads soaked in DMSO-Tris had strong expression in maxillary region and maxilla-mandibular cleft. In RA-Tris treated embryos was very weak MORN5 expression. After Noggin-DMSO treatment, expression was rapidly decreased. No expression of MORN5 was observed after Noggin-RA treatment. Nog, Noggin; RA, Retinoic acid; TD, Tris-DMSO. Scale bars = 100µm. t-test; \*p < 0.05.

function. The most highly upregulated gene, GDF2, is known to associate with Endoglin (Castonguay et al., 2011) a glycoprotein located on cell surfaces that serves as a co-receptor for members of the Transforming growth factor-β superfamily (Cheifetz et al., 1992). This suggests that activity of TGFβ family is normally repressed by MORN5. Other changes such as the increase in the antagonist FST (Follistatin) may indicate that MORN5 operates in another way to regulate TGF signaling. MORN5 may normally repress this antagonist, which binds members of the TGFβ superfamily with a particular focus on activin (Lambert-Messerlian et al., 2007). We have shown that FST does not induce skeletal changes in the palate as compared to Noggin (Celá et al., 2016) therefore FST regulation by MORN5 may serve different functions outside of facial morphogenesis. Several other genes known to be in the TGFB pathway and essential for mouse palate development were upregulated including RUNX1 (Yamashiro et al., 2002), TGFβ3 (Cui et al., 1998) and ID1 (Rice et al., 2005). In summary, we discovered that MORN5 is involved in TGFB signaling at all levels. In conclusion, BMP signaling is required for MORN5 expression and reduction of MORN5 derepresses several genes in the BMP and TGFβ signaling pathways. Furthermore, MORN5 has two potential roles in facial patterning, to specify

# REFERENCES

maxillary identity and to regulate lip fusion that warrant further study in animal models.

# AUTHOR CONTRIBUTIONS

MB, JR, and PC conceived the study. PC, MH, KF, MK conducted the experiments. PK provided intellectual contribution. PC, JR, and MB wrote the manuscript. All authors reviewed and approved the final manuscript.

# ACKNOWLEDGMENTS

This study was supported by the Grant Agency of the Czech Republic (14-37368G to MB lab), Agency for Healthcare Research of the Czech Republic (15-33232A to PK lab) and institutional support (RVO:67985904).

# SUPPLEMENTARY MATERIAL

The Supplementary Material for this article can be found online at: http://journal.frontiersin.org/article/10.3389/fphys. 2016.00378

fetal palate tissues. Pediatr. Dev. Pathol. 10, 436–445. doi: 10.2350/06-05- 0087.1

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2016 Cela, Hampl, Fu, Kunova Bosakova, Krejci, Richman and Buchtova. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# MicroRNA Profiling during Craniofacial Development: Potential Roles for Mir23b and Mir133b

Hai-Lei Ding1 † , Joan E. Hooper 2 , Peter Batzel 3 , B. Frank Eames 3, 4, John H. Postlethwait 3 , Kristin B. Artinger 1 \* and David E. Clouthier 1 \*

### Edited by:

Thimios Mitsiadis, University of Zurich, Switzerland

### Reviewed by:

Ralf Kist, Newcastle University, UK Brad A. Amendt, The University of Iowa, USA Claudio Cantù, University of Zurich, Switzerland

### \*Correspondence:

Kristin B. Artinger<EMAIL_ADDRESS>David E. Clouthier<EMAIL_ADDRESS>
### †Present Address:

Hai-Lei Ding, Laboratory of Anesthesiology, Xuzhou Medical College, Xuzhou, Jiangsu, China

### Specialty section:

This article was submitted to Craniofacial Biology, a section of the journal Frontiers in Physiology

Received: 01 December 2015 Accepted: 21 June 2016 Published: 14 July 2016

### Citation:

Ding H-L, Hooper JE, Batzel P, Eames BF, Postlethwait JH, Artinger KB and Clouthier DE (2016) MicroRNA Profiling during Craniofacial Development: Potential Roles for Mir23b and Mir133b. Front. Physiol. 7:281. doi: 10.3389/fphys.2016.00281 1 Department of Craniofacial Biology, School of Dental Medicine, University of Colorado Anschutz Medical Campus, Aurora, CO, USA, 2 Department of Cell and Developmental Biology, School of Medicine, University of Colorado Anschutz Medical Campus, Aurora, CO, USA, 3 Department of Neuroscience, University of Oregon, Eugene, OR, USA, 4 Department of Anatomy and Cell Biology, University of Saskatchewan, Saskatoon, SK, Canada

Defects in mid-facial development, including cleft lip/palate, account for a large number of human birth defects annually. In many cases, aberrant gene expression results in either a reduction in the number of neural crest cells (NCCs) that reach the frontonasal region and form much of the facial skeleton or subsequent failure of NCC patterning and differentiation into bone and cartilage. While loss of gene expression is often associated with developmental defects, aberrant upregulation of expression can also be detrimental. microRNAs (miRNAs) are a class of non-coding RNAs that normally repress gene expression by binding to recognition sequences located in the 3′ UTR of target mRNAs. miRNAs play important roles in many developmental systems, including midfacial development. Here, we take advantage of high throughput RNA sequencing (RNA-seq) from different tissues of the developing mouse midface to interrogate the miRs that are expressed in the midface and select a subset for further expression analysis. Among those examined, we focused on four that showed the highest expression level in in situ hybridization analysis. Mir23b and Mir24.1 are specifically expressed in the developing mouse frontonasal region, in addition to areas in the perichondrium, tongue musculature and cranial ganglia. Mir23b is also expressed in the palatal shelves and in anterior epithelium of the palate. In contrast, Mir133b and Mir128.2 are mainly expressed in head and trunk musculature. Expression analysis of mir23b and mir133b in zebrafish suggests that mir23b is expressed in the pharyngeal arch, otic vesicle, and trunk muscle while mir133b is similarly expressed in head and trunk muscle. Functional analysis by overexpression of mir23b in zebrafish leads to broadening of the ethmoid plate and aberrant cartilage structures in the viscerocranium, while overexpression of mir133b causes a reduction in ethmoid plate size and a significant midfacial cleft. These data illustrate that miRs are expressed in the developing midface and that Mir23b and Mir133b may have roles in this developmental process.

Keywords: craniofacial development, neural crest cell, mouse, zebrafish, RNA duplex, facebase

# INTRODUCTION

Morphogenesis of the vertebrate face is a complex event requiring coordination among a variety of signaling cascades. Human craniofacial birth defects occur at a world-wide rate of 1:800 births (Schutte and Murray, 1999; Spritz, 2001), with structural defects often resulting from failure of spatio-temporal integration of these signaling cascades. While the vertebrate mid-face has a complex embryonic origin, most facial birth defects result from disruption of cranial neural crest cell (NCC) patterning and differentiation (Chai and Maxson, 2006; Knight and Schilling, 2006; Walker and Trainor, 2006; Dixon et al., 2011; Clouthier et al., 2013). Cranial NCCs arise at the border between the neural and non-neural ectoderm and subsequently migrate to the frontonasal region (Le Douarin, 1982; Bronner-Fraser, 1995). These cells eventually give rise to most of the bone, cartilage, and connective tissue of the mid-face and neck (Couly et al., 1996; Köntges and Lumsden, 1996). NCC development relies on intricate regulation of patterning cues within specific boundaries of the head. While boundaries are established through continuous refinement of gene expression, the mechanisms required for this refinement are less clear.