Rbms3 functions in craniofacial development by posttranscriptionally modulating TGF-β signaling

RNA processing, localization, and stability from the nucleus to the cytoplasm is coordinated by a multitude of RNA-binding proteins (RBPs) and microRNAs (Keene, 2007). Although the vertebrate genome encodes for hundreds of RBPs, only a small fraction of RBP targets and regulatory roles are known to date (Halbeisen et al., 2008; Hogan et al., 2008).

Neural crest cells are a transient embryonic stem cell population that contributes to important derivatives in vertebrate embryos, including the craniofacial skeleton and peripheral nervous system. Although many critical events in neural crest development are transcriptionally regulated (Sauka-Spengler and Bronner-Fraser, 2008), it is becoming increasingly clear that posttranscriptional modifications, such as SUMOylation (Taylor and LaBonne, 2005) and RNA stabilization/localization, also play important roles (Hogan et al., 2008). However, few studies have examined the role of RNA stabilization in developmental processes.

Interestingly, defects in RNA processing can lead to neural crest-related craniofacial abnormalities. For instance, in Treacher Collins Syndrome, disruption of Tcof1, which is involved in ribosome biosynthesis, leads to craniofacial abnormalities in mice (Dixon et al., 2006). Given that cranial neural crest (CNC) anomalies are among the most common of human birth defects (McKeown and Bronner-Fraser, 2008), it is clearly important to understand the developmental role of RNA stabilization therein.

To this end, we describe the expression and developmental function of an RBP Rbms3 in craniofacial morphogenesis. Isolated in a gene and protein trap screen in zebrafish (Trinh et al., 2011), the Rbms3 RBP belongs to the small family of c-myc single-stranded binding proteins (MSSPs; Penkov et al., 2000). MSSPs contain two ribonucleoprotein (RRM) domains that were originally described in RBPs (Siomi et al., 1993) and required for c-myc promoter binding (Takai et al., 1994). Previously, Rbms3 was shown to exhibit tumor suppressor function via regulation of c-Myc (Li et al., 2011) and to bind/stabilize RNA in vitro (Penkov et al., 2000; Fritz and Stefanovic, 2007; Lu et al., 2012). However, the in vivo function for Rbms3 during development has remained unexplored.

Here, we report that Rbms3 is expressed transiently in condensing CNC in the pharyngeal arches (Penkov et al., 2000). Because our protein trap screen labels endogenous Rbms3, we find that it is predominantly expressed in the cytoplasm of CNC, which suggests that its primary function is in RNA metabolism rather than transcriptional regulation of c-Myc. Embryonic knockdown of rbms3 results in severe craniofacial defects and a reduction in activated Smad2, a downstream effector of TGF-β receptor (TGF-βr) signaling. We present evidence that Rbms3 binds to and stabilizes Smad2 transcripts. Significantly, our results suggest that zebrafish Rbms3 is a novel posttranscriptional regulator of one of the major cartilage differentiation effectors, the TGF-βr pathway. Other novel targets for Rbms3 also include cell cycle regulators (rac1 and cyclin D1) and smad1, which suggests that Rbm3 is a global chondrogenic regulator important for driving CNC down a chondrogenic lineage.

In the Rbms3 trapped line, Gt(rbms3-citrine)^ct30a (Trinh et al., 2011), the insertion of the Citrine tag was localized to intron 11. 5′ rapid amplification of cDNA ends (RACE) was performed to retrieve full-length and two C-terminal variants (Fig. 1 A). The full-length rbms3 (exons 2–14) is a 415–amino acid protein; exons 3–7 code for two putative RNA binding domains (RRM).

The Citrine in Gt(rbms3-citrine)^ct30a is flanked by splice acceptor and donor sites and is incorporated as an artificial exon into the Rbms3 protein (Trinh et al., 2011). Homozygous Gt(rbms3-citrine)^ct30a fish are viable, suggesting that Citrine incorporation does not interfere with protein function. Citrine (labeling all three rbms3 variants) was observed in the cranial mesenchyme, pharyngeal arches neurons within the neural tube, the isthmus of the midbrain-hindbrain, fin buds, and tail bud/somites (Fig. 1 C). By 48–60 h postfertilization (hpf), Citrine was absent in the tail. At the 10 somite stage (ss; 14 hpf), citrine is expressed in the hatching gland, the mesoderm of the tail bud and somites. Expression for rbms3 was similar to citrine, which confirmed that tagged and endogenous Rbms3 expression patterns overlapped at 24–60 hpf. By 48 hpf, expression in the heart was also apparent (Fig. 1 D). Quantitative PCR (QPCR) analysis showed that onset of rbms3 expression was around 6–10 hpf (gastrulation to 3 ss; Fig. 1 E). By 16–18 hpf, rbms3 was also expressed in the nervous system and cranial mesenchyme (Fig. S1).

CNC emigrate from the neural tube (5–10 ss/11.5–14 hpf in zebrafish) in a segmental fashion, invade the pharyngeal arches, and form prechondrogenic mesenchyme (Schilling and Kimmel, 1994; Trainor and Krumlauf, 2001; Knecht and Bronner-Fraser, 2002). In Gt(rbms3-citrine)^ct30a; Tg(sox10(−7.2):mrfp) fish at 31 hpf, Rbms3 (green) colocalizes with CNC cells (red) in the pharyngeal arches (Fig. 2, top and middle) in the cytoplasm (Fig. 2, middle), which is consistent with a role during RNA metabolism (Penkov et al., 2000; Fritz and Stefanovic, 2007; Lu et al., 2012). By 72 hpf, differentiated cartilage lack Rbms3, but surrounding tissue maintain expression (Fig. 2, bottom, arrowheads). The expression pattern of Rbms3 in prechondrogenic crest (PCC) suggests it may function transiently during cartilage differentiation.

We designed two morpholino oligonucleotides (MOs), a translation blocking translation start site rbms3 morpholino (Atg MO) and a splice site rbms3 morpholino (Sp MO), to perform rbms3 loss-of-function analysis during zebrafish development. Both MOs efficiently blocked Rbms3 production when injected into the Gt(rbms3-citrine)^ct30a line, as analyzed by Western blotting using an anti-GFP antibody (Fig. 3 A). The Sp MO was found to skip exon 3 (Fig. 3 B), creating a truncated transcript; sequencing of other minor PCR products also produced other truncated transcripts of rbms3 (Fig. 2 A). Survival of morphant larvae was reduced past 4 d postfertilization (dpf).

Compared with controls, rbms3 morphants displayed cardiac edema, with varying penetrance, smaller body size, a hooked head, and craniofacial defects by 3 dpf (Fig. 3 B).

Rbms3 morphants displayed major disruption to neurocranial and pharyngeal cartilage elements (Fig. 3 C; see Fig. 6 A). About 80–90% of morphants displayed malformed mandibular and hyoid cartilage elements, and with varying penetrance, the cartilage of pharyngeal arches 3–7 was either entirely missing or malformed (Fig. 3, C and D). Co-injecting full-length rbms3 with Atg MO was able to fully or partially rescue the severe phenotype (Fig. 3 E; 74% severe phenotype with Atg MO, n = 97, vs. 20% severe phenotype with Atg MO coinjected with rbms3, n = 193, P < 0.001).

Tissue interactions and signaling during gastrulation are essential for proper CNC formation in the embryo (Rogers et al., 2012). The cartilage abnormalities observed in rbms3 morphants may be caused by early patterning defects, which subsequently affect early CNC formation and migration (Fig. 4).

Analysis of the neural plate boarder markers snail1b and foxd3 at 3 ss (10 hpf) showed no differences between controls and rbms3 morphants (Fig. 4 A). No significant changes were also observed in expression of CNC markers sox10 and foxd3 at 10 ss (Fig. 4 A).

Rbms3 is expressed in the isthmus at the midbrain–hindbrain junction. To rule out the possibility that the cartilage defects in rbms3 morphants may be secondary to midbrain–hindbrain patterning defects, we analyzed markers of forebrain (pax6a), isthmus (fgf3/pax2a), and hindbrain (hoxa2) at 20 ss (19 hpf). No significant differences between controls and rbms3 morphants were observed (Fig. 4 C). Collectively, the results suggest that early patterning in rbms3 morphants is normal, and that the cartilage phenotype is not a secondary defect.

The cartilage phenotype may result from defects in PCC condensation and/or differentiation because early CNC development was normal in rbms3 morphants. Analysis of dlx2a revealed no significant differences in PCC condensation at 24 hpf in the pharyngeal arches (Fig. 5 A). At this stage, Rbms3 is only weakly expressed and later up-regulated in the pharyngeal arches by 31 hpf (Fig. 1 C).

In contrast, significant effects on PCC and its derivatives were observed in rbms3 morphants from 31 hpf to 72 hpf. Barx1, sox9a (PCC/chondrocytes), and col2a1a (mature chondrocytes) domains of expression were reduced/missing in morphants (Fig. 5 A). QPCR analysis at 48 hpf showed a reduction in sox9a and barx1 levels in rbms3 morphants (P < 0.005), yet no significant difference in prrx1a (cranial mesenchyme) was observed (Fig. 5 B; P = 0.14), which suggests that in rbms3 morphants, CNC-derived mesenchyme in the pharyngeal arches is specifically affected. Examination of runx2b (osteogenic lineage) also showed altered expression in rbms3 morphants at 60 hpf (Fig. 5 A).

Nonspecific MO effects may indirectly cause craniofacial defects through the activation of the p53–apoptotic pathway (Robu et al., 2007). Tg(sox10(−4.7):egfp) embryos coinjected with Atg MO and p53 MO failed to rescue the rbms3 morphant cartilage phenotype, which suggests that the observed craniofacial defects were specifically caused by loss of rbms3 (Fig. 6, A and B). Moreover, no significant difference in the numbers of Caspase-3⁺; EGFP⁺ cells were observed in the pharyngeal arches at 31 hpf between controls and rbms3 morphants (Fig. 6 C).

In contrast, cell proliferation was markedly reduced in rbms3 morphants at 31 hpf (Co MO, n = 31; Atg MO, n = 27) and 48 hpf (Co MO, n = 33; Atg MO, n = 34), as assayed by PH3 expression in Tg(sox10(−4.7):egfp) fish (Fig. 6 D; P < 0.001). Comparison of sox10:egfp⁺ pharyngeal arch domains between controls and rbms3 morphants at 24 hpf showed no significant differences (Fig. 6 E), which suggests that the proliferative differences observed between treatments at 31 hpf are not caused by significant loss of pharyngeal arch cells at early stages. Thus, Rbms3 functions in PCC by regulating cell proliferation.

The CNC phenotype observed in rbms3 morphants is reminiscent of knockdown of mouse TGF-βr2 in Wnt1-Cre mice (Ito et al., 2003; Wurdak et al., 2005). In both cases, the primary deficit is in the capacity of PCC to differentiate along the chondrogenic lineage because of an inability to maintain Sox9 expression. Furthermore, Smad2/3, effectors of the TGF-βr2 pathway, promote Sox9-dependent transcriptional activity on the a1(II) collagen gene (Col2A1) enhancer and induce chondrogenesis in vitro (Furumatsu et al., 2005).

In zebrafish, smad2 is expressed by PCC at 26 hpf (Müller et al., 1999) and 31 hpf (Fig. 7 A). At 48 hpf, there was a reduction in transcription of both smad2 and tgf-βr2a receptor in the pharyngeal arches of rbms3 morphants (Fig. 7 B; P < 0.005). This also corresponded to reduced Smad2 protein levels at 48 hpf in rbms3 morphant arches (Fig. 7 C); quantitation of band intensity revealed a ratio of 1:0.3 ± 0.05 (n = 3) in controls/morphants after normalization to α-tubulin (Fig. 7 C). A reduction in phosphorylated Smad2 (pSmad2, reflecting active Tgf-β signaling) levels in rbms3 morphants at 31 hpf and 48 hpf was also observed in the pharyngeal arches (Fig. 7 C). These results further support the hypothesis that the rbms3 morphant cartilage phenotype may be caused by defects in Tgf-βr signaling and that smad2 is a putative Rbms3 target.

We next tested whether constitutively activated smad2 (smad2ca), which signals independently of TGF-βr, was able to rescue the phenotype. Because injecting smad2ca alone produces a dorsalized phenotype (Müller et al., 1999; Dick et al., 2000), we injected RNA below the threshold of an observable phenotype and excluded any embryos exhibiting axis defects from analysis (Fig. 7, D and E). To improve survival of smad2ca and Atg MO coinjected embryos, we also reduced Atg MO concentration (∼150 µM), which in turn reduced the severe phenotype to ∼60%. Co-injecting smad2ca RNA with Atg MO nearly doubled the number of wild-type larvae (∼40%, n = 321 vs. Atg MO alone, ∼20%, n = 251; Fig. 7 D; P < 0.001). Therefore, the rbms3 morphant cartilage defect is due, at least in part, to reduction in Tgf-βr/Smad2 signaling.

In PCC, Rbms3 may regulate the stability of smad2 transcript and thus affect the pool of protein available for signaling, as evidence suggests that Rbms3 can bind to the 3′ UTRs (Fritz and Stefanovic, 2007; Lu et al., 2012). To test if binding sites were harbored within the 3′ UTR of zebrafish smad2, embryos were coinjected with full-length Rbms3 and destabilized egfp (d1egfp) mRNA fused to smad2 3′ UTR (d1egfp-smad2 3′ UTR; Fig. 8, A–C). Comparing fish at 22–24 hpf injected with a truncated C-terminal version of rbms3 (dRRM-rbms3; Fig. 8 A) versus full-length rbms3 showed higher d1EGFP protein and transcript levels (Fig. 8, B and C), which suggests that the d1egfp-smad2 3′ UTR transcript half-life was extended with full-length Rbms3 (1.7 ± 0.3-fold increase, P = 0.05; Fig. 8 C, asterisk). In contrast, no difference in fluorescence was observed with smad2 3′ UTR in the reverse orientation (smad2 3′ UTR R), since it would not bind directly/indirectly to Rbms3 (P > 0.05) when coinjected with either rbms3 or dRRM-rbms3 (Fig. 8, B and C). The last 150 bp of smad2 3′ UTR was found to harbor Rbms3-mediated stabilization sites (2.8 ± 0.2 fold, P < 0.05; Fig. 8 C, asterisk). However, the first 200 bp region of smad2 3′ UTR was found to destabilize the reporter transcript (Fig. 8 C; *, P < 0.05). Additionally, Rbms3 has no effect on sox9a 3′ UTR (P = 0.7; Fig. 8 C). To further extend these findings, we tested d1egfp-smad2-150bp 3′ UTR stability in the absence of endogenous Rbms3. Co- or Atg MO–treated embryos were coinjected with d1egfp-smad2-150bp 3′ UTR, and Atg MO–treated embryos had significantly less d1egfp transcript (*, P < 0.05) and protein levels compared with controls at ∼22–24 hpf (Fig. 8 D). Collectively, these results suggest that Rbms3 stabilizes smad2 transcripts by acting on the last 150 bp of the smad2 3′ UTR.

To test for direct binding, we performed RNA immunoprecipitation (RIP). Either full-length Rbms3-Myc or truncated Rbms3 lacking RNA binding domains (dRRM-Rbms3)-Myc was injected into zebrafish embryos, and c-Myc antibody was used to pull down associated mRNA transcripts (Fig. 8, E and F). Analysis of the Tgf-βr pathway components in the Rbms3-Myc pull-downs revealed higher enrichment for smad2, smad3a, and smad3b compared with tgf-β receptors (compare purple with blue; Fig. 8 F). Other transcripts that were significantly enriched (more than fivefold; Fig. 8 F, broken red line) for full-length Rbms3 were cell cycle regulators (cyclin D1 and rac1) and smad1.

We tested specificity of binding for Rbms3 and found very little enrichment for the following transcripts. These transcripts overlapped with observed domains of expression for endogenous Rbms3 around 10–24 hpf: wnt3a, expressed in the isthmus and other neural tissues (Clements et al., 2009); the adhesion molecule cad2a (n-cad), ubiquitously expressed yet enriched in neural tissues (Harrington et al., 2007); and transcription factors (myod and sox9a), both expressed in somites, with the latter also being expressed in the otic placodes (Fig. 8 F; Weinberg et al., 1996; Yan et al., 2005). These results suggest that posttranscriptional binding and stabilization of smad2 and other transcripts associated with the Tgf-βr signaling pathway may be a direct mechanism by which Rbms3 regulates PCC differentiation into cartilage.

From a protein trap screen (Trinh et al., 2011), we isolated zebrafish rbms3, an RBP that belongs to the small family of MSSP (Penkov et al., 2000). For the first time, we have analyzed the in vivo developmental function for Rbms3. Here we report that Rbms3 is predominantly restricted to the cytoplasm and that rbms3 morphants exhibit severe craniofacial defects. Significantly, our results highlight a posttranscriptional role for Rbms3 in regulating one of the major pathways that promote chondrogenesis, the TGF-βr pathway.

Knockdown of rbms3 with two different MOs resulted in severe craniofacial defects (Fig. 3). Early neural crest development proceeded normally (Fig. 4). Although CNC migration to and condensation within the pharyngeal arches appeared unaffected in rbms3 morphants, pharyngeal arch expression domains of barx1, sox9a, col2a1a, and runx2b were severely reduced or affected (Fig. 5), which suggests a major role for Rbms3 during chondrogenesis.

CNC migration and condensation relies on interactions between CNC and their environment (Trainor and Tam, 1995; Trainor and Krumlauf, 2001; García-Castro et al., 2002; Trainor et al., 2002; Crump et al., 2004; Eberhart et al., 2006). Citrine tagged Rbms3 is expressed in the pharyngeal arches and the paraxial cranial mesoderm during CNC condensation in the arches (Figs. 1 and 2). High-resolution analysis in the Tg(sox10(−7.2):mrfp) line revealed that Rbms3 is transiently expressed by PCC and excluded from differentiated cartilage while being maintained by surrounding tissues (Figs. 1 and 2). Collectively, the results suggest that Rbms3 promotes differentiation of PCC to cartilage in a cell-autonomous manner.

A fine balance between cell proliferation and apoptosis is essential for proper cranial cartilage formation (Plaster et al., 2006; Jones et al., 2008). Rbms3 morphants show a reduction in proliferating CNC (Fig. 6). This may be caused by (a) proliferative/apoptotic defects in the neuroepithelium, which influence the number of emigrating CNC; or (b) proliferative/apoptotic defects in the condensing ectomesenchyme. The former seems unlikely because Rbms3 is not expressed in the neuroepithelium during CNC induction. Additionally, condensation of CNC in the pharyngeal arches at 24 hpf is unaffected, and co-injecting rbms3 MO with p53 MO does not rescue the cartilage phenotype (Fig. 6). This differs from the flathead mutants, where decreased proliferation of CNC cells at 48 hpf and activation of the p53 pathway lead to a deficit in zygotic DNA polymerase δ1, thus affecting cartilage morphogenesis. In contrast, in rbms3 morphants, lack of proliferation does not activate the p53 pathway or increase apoptosis (Fig. 6).

We hypothesized that Rbms3 may modulate the TGF-βr pathway. Consistent with this possibility, rbms3 and smad2 (an effector of the TGF-βr pathway) exhibit overlapping expression during CNC condensation in the pharyngeal arches (Figs. 1 and 7). Mice conditionally deficient for the TGF-βr2 pathway exhibit a similar cartilage phenotype to rbms3 morphants (Ito et al., 2003; Wurdak et al., 2005), although previous work in these mutant mice only concentrated on palatogenesis and no comment was made on the more posterior cartilage defects. As observed in Tgf-βr2^fl/fl; Wnt1-Cre mice, rbms3 morphants show a reduction in sox9a expression (Fig. 5) and cell proliferation of PCC (Fig. 6), whereas pre-migratory and early migrating CNC appear (Fig. 4) normal, and cell death is unaffected (Fig. 6).

The phenotypes observed in other TGF-βr pathway mutants are not consistent with the overall rbms3 MO phenotype. In tgf-βr3 morphants, survival of early neural crest is affected, resulting in a secondary cartilage phenotype (Cheah et al., 2010). The requirement for Tgf-βr1 (Dudas et al., 2006) is significantly different from Tgf-βr2 during chondrogenesis, suggesting that Tgf-βr1 may participate in diverse signaling cascades during craniofacial development. Furthermore, signaling via Tgf-βr1 is not necessary for PCC proliferation but is required for survival (Dudas et al., 2006). Although it is true that Tgf-βr1 can activate pSmad2, we suggest that reduction in observed Smad2 signaling is primarily caused by the Tgf-βr2–pSmad2 pathway deficiency.

Early PCC proliferation is likely to be regulated mainly through the TGF-βr receptor pathway in a Smad-independent manner. Although Tgf-βr2 null mice exhibit a cell proliferation defect (Ito et al., 2003), the mechanism by which this occurs has not been established during chondrocyte differentiation (Fig. 9 A, broken line). Previously, the small GTP-binding proteins Cdc42/Rac1 were shown to promote chondrocyte differentiation in mesenchyme condensations by promoting Sox9, Col2A, and N-cadherin expression (Woods et al., 2007). Additionally in a cell-dependent context, Cdc42 can activate cell proliferation in a Smad-independent manner after TGF-βr activation in fibroblasts (Wilkes et al., 2003). Collectively, the decreased proliferation observed in rbms3 morphants is hypothesized to be a direct consequence of down-regulating components of the TGF-βr pathway, likely in a Smad-independent manner. In support of this, we find that the Tgf-β receptor transcripts are enriched after immunoprecipitation for Rbms3 (Fig. 8). The direct interactions between TGF-βr signaling, Cdc42/Rac1, and cell cycle effectors remain unclear, but this study demonstrates that rac1 and cyclin D1 can be independently regulated by Rbms3 (Fig. 8).

Although the cell proliferation defect is likely to be mainly Smad independent yet TGF-βr signaling dependent, we suggest that an inability to generate cartilage and bone is caused by a Smad2 signaling deficiency in PCC progenitors (Fig. 9). Atg MO treatment resulted in reduced levels of pSmad2 (indicative of active Tgf-βr signaling), probably because of reduced Smad2 transcript and total protein levels in morphants. Importantly, coinjection of capped smad2ca RNA with rbms3 MO significantly rescued the severe cartilage phenotype (Fig. 7), further supporting the notion that Rbms3 modulates the Tgf-βr2 pathway during cartilage formation (Fig. 9).

The Tgf-βr pathway is highly modulated during development to control the precise timing and duration of signaling (Liu, 2003; Massagué et al., 2005). Steady-state levels of receptor Smads (such as Smad2) are regulated by the ubiquitin pathway in a signal-dependent and -independent manner (Izzi and Attisano, 2004). Another possible mechanism for regulating protein levels is by altering the levels of transcript either by modulating the half-life of mRNA or altering accessibility of mRNA to translational machinery through compartmentalization of transcripts to different regions within the cell. This is facilitated by a multitude of RBPs and microRNAs (Lasko, 2003; Keene, 2007). Evidence from the in vivo stability assay suggests that the smad2 3′ UTR harbors Rbms3-binding sites (Fig. 8), thereby lending further support to the idea that smad2 levels and thus Smad2 steady-state protein levels are regulated by Rbms3. The region localized to the last 150 bp of smad2 3′ UTR is sufficient to stabilize the reporter transcript. However, the first 200 bp of the smad2 3′ UTR caused the reporter transcript to be destabilized, suggesting that there are “functional” domains within the smad2 3′ UTR that both positively and negatively regulated by Rbms3. It may also reflect Rbms3 dependence for RNA secondary structure to facilitate binding of the Rbms3–RNP complex (Hiller et al., 2006; Li et al., 2010). The requirement for endogenous Rbms3 to stabilize smad2 150 bp 3′ UTR and significant enrichment of smad2/3 transcripts after RIP for Rbms3 further provides evidence that Smad2 is a direct target (Fig. 8). Smad3 also participates during chondrocyte differentiation (Ferguson et al., 2000; Yang et al., 2001) and is thus likely to be regulated directly by Rbms3.

The reduction observed in sox9a expression (Fig. 5) is also consistent with the hypothesis that Rbms3 is a modulator of the TGF-βr pathway. Sustained Sox9 expression by condensing prechondrogenic CNC requires active TGF-βr2 signaling (Ito et al., 2003; Wurdak et al., 2005). Analysis of Sox9^−/− chimeric mice reveals that loss of Sox9 expression prevents PCC from differentiating into cartilage, which was formed entirely from wild-type cells, whereas Sox9^−/− cells, which also lacked Col2a1, were excluded (Bi et al., 1999). Conditional inactivation of Sox9 during CNC condensation maintains these cells in a mesenchymal state, as observed by a lack of Col2A1-positive cartilage extracellular matrix production (Akiyama et al., 2002). This is consistent with the finding that Smad2/3, effectors of the TGF-βr2 pathway, promote Sox9-dependent transcription of Col2A1 in vitro (Furumatsu et al., 2005). Further support of the notion that loss of Smad2 is the primary defect and that sox9a loss in rbms3 morphants is a secondary outcome comes from the finding that the sox9a 3′ UTR is unresponsive to Rbms3 and is not highly enriched during RIP (Fig. 8).

While TGF-βr signaling is a major pathway that promotes chondrogenesis, other signaling pathways also contribute to this process (Walshe and Mason, 2003a; Wada et al., 2005; Yoon et al., 2005; Eberhart et al., 2006; Sakai et al., 2006; de Frutos et al., 2007; Oka et al., 2008). In support of this, our data suggest that smad2ca alone is not sufficient to fully rescue the rbms3 morphant phenotype (Fig. 7). Mice deficient for Bmpr1a and Bmpr1b exhibit defects in PCC proliferation and differentiation (Yoon et al., 2005). Additionally, a zebrafish transgenic reporter for Smad1/5/8 and expression of smad1 also highlight a requirement for BMP signaling during chondrocyte development (Alexander et al., 2011; Laux et al., 2011). Collectively, we suggest that the cell proliferation defect and the loss in sox9a expression could be partially caused by a loss of BMP–Smad1 signaling and rac1 regulation by Rbms3 (Fig. 8).

Integrating these data, we propose a model for Rbms3 function during prechondrogenic CNC development in which Rbms3 posttranscriptionally regulates components of the Tgf-βr pathway to influence the proliferative state and initial differentiation of prechondrogenic CNC (Fig. 9). Previously, Prx1 and Ptf1a mRNA was demonstrated to be up-regulated by Rbms3 through direct interaction with the Prx1/Ptf1a 3′ UTR sequence (Fritz and Stefanovic, 2007; Lu et al., 2012). In a similar manner, we predict that Rbms3 influences transcript stability of Tgf-βr pathway components (particularly smad2) and therefore transiently influences the timing and duration of Tgf-βr signaling within CNC cells. Additionally, Rbms3 is likely to also regulate other components of the Smad2-independent pathway, such as cell cycle regulators and the BMP–Smad1 pathway. Once these prechondrogenic cells become committed to form round chondrocytes, rbms3 expression is shut down in these cells. Under normal conditions, PCC are likely to be sensitive to levels of Tgf-βr/BMP signaling and require a certain threshold of signaling to drive them down the chondrogenic lineage. In rbms3 morphants, a reduction in the pool of smad2 transcripts caused by reduced levels of Rbms3 is likely to directly influence the pool of Smad2 protein available in the cytoplasm that is able to respond to external Tgf-β signals, thereby maintaining these cells in a mesenchymal state. Once prechondrogenic cells are driven into the cartilage lineage, we predict that the function of Rmbs3 in the CNC is complete. At this juncture, perhaps other RBPs or other mechanisms help maintain these cells in a committed chondrogenic state.

Wild-type (AB or AB/TL background) and transgenic zebrafish lines (see below) were maintained according to institutional animal care and use protocols. Embryos were collected in egg water, maintained at 28°C, and staged according to Kimmel et al. (1995) based on somite number or landmark structural features.

Gt(rbms3-citrine)^ct30a were maintained in a mixed AB/TL background. Previously published Tg(sox10(−7.2):mrfp) (Kucenas et al., 2009) or Tg(sox10(−4.7):egfp) (Dutton et al., 2008) were maintained in the AB background.

To retrieve the full-length rbms3, wild-type 42–48 hpf cDNA library was used to amplify full length and variants. 5′ RACE was performed using the Generacer kit (Invitrogen) using 5′-AGAGCGTTCCAGCCTCATCACAGGACGC-3′ primer. 3′ RACE was performed as described previously (Jayasena et al., 2011) using a 5′-AGAGCGTTCCAGCCTCATCACAGGACGC-3′ primer. For subcloning of full-length rbms3 containing 5′ UTR and open reading frame into pCS2⁺ at the Cla1–Xba1 sites are: 5′-GATCCCATCGATACAGCCAGAAGGATTCCAGCTAC-3′ and 5′-TGAGGAGTAGAACCCGTTCTGCATTG-3′.

MOs were injected either into AB or AB/TL wild-type background. The translation start site MO (Atg MO) is as follows: 5′-CGTTTGCCCATGTAGCTGGAATCCT-3′. The splice MO (Sp MO) sequence is as follows: 5′-CATCTGGGAGAGACACTTACGGTTG-3′. Atg MO stock injecting solution was at 160–120 µM and Sp MO was at 50 µM (unless otherwise specified). 2.3 nl of the stock solution was injected per embryo at the 1–2-cell stage. For control injections, the standard MO, 5′-CCTCTTACCTCAGTTACAATTTATA-3′ (Co MO), was used. p53 MO injection stock solution was at 250 µM (Robu et al., 2007). All MOs were purchased from Gene Tools, LLC. To detect splice variants generated by Sp MO, the following PCR primer pair was used: Exon2F, 5′-AGAACCACAAGCGGAGGCTTT-3′; with Exon5/6R, 5′-GTCCTGCTCTTGTTGCTTTGCCAT-3′.

For rescue experiments with the Atg MO, a kozac sequence (5′-GCCACC-3′) was cloned immediately upstream of a mutagenized full-length rbms3 into pCS2⁺ at the Cla1–Xba1 sites. The endogenous first two codons (indicated by underlines) following the start site (shown in bold letters; 5′-ATGGGCAAA-3′) of rbms3 were changed to 5′-ATGGGAAAG-3′.

For live confocal imaging, Gt(pwp2h-citrine)^ct30a fish were soaked in BODIPY TR methyl ester vital dye (Invitrogen) for 30–60 min at 28°C to label all cells before mounting. Fish were anesthetized using 0.1% Tricaine, pH 7.2, in egg water and mounted laterally onto imaging molds (1% agarose in egg water).

All confocal imaging was performed at room temperature. Confocal z-stack images ranging from 40 to 100 µM were acquired using a microscope (LSM 510) and 25× LD LCI Plan-Apochromat NA 0.80 DIC 1 mm Korr or LD C-Apochromat 40×/1.1 W Korr UV-VIS-IR objective lenses (all from Carl Zeiss). LSM 510 version 4 software (Carl Zeiss) was used to acquire images.

Imaging of antibody-stained fish in whole-mount was performed as described for live confocal imaging earlier. Details of used fluorochromes are described further on. Imaging for whole-mount in situ hybridization and Alcian blue staining was performed using a microscope (Stemi SV 11; Carl Zeiss) with a Plan-Axiochromatic S 1.6× objective lens. Whole-mount images were acquired using AxioVision 4.6.3 software (Carl Zeiss).

All confocal images were processed using ImageJ 1.38×. Brightness/contrast processing of images was performed using Photoshop 7 (Adobe).

A combination of two probes for rbms3 were used: the first contained 5′ UTR and dRRM domains of rbms3 and was generated using 5′-GTGTGGAGCAACCGGCAG-3′ with 5′-TAATACGACTCACTATAGGGTCTCAGTGGACTCCATTCTGGCAA-3′ containing a T7 promoter (underlined) site. The second probe template spanned from the dRRM domains to the stop codon in pPCRII-TOPO (Invitrogen) and was generated using Not1 and with SP6 RNA polymerase. In situ hybridization was performed as described previously (Jayasena et al., 2008). In brief, embryos or larvae were treated with phenylthiourea (PTU) to prevent pigmentation and fixed in 4% PFA. Fish older than 24 hpf were Proteinase K treated (20–100 µg/ml depending on the stage) at room temperature. RNA probes were generated using SP6, T7, or T3 RNA polymerases, and hybridization was performed at 70°C. The citrine (Jayasena et al., 2011), dlx2a (Sperber et al., 2008), fgf3 (Walshe and Mason, 2003b), sox9a (Yan et al., 2005), sox10 (Dutton et al., 2001), foxd3 (Kelsh et al., 2000), pax2a (Pfeffer et al., 1998), barx1 (Sperber and Dawid, 2008), pax6 (Krauss et al., 1991), hox2a (Prince et al., 1998), runx2b (Flores et al., 2004), col2a1a (Yan et al., 2002), and smad2 (Müller et al., 1999) probes have been described previously. Processed fish were imaged (as described earlier) in either PBS or 70% glycerol.

Immunofluorescent staining on deyolked fish was essentially performed as described previously (Ohyama et al., 2006). In brief, fish were fixed in 4% PFA made in 0.1 M phosphate buffer. Blocking, primary, and secondary antibody incubations were performed in 10% goat serum containing 0.5% Triton X-100 in PBS at either room temperature (blocking step, 2–5 h) or 4°C (antibody incubations, 16–48 h). For confocal imaging, embryos were imaged in PBS. The following antibodies were used: rabbit anti–phospho-histone H3 (PH3; EMD Millipore), rabbit anti-activated Caspase3 (R&D Systems), rabbit anti-GFP (Molecular Probes), goat anti–rabbit Alexa Fluor 488, and goat anti–rabbit Alexa Fluor 594 (Molecular Probes).

To detect knockdown with Atg MO, Gt(rbms3-citrine)^ct30a embryos treated with either Atg MO or Co MO at 48 hpf were deyolked in cold Ringer’s solution with 0.5 mM DTT and 0.2 mM phenylmethanesulfonylfluoride (PMSF) supplemented with Complete protease inhibitors (EDTA free; Roche). For detecting Smad2 levels, 31-hpf head chunks (the region immediately posterior of the eyes and anterior of the fin buds) were collected, and pharyngeal arches from the same region (without any neural tissue) at 48 hpf were dissected in cold Ringer’s solution. Experiments were performed from at least three independent pools of protein extract. The following antibodies were used: mouse anti–α-tubulin (loading control; Sigma-Aldrich), rabbit anti-phosphorylated Smad2 (pSmad2; Cell Signaling Technology), rabbit anti-GFP (Invitrogen), rabbit anti-Smad2/3 (Cell Signaling Technology), secondary anti-mouse HRP (Promega), and anti–rabbit HRP (Promega). Comparison of band intensity between loading control and Smad2 was performed in ImageJ 1.38×. Loading control intensity was set to 1 and data were expressed as mean ± SD.

Cartilage staining was performed essentially as described previously (Robu et al., 2007) in 0.1% Alcian blue, 70% ethanol, and 0.37% hydrochloric acid. Larvae were bleached with 3% H₂O₂ plus 1% KOH in PBS for ∼20 min and treated with Trypsin (1 mg/ml; Sigma-Aldrich) for 30–60 min at 37°C. Experiments were performed at least three times to determine consistency. n values represent total number of larvae analyzed for the cartilage phenotype. Data were expressed as mean ± SD. To determine if there were statistical differences in phenotypes, between treatments, a χ² test was performed.

MO was injected into the Tg(sox10(−4.7):egfp) background and imaged confocally. Cells positive for both EGFP and a marker of interest within the pharyngeal arches were counted. Data were expressed as bifluorescent cells/arch. The results were expressed as mean ± SEM, and statistical analysis was performed using a two-tailed Student’s t test. For pharyngeal arch area measurements, 0.1% Tricaine-, pH 7.2, treated embryos were imaged in whole-mount at 24 hpf in egg water at room temperature, as described for confocal imaging (Jayasena et al., 2011). Images were acquired using a microscope (MVX10; Olympus) attached to a 1× objective lens (MV PLAPO; Olympus) and AxioVision version 4.6.3 software (Carl Zeiss). Images were processed using the ImageJ 1.38× measure tool.

Pharyngeal arches removed of contaminating neural tissues/whole embryos were collected. Total RNA was extracted using the RNAqueous-Micro kit (Ambion) and treated with DNase1 (Invitrogen), then cDNA was generated using random hexamers with Superscript III (Invitrogen).

QPCR was performed using SYBR green reagent (Applied Biosystems) on 5–7 independent RNA pools (generated from 2–3 experiments). For each QPCR primer set, amplification was performed in triplicate per pool of cDNA and validated for each MO treatment condition at least three times. The housekeeping gene ef1α (Plaster et al., 2006) was used as an internal control, and the results were analyzed using the comparative cycle threshold method for relative quantitation and normalized to ef1α levels.

For primer sequence details, see Table 1. All primer pairs showed 100% efficiency. Because of high variation between experiments for controls, the fold change was set to 1 to facilitate direct comparison between experiments (different pools of RNA from different injection experiments). All samples were run in triplicate per primer pair. Error bars represent SD from biological replicates (independent pools of RNA) unless otherwise stated. Statistical tests were performed using the two-tailed Student’s t test.

All DNA coding for desired mRNA were cloned into a pCS2⁺ vector and linearized with either Not1 or Kpn1 for generating capped mRNA using mMESSAGE mMACHINE SP6 kit (Ambion). For rescue of the MO phenotype, full-length mutagenized rbms3 (described earlier) was injected ∼20–30 pg/embryo (as already described).

Smad2ca (Dick et al., 2000) was injected ∼15–30 pg/embryo, and Atg MO was coinjected at 150 µM. Trace amounts of egfp mRNA (∼5 pg/embryo) were also coinjected as a selection marker.

Truncated rbms3 (dRRM-rbms3, lacking the putative RNA binding domains [RRM], amino acids 210–415) was cloned into Cla1–Xba1 site in pCS2⁺. d1egfp from the O. Pourquie laboratory (Institute of Genetics and Molecular and Cellular Biology, Strasbourg, France) containing the PEST domain of mouse ornithine decarboxylase (d1egfp) was subcloned into pCS2⁺ with EcoR1–Xba1 after the stop codon. The candidate 3′ UTR sequences were cloned into EcoR1–Xba1 site of pCS2⁺-d1egfp. pCS2⁺-cherryh2b was used as an injection control. Each embryo was injected with 70 pg of candidate 3′ UTR, 60 pg rbms3/dRRM-rbms3, and 20 pg cherryh2b. Levels of d1egfp were normalized to levels of cherryh2b. For QPCR, RNA was extracted from 4–6 individual embryos collected from at least two independent experiments per condition. Normalized fold change was graphed as mean ± SEM or SD, and a two-tailed Student’s t test was performed. For 3′ UTR and QPCR primer sequence details, see Tables 1 and 2.

RIP with full-length Rbms3-Myc and dRRM-Rbms3-Myc was performed as described previously (Keene et al., 2006), at 4°C unless otherwise stated. Approximately 50 pg of rbms3-Myc or dRRM-rbms3-Myc capped mRNA was injected at the one-cell stage. Embryos (16–24 hpf) were deyolked manually with a pulled glass Pasteur pipette in cold zebrafish Ringer’s solution (116 mM NaCl, 2.9 mM KCl, 5 mM Hepes, pH 7, 20 mM vanadyl ribonucleoside complex [VRC; Sigma-Aldrich], and 1 mM DTT). Per 10 ml of zebrafish Ringer’s solution, the following was added: 1 mini tablet EDTA-free protease inhibitors (Roche) and 10 µl of 0.2 M phenylmethanesulfonylfluoride (PMSF). To isolate total protein (∼0.5–1 mg), an Eppendorf pestle was used. About 100–150 µl of cold polysome lysis buffer (10 mM Hepes, pH 7.9, 100 mM KCl, 5 mM MgCl2, 1% IGEPAL CA-630, 1 mM DTT, 7× complete protease inhibitors [EDTA free; Roche], 100 U/ml SuperaseIn (Ambion), and 250 U/ml RNaseOUT [Invitrogen]) was used, and lysate was stored at −80°C. Approximately 5 µg of mouse anti–c-Myc antibody (clone 9E10; Santa Cruz Biotechnology, Inc.) or mouse IgG (Santa Cruz Biotechnology, Inc.) were incubated with 50 µl of Dynabeads Protein G (Invitrogen) overnight in NT2 (50 mM Tris, pH 7.4, 150 mM NaCl, and 1 mM MgCl₂) containing 5% BSA and 0.05% IGEPAL CA-630. The beads were washed several times in NT2 plus 0.05% IGEPAL CA-630. Thawed lysate was centrifuged at 20,000 g for 10 min. The resulting supernatant (0.1× final total volume) was incubated with the beads in NT2 supplemented with 200 U/ml RNaseOut, 1 mM DTT, 0.05% IGEPAL CA-630, and 20 mM EDTA for 2 h. All post-antibody incubation washes were performed in NT2 plus 0.5% IGEPAL CA-630; the final three washes were also supplemented with 1 M urea. 2–3 independent experiments were performed (mean ± SD). For calculations, QPCR results were initially normalized to input samples before calculating fold enrichment. Transcripts enriched greater than fivefold were considered to be significantly enriched. For primer sequence details, see Table 1.

Fig. S1 shows rbms3 expression at 16 and 18 hpf.

We wish to thank the following people: Dr. Le. A. Trinh and Bronner laboratory members for helpful discussions and technical help, Ho-yin Leung for performing 3′ RACE, and Leigh Ann Fletcher and David Mayorga from the Caltech Centers of Excellence in Genomic Science fish facility for fish maintenance.

This work was supported by the US Public Health Service Corps grants P50HG004071, DE017911, and HD037105 to M.E. Bronner.

Author contributions: C.S. Jayasena designed and performed the experiments, executed the data analysis, and wrote and edited the manuscript. M. Bronner performed data analysis and edited the manuscript.