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. 2015 Aug 18;4:e07048. doi: 10.7554/eLife.07048

A shift in anterior–posterior positional information underlies the fin-to-limb evolution

Koh Onimaru 1,2, Shigehiro Kuraku 3, Wataru Takagi 4, Susumu Hyodo 4, James Sharpe 2,5,*, Mikiko Tanaka 1,*
Editor: Marianne E Bronner6
PMCID: PMC4538735  PMID: 26283004

Abstract

The pectoral fins of ancestral fishes had multiple proximal elements connected to their pectoral girdles. During the fin-to-limb transition, anterior proximal elements were lost and only the most posterior one remained as the humerus. Thus, we hypothesised that an evolutionary alteration occurred in the anterior–posterior (AP) patterning system of limb buds. In this study, we examined the pectoral fin development of catshark (Scyliorhinus canicula) and revealed that the AP positional values in fin buds are shifted more posteriorly than mouse limb buds. Furthermore, examination of Gli3 function and regulation shows that catshark fins lack a specific AP patterning mechanism, which restricts its expression to an anterior domain in tetrapods. Finally, experimental perturbation of AP patterning in catshark fin buds results in an expansion of posterior values and loss of anterior skeletal elements. Together, these results suggest that a key genetic event of the fin-to-limb transformation was alteration of the AP patterning network.

DOI: http://dx.doi.org/10.7554/eLife.07048.001

Research organism: chicken, mouse

eLife digest

Humans, mice, and other animals with four limbs belong to a group of land-dwelling animals known as the tetrapods. This group of animals evolved from ancient fish and one crucial adaptation to life on land involved the modification of fins to form limbs. The front pair of limbs (the ‘arms’) evolved from the ‘pectoral’ fins of the ancient fish. These fins contain numerous bones that fan out from a set of bones called the pectoral girdle. However, most of the bones nearer the front side (the thumb side in the human limb) were lost in the ancestors of tetrapods as they moved onto land. Only the bone nearest the back remained as the ‘humerus’, which forms the upper part of the limb (i.e., the upper arm of humans).

In the embryos of mice and other animals, the limbs develop from structures called limb buds. For the limb to develop properly, the cells in the limb bud need to receive specific instructions that depend on their position in the bud. A protein called Gli3R provides cells with information about their position along the ‘anterior–posterior’ (or thumb-to-little finger) axis of the bud. This protein regulates several genes that are involved in limb development, and this results in different genes being expressed in cells along the anterior–posterior axis. For example, Alx4 is only expressed in a small area at the anterior end of the bud, while Hand2 expression is found in a large area towards the posterior part.

Gli3R is also found in a fish called the catshark, but it is not clear how it controls the formation of fins. Onimaru et al. show that the pattern of gene expression in the catshark fin bud is different to that of the mouse limb bud. For example, Alx4 is expressed in a larger area of the fin bud that extends further towards the posterior, while Hand2 is only found in a much smaller area at the posterior end of the bud. The experiments also suggest that Gli3R is active in a much larger area of the fin bud than in the limb bud.

Next, Onimaru et al. used a drug on the catshark embryos to increase the activity of another protein that can inhibit Gli3R. The fin buds of these shark had anterior shift in several gene expression domains, and the fins that formed were missing several anterior bones and had only a single bone connected to the pectoral girdle. Onimaru et al.'s findings suggest that during the evolution of the tetrapods, there may have been a shift in the anterior–posterior patterning of the fin bud to form a limb. An important area for future work will be to use genome-wide studies to study the fin/limb buds of other species.

DOI: http://dx.doi.org/10.7554/eLife.07048.002

Introduction

Regulatory interactions between transcriptional factors play important roles for interpreting a morphogen gradient as positional information (Balaskas et al., 2012). Changes in these regulatory interactions may therefore be key players for patterning changes during morphological evolution. The fin-to-limb transformation is a prominent but still unsolved example of morphological evolution. 150 years ago Carl Gegenbaur subdivided the skeletal elements of shark pectoral fins into three segments along the anterior–posterior (AP) axis: propterygium, mesopterygium, and metapterygium (Gegenbaur, 1865) (Figure 1A), which are also found in the majority of chondrichthyans, none-teleost actinopterygians, placoderms, and acanthodians (Orvig, 1962; Coates, 1994, 2003). Therefore, possession of propterygium, mesopterygium, and metapterygium is considered to be a plesiomorphic state for gnathostomes. In the sarcopterygians (lobe-finned fishes including tetrapods), the propterygium and mesopterygium have been lost (Coates, 2003), thus, suggesting that anterior positional values have been lost or reduced during tetrapod evolution.

Figure 1. Anterior–posterior patterning in Scyliorhinus canicula pectoral fin buds.

(A) Skeletal patterns of S. canicula pectoral fin and mouse limb. Blue colours, homologous elements. (B–G) In situ hybridisation for Alx4 (B), Pax9 (C), Hand1 (D), Zic3 (E), Hand2 (F), and Tbx2 (G) in S. canicula pectoral fin buds at stage 30 and mouse limb bud at E11.5 (left panel in G). Arrowheads in G, anterior boundary of posterior Tbx2 expression. Dorsal view; anterior is to the top. Scale bars, 100 μm. (H) Schematic of the gene expression patterns. Arrowheads, Hand2 expression boundary. Expressions of mouse limb buds at E11.5 are after EMBRYS database (Yokoyama et al., 2009; Fernandez-Teran et al., 2003).

DOI: http://dx.doi.org/10.7554/eLife.07048.003

Figure 1.

Figure 1—figure supplement 1. Molecular phylogenetic trees of relevant S. canicula genes.

Figure 1—figure supplement 1.

(A–E), Trees for Alx4 (A), Pax9 (B), Hand1 (C), Zic3 (D), Tbx2 (E), and Ptch1 (F) were generated from amino acid sequences of the homeodomain and neighbouring sequences (A), the paired box and C-terminal sequences (B), Helix-loop-helix domain and C-terminal sequences (C), N-terminal sequences (D), and C-terminal sequences (E and F). The neighbour-joining method was used for constructing the trees. The numbers at nodes indicate bootstrap probabilities with 1000 replicates.
DOI: http://dx.doi.org/10.7554/eLife.07048.004

Figure 1—figure supplement 2. Temporal expression analysis of Alx4, Pax9, Hand2, and Hoxa13 in S. canicula pectoral fins.

Figure 1—figure supplement 2.

(A–D) Expression of Alx4 (A), Pax9 (B), Hand2 (C), and Hoxa13 (D) at the indicated stages. Dorsal view of right pectoral fin buds of S. canicula embryos (anterior to the top). (A) Alx4 was initially expressed throughout the fin buds at stage 25, but the posterior expression of Alx4 was reduced by stage 27. Alx4 expression was then restricted to the anterior two-thirds of the fin bud at stage 29 and to the anterior half of the fin bud at stage 31. (B) Pax9 expression was not detected at stage 27 but was present in anterior distal fin buds at stage 29. This expression persisted until at least stage 31. (C) Hand2 was initially expressed throughout the fin bud, although slightly more-robust expression was observed at the posterior side. Subsequently, Hand2 expression became posteriorly restricted by stage 27. (D) Hoxa13, which is a marker for a late developmental stage in tetrapod limb buds, was detected since stage 29 in a distal domain. (E) OPT scans of Sox9 expressions (red) stained with propidium iodide (green) at stage 28 (left) and late stage 29 (right). pg, pectoral girdle; ms, mesopterygium; mt, metapterygium. (F) Comparison of Alx4 (black), Pax9 (grey), Hand2 (blue), and Hoxa13 (light green) expression patterns between S. canicula fin buds and chick forelimb and hindlimb buds (see Figure 1—figure supplement 3 for detailed gene expression in chick limb buds). st, stage. Scale bars, 100 μm.
DOI: http://dx.doi.org/10.7554/eLife.07048.005

Figure 1—figure supplement 3. Temporal expression analysis of Alx4, Pax9, Hand2, and Hoxa13 in chick limb buds.

Figure 1—figure supplement 3.

(A–H) Expression of Alx4 (A, B), Pax9 (C, D), Hand2 (E, F), and Hoxa13 (G, H) at the indicated stages. Dorsal views of chick forelimb (A, C, E, G) and hindlimb (B, D, F, H) buds (anterior to the top; distal to the right). (A, B) At stage 19, Alx4 expression was broad but was slightly weaker in the posterior side of forelimb (A) and hindlimb (B) buds. At later stages, its expression became restricted to the anterior one-third of the limb buds, with subsequent further restriction to the anterior proximal region both in forelimb (A) and hindlimb (B) buds. Alx4 expression also appeared in the posterior distinct region of chick forelimb (A) and hindlimb (B) buds at stage 25. (C, D) Pax9 expression was first present in the anterior region at stage 24 in forelimb buds (C) and stage 23 in hindlimb buds (D) but was more distally restricted than Alx4 expression. At stage 25, expression of Pax9 was robust, but the extent of its overlap with Alx4 expression was limited both in forelimb (C) and hindlimb (D) buds. (E, F) Hand2 expression was initially complementary with Alx4 and subsequently with Pax9 expression in forelimb (E) and hindlimb (F) buds. (G, H) At stage 21, Hoxa13 expression fully overlapped with the distal Hand2 expression domain in forelimb (G) and hindlimb (H) buds. At later stages, co-expression of Pax9 and Hoxa13 was observed in the anterior distal portion of the limb buds, but this overlap was not as extensive as that between the Hand2 and Hoxa13 expression domains. st, stage. Scale bars, 100 μm.
DOI: http://dx.doi.org/10.7554/eLife.07048.006
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In mouse limb buds, Hand2, Gli3, and Shh are key genes for controlling AP patterning (Riddle et al., 1993; Te Welscher et al., 2002). One of the earliest patterning events is the mutual transcriptional repression between Gli3 in the anterior tissue and Hand2 in the posterior (Te Welscher et al., 2002; Osterwalder et al., 2014). This early polarity in expression contributes to the subsequent posterior localized expression of Shh, and this morphogen in turn reinforces the anteriorly restricted Gli3 protein activity (Shh inhibits the default processing of the Gli3 protein to its repressor form (Gli3R), thus, creating a gradient of Gli3R along the AP axis) (Wang et al., 2000). Several studies on fin development of actinopterygians and chondrichthyans have revealed that posterior Shh expression is conserved among gnathostomes (Dahn et al., 2007; Davis et al., 2007; Yonei-Tamura et al., 2008; Sakamoto et al., 2009a). However, in fish fin development, the detailed roles of Shh signalling for AP patterning are not well studied and the role of the Hand2-Gli3 mutual interaction remains to be elucidated.

Results and discussion

To investigate changes in AP patterning during the fin-to-limb transition, we first cloned a number of AP patterning genes from the non-model species Scyliorhinus canicula (Figure 1B–H and Figure 1—figure supplement 1 for phylogentic analyses). In the mouse limb bud, Alx4, Pax9, Hand1, and Zic3 are positively regulated by Gli3R (Te Welscher et al., 2002; Fernandez-Teran et al., 2003; McGlinn et al., 2005; Vokes et al., 2008) and thus are expressed in a localized anterior domain (one-third of the axis), while Hand2 and Tbx2 show broad posterior expression domains (two-thirds and one-half of the axis, respectively). In stage 30 S. canicula embryos (staged according to Ballard et al., 1993), we found instead that the anterior genes Alx4, Pax9, Hand1, and Zic3 were expressed in broad domains, which extend more posteriorly than in the mouse (half the fin bud for Alx4, two-thirds for Pax9 and Hand1, and the whole axis for Zic3, Figure 1B–E). By contrast, the Hand2 and Tbx2 domains were more posteriorly restricted in S. canicula fin buds than in mouse limb buds (Figure 1F,G). All of these 6 AP patterning genes show the same trend—their expression boundaries are more posterior in S. canicula fin buds (Figure 1H), apparently reflecting a gross shift in the AP coordinate system. We chose 3 of these genes to test at multiple time-points to determine whether this was a transient gene expression state (Figure 1—figure supplements 2, 3), but in all cases these shifts were observed from stage 29 to stage 31 (which covers ∼30 days of S. canicula development). In particular, stage 29 is a stage where Sox9 expression (a prechondrogenic marker) starts in the proximal part of the pectoral fin buds (Figure 1—figure supplement 2E), which suggest that the observed shift of AP values would affect proximal skeletal elements as well as distal.

Since the above AP patterning genes are regulated by Shh–Gli3 pathway (Te Welscher et al., 2002; Fernandez-Teran et al., 2003; McGlinn et al., 2005; Galli et al., 2010), we cloned Gli3 from S. canicula fin buds (Figure 2A and Figure 2—figure supplement 1A for phylogenetic tree) and analysed its expression in pectoral fin buds. In striking contrast to tetrapod limb buds (Büscher et al., 1997; Schweitzer et al., 2000), Gli3 expression is not restricted to the anterior region—thus again indicating a general posterior shift of AP positional values in the S. canicula. To address whether this situation is conserved in other chondrichthyans, we also cloned and analysed the expression of Gli3 in pectoral fin buds of a holocephalian, Callorhinchus milii, which has propterygium (Figure 2B), and again found expression in the posterior part of pectoral fin bud at stage 31 (staged according to Didier et al., 1998; Figure 2B). Since Hand2 is expressed posteriorly and thus now overlaps with Gli3, this strongly suggests that the Hand2–Gli3 mutual inhibition seen in tetrapods is weak or non-existent in chondrichthyans.

Figure 2. Expression and processing of Gli3 and Gli2 in S. canicula embryos.

(A) Expression of Gli3 in S. canicula pectoral fins. (B) Alcian blue staining of C. milii pectoral fin at stage 35 (top, the ventral view of a right fin flipped horizontally) and Gli3 expression at stage 31 (bottom, a left pectoral fin flipped horizontally). pro, propterygium. (C) Expression of Gli2 in S. canicula pectoral fin buds. Scale bars, 100 μm. (D) The Gli3 chimera constructs. hGli3 PDD, full-length human Gli3 (grey box) with Myc tags. hGli2, ScGli2 and ScGli3 PDD, chimeric Gli3 genes recombined at the processing determinant domain (PDD) with human Gli2, S. canicula Gli2 and Gli3, respectively. (E) Protein processing of the chimeric constructs in cell cultures treated with either FSK (+) or DMSO (−). Truncated Gli3 is detected only in hGli3 PDD (lane 4) and ScGli3 PDD (lane 8).

DOI: http://dx.doi.org/10.7554/eLife.07048.007

Figure 2.

Figure 2—figure supplement 1. Phylogenetic tree of Gli2 and Gli3, and PDD amino acid sequences.

Figure 2—figure supplement 1.

(A) Trees for Gli2 and Gli3 were generated from amino acid sequences of the zinc finger domain and PDD. The neighbour-joining method was used for constructing the trees. The numbers at nodes indicate bootstrap probabilities with 1000 replicates. (B) Human Gli3 (the upper diagram) is composed of an N-terminal repressor domain (Repressor), a DNA-binding domain (Zinc finger), and a C-terminal activator domain (Activator). The alignment shows the homologous PDD domains from both human and S. canicula Gli2 and Gli3.
DOI: http://dx.doi.org/10.7554/eLife.07048.008
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In chick and mouse limb buds, Gli2 does not play a major role in AP patterning because of its weak processing efficiency to produce its repressor form (Wang et al., 2000). However, in zebrafish, Gli2 does indeed act as a repressor (Maurya et al., 2013), so we checked whether Gli2 could be playing the repressor role in S. canicula fin buds. First, we analysed Gli2 expression in S. canicula embryos and found it to be uniform until stage 29 (Figure 2C) and then subsequently restricted to the posterior region (Figure 2C). Second, we checked whether Gli3 and Gli2 of S. canicula have the repressor function, by measuring their processing efficiencies. We analysed the processing determinant domain (PDD), which determines the differential processing efficiencies of Gli3 and Gli2 in mice and humans (Pan and Wang, 2007). We inserted the PDDs from human Gli2 and S. canicula Gli2 or Gli3 into the human Gli3 PDD region (Figure 2D and Figure 2—figure supplement 1 for the amino acid sequences), transfected these constructs into HEK293 cells, and treated the cells with forskolin (FSK) to induce Gli processing. Human and S. canicula Gli2 PDD did not induce Gli3R, whereas their Gli3 PDDs did (Figure 2E). Thus, in S. canicula (as in chick and mouse), Gli3, but not Gli2, plays the major role in repressor production.

We next wished to explore if a genetic explanation could be found for the lack of Gli3 repression in the posterior part of pectoral fin buds of S. canicula and C. milii. To compare Gli3 enhancers in chondrichthyans and tetrapods, we used the VISTA enhancer browser (Visel et al., 2007) and found a limb-specific Gli3 enhancer, element 1586, which replicates anterior Gli3 expression in mouse limb buds. We identified the homologues of element 1586 in S. canicula and C. milii and compared them with those from other vertebrates. Consistent with the slow evolutionary rate of chondrichthyans and coelacanth (Amemiya et al., 2013; Renz et al., 2013; Venkatesh et al., 2014), element 1586 is conserved in tetrapods, coelacanth, and chondrichthyans, but not in gar, medaka, and zebrafish (Figure 3A). To assess whether the element 1586 in different species has different functionalities, we cloned this element from chick, S. canicula, and C. milii in front of a basal promoter followed by a GFP reporter (Ochi et al., 2012; Figure 3B). These constructs were electroporated into chick forelimb buds with a constitutively active RFP vector (to determine the spatial efficiency of electroporation). As with endogenous Gli3 expression (Büscher et al., 1997), the chick element 1586 drove GFP expression specifically in anterior tissue and was repressed in the posterior region, even though RFP was expressed throughout the buds (Figure 3C). The element 1586 from both S. canicula and C. milii also drove GFP expression in the chick limb buds, confirming that its general activity is conserved from sharks to tetrapods. However, in both cases, the specific posterior repression observed in the chick element was absent (Figure 3D,E). Thus, the differential activity of this enhancer (with tetrapods showing posterior repression, and chondrichthyans not) recapitulates the differences in Gli3 expression within these groups. Furthermore, by recombining S. canicula and chick enhancers, we identified a sequence that can exert the posterior repression when inserted into the S. canicula enhancer (Figure 3F,G and Figure 3—figure supplement 1). This sequence contains tetrapod or sarcopterygian-specific sequences, suggesting that the posterior repressive activity would have been acquired in a stepwise fashion.

Figure 3. The Gli3 limb-specific enhancer of S. canicula and C. milii.

(A) VISTA plots of Gli3 intron 3 from indicated animals. Blue vertical bars, exons of human Gli3; black rectangle, element 1586. Regions with >70% identity are indicated: blue, exon; pink, non-coding sequences. (B) The enhancer construct. (C) GFP expression in chick forelimb buds driven by chick element 1586 at stage 19 (top, n = 3/3), stage 23 (middle, n = 14/14), and empty vector (bottom, n = 0/7). pCAGGS-RFP (right). (D–F) GFP expression driven by element 1586 of S. canicula (D, n = 11/11), C. milii (E, n = 10/10) and Sc1586mt (F, n = 4/4). Scale bars, 100 μm. (G) Scheme of Sc1586mt, S. canicula enhancer (blue) partially replaced by chick sequence (green) and alignment. Boxes indicate tetrapod (i) and sarcopterygian (ii) specific sequences.

DOI: http://dx.doi.org/10.7554/eLife.07048.009

Figure 3.

Figure 3—figure supplement 1. Detailed functional analyses of element 1586.

Figure 3—figure supplement 1.

(A) Alignment of element 1586 sequences. Points where chick and S. cacnicula enhancers are recombined in (B) are indicated by arrows a and b. Box i and Box ii are sequences that are replaced with S. canicula sequence in (C), activities of element 1586 recombined at the indicated points. Note that the posterior repressive activity is not altered at arrow a, but partially destroyed at arrow b, suggesting that the repressive sequences are located between arrow a and b and after arrow b. (C) Activities of chick element 1586 that are partially recombined with S. canicula sequences at Box i and Box ii in (A). The sequences were chosen because of specific conservation among tetrapods (Box i) and sarcopterygians (tetrapod and coelacanth, Box ii). Each of replacement can slightly alter the posterior repression of GFP (n = 3/3). When the chick enhancer is replaced by both sequence (the most right panel), the distal GFP expression is significantly shifted to the posterior limb buds (n = 4/4, white arrow). And weak expressions are also detected in the posterior proximal limb buds (n = 2/4. arrow head in the bottom panel, magnified view of the above white box with enhanced contrast), suggesting that the sequences at Box i and Box ii are partially responsible for the posterior repression. Scale bars, 100 μm.
DOI: http://dx.doi.org/10.7554/eLife.07048.010
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Finally, we wished to address whether changes to AP positional information could modify skeletal arrangement of the propterygium and mesopterygium in catshark. For this purpose, we explored methodologies for performing manipulative experiments on this very slow-developing non-model fish (see ‘Materials and methods’). We treated S. canicula embryos with retinoic acid (RA) to increase Shh-signalling activity (at stage 29 with 1 μg/ml of RA for 4 days). Activation of Shh signalling by RA is known to be conserved among vertebrate limbs/fins (Riddle et al., 1993; Hoffman et al., 2002; Dahn et al., 2007), and as expected, the most reliable Shh target gene, Ptch1 expression (Marigo et al., 1996; Vokes et al., 2008; and see Figure 1—figure supplement 1F for phylogenetic analysis) was increased and expanded anteriorly (Figure 4A). Consistent with this, Hand2 expression also extended anteriorly (Figure 4B)—probably due to inhibition of Gli3 repressor formation by ectopic activation of Shh signalling (as revealed by the extended Ptch1 expression). On the other hand, Pax9 expression (an anterior marker) was significantly downregulated and showed only weak expression in the anterior part of the fin buds (Figure 4C). The most anterior regions may not be sensitive to this treatment, as expression of Alx4 was not significantly shifted (Figure 4D), and this may be due to the lack of inhibitory regulation from Hand2 to Gli3 described above. To test whether the results of RA treatment were due to specific effects on AP patterning or instead due to a more general interference with limb development, we examined a marker for proximal-distal (PD) patterning in mouse and chick limb buds—Hoxa13 (Tamura et al., 1997; Mercader et al., 2000; Yashiro et al., 2004). In RA-treated pectoral fin buds, Hoxa13 expression was weaker than in control, but a shift in its expression domain was not seen (Figure 4E), showing that the impact of RA in these experiments is primarily on the AP patterning (the shifts of Ptch1, Hand2, and Pax9, Figure 4A–C), rather than on PD patterning or a general impact on development. Most intriguingly, we examined skeletal patterns of S. canicula pectoral fins in these partially ‘posteriorised’ fin buds (Figure 4F). Phenotypes varied from mild to severe but in all cases the appearance of distinct anterior elements (propterygium and mesopterygium) was lost. In the mild cases, a proximal element anterior to the metapterygium is attached to the pectoral girdle (single asterisk in Figure 4F). This proximal element may result from a fusion of the proximal parts of propterygium and mesopterygium. Whereas, the severe cases still have a fused element anterior to the metapterygial axis (double asterisk in Figure 4F), but this element is not directly attached to the pectoral girdle, indicating that the pectoral fin of the severe phenotype has lost the anterior proximal elements. By contrast, the posterior metapterygium itself was larger than normal, but retained its strong identity as the primary axis from which radial branching was observed. Although RA treatment potentially cause non-specific effects, given the clear affect on AP patterning (the shifts of Ptch1, Hand2, and Pax9, Figure 4A–C), while causing no obvious effects on PD patterning, the main cause of the phenotype is likely caused by the AP pattern change.

Figure 4. RA treatment causes ectopic activation of Shh signalling and loss of anterior skeletal elements.

Figure 4.

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(A–E) In situ hybridisation of S. canicula pectoral fin buds for Ptch1 (A; left fins flipped horizontally), Hand2 (B), Pax9 (C), Alx4 (D), and Hoxa13 (E) treated with 1% DMSO or 1 μg/ml retinoic acid (RA) (n = 2/2 for each for each except n = 4/4 for Hand2). Arrowheads in C, a weak expression of Pax9. White brackets in E, width of Hoxa13 expression domain along the proximal-distal (PD) axis. (F) Pectoral fin skeletal patterns of 1% DMSO control (n = 4) and 1–2 μg/ml RA (n = 4). Right panels, schematics of interpretive skeletal patterns. *, an anterior proximal radial; **, a fused radial attached to the metapterygium; pg, pectoral girdle; ppr, pms and pmt, proximal propterygium, mesopterygium and metapterygium. Scale bars, 100 μm. (G) Comparison between S. canicula fin and mouse limb. Green and blue colours represent anterior–posterior (AP) positional information. ppr, pms, and pmt denote proximal propterygium, mesopterygium, and metapterygium, respectively.

DOI: http://dx.doi.org/10.7554/eLife.07048.011

In the present study, we have found that S. canicula pectoral fin buds have a gross posterior shift in the AP coordinate system compared to mouse limb buds. We show that S. canicula and C. milii lack a specific enhancer activity for Gli3, which in tetrapods mediates the posterior repression, and that this genetic difference likely contributes to the shift of AP positional information. Finally, RA treatment analyses suggest that a partial posteriorisation of S. canicula fin buds leads to a loss of anterior proximal elements (propterygium and mesopterygium). Thus, while the loss of the anterior proximal elements during evolution was associated with cis-regulatory changes of Gli3 in the RA experiments, it was driven by a Shh-mediated affect on the Gli3 protein itself, but in both cases achieving similar phenotypic changes by anterior shift in AP pattern. In support of our observations, a recent study also showed that anterior extension of Shh signalling accompanied with an anterior shift of Gli3 expression resulted in a loss of anterior skeletal elements in mouse limbs (Li et al., 2014). Considering all these data together, we therefore propose that one of the key events during the fin-to-limb transition was an anterior shift of AP positional information (a posteriorisation), which caused the loss of anterior proximal elements (Figure 4G).

In the RA treatment experiments, we also observed that the anterior distal radials reduced and only metapterygial radials retained, suggesting that anterior shift in AP positional information may also have had an impact on the distal radials during the fin-to-limb transformation. Interestingly, nearly 30 years ago, a classic study proposed that the distal end of the metapterygial axis (which has a uniformly posterior position in chondrychthians) bent anteriorly during acquisition of digits—the so-called digital arch model (Shubin and Alberch, 1986; Oster et al., 1988). Although the detailed validity of this model is unclear (Wagner and Larsson, 2007), there is a possibility that an AP shift in molecular patterning was involved in the acquisition of digits. In addition to our RA treatment analysis, knockdown analyses of actinotrichia proteins, which are components of fin rays and lost in tetrapod, show an anterior shift in several gene expressions in zebrafish pectoral fin buds (Zhang et al., 2010). Therefore, it is interesting to speculate that AP positional information may have shifted several times until the acquisition of digits.

We have shown that the Gli3 regulatory region of S. canicula and C. milii lacks the tetrapod-specific repressive element, which is likely needed for the Gli3–Hand2 interaction in mouse limb buds. In mice, Gli3−/−; Hand2−/c limbs show a severe dysplastic humerus (some of them have ectopic protrusion in humerus; Osterwalder et al., 2014), suggesting that the Gli3–Hand2 interaction has an important role for patterning the proximal elements. However, how Gli3 regulates the proximal skeletal pattern is not well understood even in mice. Although Gli3 is involved in the stylopod (humerus/femur) formation in mice, the phenotype in stylopod always appears with combination of other gene knockouts. For example, Gli3−/−; Plzf−/− mice lack a femur, and Gli3−/−; Alx4−/− mice exhibit humerus malformation (Barna et al., 2005; Panman et al., 2005). These facts suggest that evolutionary modification of Gli3 regulation is likely necessary, but additional regulatory modifications are required for the loss of the anterior elements. Since Alx4 and Hand2 are expressed in S. canicula pectoral fin bud, and Plzf is involved only in hindlimb development, currently there is no obvious candidate that would be involved in the loss of propterygium and mesopterygium. Although S. cacnicula genome has not been sequenced, systematic studies at genome-wide level such as ChIP-seq in S. cacnicula pectoral fin buds would be invaluable to provide a more complete picture of evolutionary mechanism of the loss of the anterior elements in the future.

In conclusion, by taking advantage of the slow evolutionary rates of chondrichthyian genomes, we were able to precisely compare the gene expression, function and regulation between pectoral fin and limb development, and discover a key difference between them. In particular, our study suggest that changes in morphogen interpretation by gene regulatory network mutations may have a major impact on morphological evolution.

Materials and methods

Animals

Experiments were performed in accordance with guidelines for animal experiments of Tokyo Tech and CRG, and experiments involved in mice were approved by animal ethics committees of CRG (JMC-07-1001P3-JS). Catshark (S. canicula) eggs were incubated at 12–16°C in seawater and staged according to (Ballard et al., 1993). C. milii eggs and embryos were collected as described (Takagi et al., 2012) and staged according to (Didier et al., 1998). C52BL/6 (Charles River Laboratories, Wilmington, MA) mouse timed-pregnant females were sacrificed at different days after gestation E11.5. Chicken (Gallus gallus) eggs were incubated at 38°C in a humidified incubator until the desired Hamburger–Hamilton (HH) stage (Hamburger and Hamilton, 1951) was reached. For in situ hybridisation, embryos were fixed overnight in 4% paraformaldehyde in phosphate-buffered saline, dehydrated in a graded methanol series, and stored in 100% methanol at −20°C.

Gene isolation and phylogenetic analysis

Total RNA was extracted from stage 24 to 29 S. canicula embryos, stage 28 chick embryos and E11.5 mouse embryos using an RNeasy kit (Qiagen, Netherlands). cDNA was synthesised by reverse transcription and used as a template for PCR. Extraction of total RNA and cDNA synthesis from C. milii embryos were carried out as described (Takagi et al., 2012). To clone S. canicula and C. milii genes, we used primers that were based on the nucleotide sequences of putative C. milii orthologues found in the Elephant Shark Genome Project database (http://esharkgenome.imcb.a-star.edu.sg/) (Venkatesh et al., 2007) for Pax9, Alx4, and Gli3; SkateBase (http://skatebase.org/) (Wang et al., 2012) for Hand1, Zic3, Tbx2, and Ptch1; and GenBank for Gli2 (EU196410) and Sox9 (EU241880): S. canicula Alx4, 5′-AGGAATGAACGGCGAGACTTG-3′ and 5′-TCATGTTGCCCAAGATATAGC-3′; S. canicula Pax9, 5′-GCTGTGTCAGCAAGATACTGG-3′ and 5′-CCGCACTGTATGTCATGTAGG-3′; S. canicula Gli3, 5′-CAGCCCAGCAGAATACTACC-3′ and 5′-GAGATCTCAGCGCCATTGATG-3′; S. canicula Gli2, 5′-GTAAAGCTTACTCACGACTCG-3′ and 5′-CGTAAGAGTCAGCCGAGCTGATG-3′; S. canicula Sox9, 5′-CCCAGGTGCTGAAGGGATAC-3′ and 5′-GGCAGGTACTGGTCGAACTC-3′; S. canicula Hand1, 5′-GAGAGCATCAACAGCGCATTCGC-3′ and 5′-TTCCTGGTCCTCAACCTGGTCAG-3′; S. canicula Zic3, 5′-GTGGCCATGGCGATGTTACTGGATGGTG-3′ and 5′-GTTTCTCGCCGGTGTGCACTCGGATGTG-3′; S. canicula Tbx2, 5′-GACACAGAAACCAGCTTCAGTCACAGTC-3′ and 5′-GAAAGTCGCGATACCCAATGTGGATCAG-3′; S. canicula Ptch1, 5′-GAGGTTTCACCTCTCGATGGGAGAACC-3′ and 5′-CCATACTAATGTGTTCTGTTCCCACTG-3′; C. milii Gli3, 5′-GAGATCTCAGCGCCATTGATG-3′ and 5′-GAGATCTCAGCGCCATTGATG-3′. To clone chick and mouse genes, we used primers that were based on the nucleotide sequences of Pax9 (NM_204912) (Muller et al., 1996), Hoxa13 (NM_204139), and Tbx2 (NM_009324) (Bollag et al., 1994a): chick Pax9, 5′-TGAGCGACACCTCGTCGTACC-3′ and 5′-GGTTATGCGATCCACTGCTA-3′; chick Hoxa13, 5′-GTCATGTTCCTCTACGACAAC-3′ and 5′-GGTGGACTTCCAGAGGTGAGG-3′; mouse Tbx2, 5′-ATCCTGAACTCCATGCACAAGTACC-3′ and 5′-GAACTGCTGCCCATGCAGGTGGCTG-3′. The gene fragments were cloned into pBluescript SK− or pCR4 (Invitrogen, Thermo Fisher Scientific Inc., Waltham, MA). The partial coding sequences for Alx4 (1112 bp), Pax9 (729 bp), Gli3 (1937 bp), Hand1 (465 bp), Tbx2 (926 bp), Zic3 (919 bp) and Ptch1 (791 bp) of S. canicula and Gli3 (428 bp) of C. milii have been submitted to GenBank under accession numbers KC507187–9, KF748129, and KP055651-KP055653, KF297620, respectively. Phylogenetic analysis was used to confirm the orthology of newly identified S. canicula and C. milii genes. Amino acid sequences were aligned using ClustalX (Thompson et al., 1997). Regions that could not be aligned were excluded from the analysis. Neighbour-joining phylogenetic trees of amino acid sequence data sets were constructed with MEGA5 (Tamura et al., 2011). Bootstrapping was carried out with 1000 replicates.

Probe synthesis and in situ hybridisation

Chick Alx4 (NM_204162) was kindly provided by Dr Toshihiko Ogura. Riboprobes for Hand2 (AY057890) and Hoxa13 (EU005550) of S. canicula and for chick Alx4 were synthesised as described (Takahashi et al., 1998; Tanaka et al., 2002a; Sakamoto et al., 2009a). The cloned genes described above were used as templates for riboprobe synthesis. Whole-mount in situ hybridisation was carried out as described (Tanaka et al., 2002a). Sox9 expressions were scanned with Optical Projection Tomography (OPT) as described (Sharpe et al., 2002) and analysed with Volviewer (Lee et al., 2006).

Gli processing analysis

Human Gli3 (clone name: pFN21AE1055) and Gli2 (Roessler et al., 2005) were obtained from the Kazusa DNA Research Institute (Nagase et al., 2008) and Addgene, respectively. pCAGGS was kindly provided by Dr Toshihiko Ogura and originated from Dr Jun-ichi Miyazaki (Niwa et al., 1991). For Western blotting analysis, the N-terminal HaloTag in the human Gli3 construct was replaced with a 6×Myc tag (Myc-hsGli3). Then, the human Gli3 PDD (amino acids 644–842) (Pan and Wang, 2007) was replaced with the homologous domain from human Gli2 and S. canicula Gli2 and Gli3 by a combination of PCR (Wurch et al., 1998) and restriction enzyme digestions. The HEK293 cell line was kindly provided by Dr Masayuki Komada. HEK293 cells were grown in Dulbecco's modified Eagle medium (Sigma–Aldrich, St. Louis, MO) supplemented with 10% foetal bovine serum (Gibco, Thermo Fisher Scientific Inc., Waltham, MA) and penicillin/streptomycin (Sigma–Aldrich) at 37°C. For Western blot analysis, cells were plated in 6-well plates without penicillin/streptomycin and transfected with 4 μg of constructs using polyethylenimine (GE Healthcare, England) for 3 hr. After the transfection, the medium was changed, and cells were cultured for 24 hr and then treated with 50 μM forskolin (FSK; Sigma) in Dimethyl sulfoxide (DMSO) DMSO or with DMSO alone for 24 hr. Whole-cell extracts were prepared by solubilisation in lysis buffer containing 50 mM Tris-HCl, pH 7.5; 150 mM NaCl; 1 mM ethylenediaminetetraacetic acid (EDTA); 1% Triton X-100; 0.1% Sodium Dodecyl Sulfate (SDS)S; 1% sodium deoxycholate; and protease inhibitor cocktail (Roche, Switzerland). Whole-cell lysates were separated by sodium dodecyl sulphate–polyacrylamide gel electrophoresis and analysed by Western blotting and anti-c-Myc (Sigma–Aldrich), anti-rabbit IgG secondary antibody conjugated with horseradish peroxidase (Jackson ImmunoResearch, West Grove, PA), and enhanced chemiluminescence detection (GE Healthcare).

Enhancer analysis

The limb-specific Gli3 enhancer was found with VISTA enhancer browser (http://enhancer.lbl.gov/) (Visel et al., 2007). The enhancer ID is hs1586, which is located in Gli3 intron 3 in the human genome (hg19). For alignment, Gli3 intron 3 sequences from mouse (Mus musculus), chick (G. gallus), frog (Xenopus tropicalis), coelacanth (Latimeria chalumnae), gar (Lepisosteus oculatus), medaka (Oryzias latipes), and zebrafish (Danio rerio) from the Ensembl and Pre Ensembl genome browsers (http://www.ensembl.org/, http://pre.ensembl.org/) were collected. The element 1586 homologue from elephant shark (C. milii) was retrieved from the genome assembly (http://esharkgenome.imcb.a-star.edu.sg/) (Venkatesh et al., 2007) by using human element 1586 sequence as the query. The GenBank accession number of the C. milii element 1586 is AAVX01295166. The S. canicula counterpart of element 1586 was amplified by PCR with primers designed from conserved sequences of the upstream exon and the distal part of element 1586: 5′-AGTGGACCCCCGAAATGGCTACATGGACC-3′ and 5′-GAACATCTTCTAATTTACTGGAATCCCAG-3. The amplified fragment was then cloned into pBluescript SK−. The sequence of S. canicula element 1586 was deposited in GenBank under accession number KF297619. The alignment was carried out with the SLAGAN method, and overall sequence similarities in the alignment were visualised with mVISTA (Mayor et al., 2000; Brudno et al., 2003; Frazer et al., 2004).

For functional analysis, the element 1586 homologues were isolated from chick and C. milii genomes by PCR. The following forward and reverse primers were used: chick element 1586, 5′-CGAGCTCCCTCCTCAGTCATTCAGTTCTGC-3′ and 5′-TGTGTGAGACATACTTTGATC-3′; C. milii element 1586, 5′-GAGCTCGTACAGTGATGACTGAAATGGTG-3′ and 5′-GAGATTTCGAGTCTCTTTGATC-3′. The amplified DNA fragments were cloned into pBluescript SK−. To subclone the S. canicula 1586 fragment, we used the following primers: 5′-CCGCTCTAGAACTAGCATCAATATGATTTGCTGAG-3′ and 5′-CGGGGGATCCACTAG GCTTCACGAGCATCAGGAAC-3′. The element 1586 sequence from each species was subcloned in front of a chicken β-actin basal promoter that is followed by a GFP reporter (Ochi et al., 2012). Recombined enhancers were created by PCR. In ovo, electroporation was carried out as described (Suzuki and Ogura, 2008). A DNA solution was prepared with Maxi Prep (Qiagen). pCAGGS-RFP was kindly provided by Dr Cheryll Tickle. Gli3 limb enhancers and empty β-actin basal promoter–GFP at ∼6 μg/μl, coloured with ∼3% fast green, and co-electroporated with pCAGGS-RFP (∼2 μg/μl) into the presumptive forelimb field of stage 13–14 embryos. A CUY21EDIT II electroporator (BEX Co., Ltd., Japan) was used. Electric pulses consisted of one short pulse (25 V, 0.05 ms) and a 0.1-ms interval, followed by five long pulses (8 V, 10 ms) with 1-ms intervals. The electric pulses were applied during injection of the DNA solution.

RA treatment

S. canicula embryos were removed from their egg shells, then placed into 6-well plates. 4–6 ml of artificial seawater containing penicillin/streptomycin was used for culturing embryos. RA was dissolved in DMSO to 2 mg/ml as a stock solution and diluted in the artificial seawater to 1–2 μg/ml. 1% DMSO in the artificial seawater was used as negative controls. Embryos at stage 28–29 were cultured with RA for 4 days for gene expression analyses. For alcian blue staining, embryos at stage 28–29 were cultured with 1–2 μg/ml RA for 20 days and additional 10–18 days after removing RA. Note that effect of RA is highly dependent on individual embryos. Some batches of embryos were lethal at 2 μg/ml of RA, probably due to season or parents' condition. In this case, embryos were treated with 1 μg/ml of RA.

Acknowledgements

We thank T Ogura, J Miyazaki, A Kuroiwa, H Ogino, and H Ochi for providing the plasmids; A Tweedale and Station Biologique de Roscoff for collecting S. canicula embryos; JA Donald, T Toop, and JD Bell for collecting C. milii embryos; M Komada for providing cell lines; M Davy, X Xu, S Kuratani, and C Tickle for comments; and K Munakata, S Ueda, and N Suda for technical advice. Sequences for Alx4, Pax9, Gli3, Hand1, Tbx2, Zic3, Ptch1, and element 1586 from S. canicula and Gli3 from C. milii are deposited in GenBank under accession numbers KC507187–9, KF748129, KP055651-KP055653, KF297619, and KF297620. Correspondence and requests for materials should be addressed to M Tanaka (mitanaka@bio.titech.ac.jp) and J Sharpe (james.sharpe@crg.eu). This work was supported in part by the Global COE Program ‘Evolving Education and Research Center for Spatio-Temporal Biological Network’ from the Ministry of Education, Culture, Sports, Science and Technology (MEXT) to KO and MT, the Japan–Australia Research Cooperative Program to SH, the Grant-in-Aid for Scientific Research on Innovative Areas, the Grant-in-Aid for Scientific Research (B) and the Inamori Foundation to MT, ICREA and the Spanish Ministry of Economy and Competitiveness, ‘Centro de Excelencia Severo Ochoa 2013-2017’, SEV-2012-0208 to JS and the Centre for Genomic Regulation to KO and JS.

Funding Statement

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Funding Information

This paper was supported by the following grants:

  • Japan Society for the Promotion of Science (JSPS) Grant-in-Aid for Scientific Research (B) to Mikiko Tanaka.

  • Ministry of Education, Culture, Sports, Science, and Technology (MEXT) Grant-in-Aid for Scientific Research on Innovative Areas to Mikiko Tanaka.

  • Japan Society for the Promotion of Science (JSPS) Japan-Australia Research Cooperative Program to Susumu Hyodo.

  • Spanish Ministry of Economy and Competitiveness (MINECO) Plan Nacional Grant (BFU2010-16428) to James Sharpe.

Additional information

Competing interests

The authors declare that no competing interests exist.

Author contributions

KO, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article, Contributed unpublished essential data or reagents.

SK, Acquisition of data, Analysis and interpretation of data, Contributed unpublished essential data or reagents.

WT, Acquisition of data, Contributed unpublished essential data or reagents.

SH, Acquisition of data, Contributed unpublished essential data or reagents.

JS, Conception and design, Analysis and interpretation of data, Drafting or revising the article.

MT, Conception and design, Analysis and interpretation of data, Drafting or revising the article.

Ethics

Animal experimentation: Experiments were performed in accordance with guidelines for animal experiments of Tokyo Tech and CRG, and experiments involved in mice were approved by Animal Ethics Committees of CRS (No. JMC-07-1001P3-JS).

Additional files

Major datasets

The following datasets were generated:

Onimaru K, Tanaka M, 2014, Scyliorhinus canicula Alx4 mRNA, complete cds, http://www.ncbi.nlm.nih.gov/nuccore/KC507187, Publicly available at the NCBI Nucleotide (Accession no: KC507187).

Onimaru K, Tanaka M, 2014, Scyliorhinus canicula Pax9 mRNA, partial cds, http://www.ncbi.nlm.nih.gov/nuccore/KC507188, Publicly available at the NCBI Nucleotide (Accession no: KC507188).

Onimaru K, Tanaka M, 2014, Scyliorhinus canicula Gli3 mRNA, partial cds, http://www.ncbi.nlm.nih.gov/nuccore/KC507189, Publicly available at the NCBI Nucleotide (Accession no: KC507189).

Onimaru K, Kuraku S, Takagi W, Hyodo S, Tanaka M, 2014, Scyliorhinus canicula element 1586, genomic sequence, a fin/limb enhancer of Gli3, http://www.ncbi.nlm.nih.gov/nuccore/KF297619, Publicly available at the NCBI Nucleotide (Accession no: KF297619).

Onimaru K, Kuraku S, Takagi W, Hyodo S, Tanaka M, 2014, Scyliorhinus canicula Hand1 mRNA, partial cds, http://www.ncbi.nlm.nih.gov/nuccore/KF748129, Publicly available at the NCBI Nucleotide (Accession no: KF748129).

Onimaru K, Kuraku S, Takagi W, Hyodo S, Sharpe J, Tanaka M, 2015, S. canicula Zic3 mRNA, partial cds, http://www.ncbi.nlm.nih.gov/nuccore/KP055652, Publicly available at the NCBI Nucleotide (Accession no: KP055652).

Onimaru K, Kuraku S, Takagi W, Hyodo S, Sharpe J, Tanaka M, 2015, S. canicula Tbx2 mRNA, partial cds, http://www.ncbi.nlm.nih.gov/nuccore/KP055651, Publicly available at the NCBI Nucleotide (Accession no: KP055651).

Onimaru K, Kuraku S, Takagi W, Hyodo S, Sharpe J, Tanaka M, 2015, S. canicula Ptch1 mRNA, partial cds, http://www.ncbi.nlm.nih.gov/nuccore/KP055653, Publicly available at the NCBI Nucleotide (Accession no: KP055653).

Onimaru K, Kuraku S, Takagi W, Hyodo S, Tanaka M, 2015, C. milii Gli3 mRNA, partial cds, http://www.ncbi.nlm.nih.gov/nuccore/KF297620, Publicly available at the NCBI Nucleotide (Accession no: KF297620).

The following previously published datasets were used:

Tanaka M, Munsterberg A, Anderson WG, Prescott AR, Hazon N, Tickle C, 2002, Scyliorhinus canicula dHand protein mRNA, partial cds, http://www.ncbi.nlm.nih.gov/nuccore/AY057890, Publicly available at the NCBI Nucleotide (Accession no: AY057890).

Sakamoto K, Onimaru K, Munakata K, Suda N, Tamura M, Ochi H, Tanaka M, 2009, Scyliorhinus canicula HoxA13 mRNA, partial cds, http://www.ncbi.nlm.nih.gov/nuccore/EU005550, Publicly available at the NCBI Nucleotide (Accession no: EU005550).

Muller TS, Ebensperger C, Neubuser A, Koseki H, Balling R, Christ B, Wilting J, 2015, Gallus gallus paired box 9 (PAX9), mRNA, http://www.ncbi.nlm.nih.gov/nuccore/NM_204912, Publicly available at the NCBI Nucleotide (Accession no: NM_204912).

Takahashi M, Tamura K, Buscher D, Masuya H, Yonei-Tamura S, Matsumoto K, Naitoh-Matsuo M, Takeuchi J, Ogura K, Shiroishi T, Ogura T, Izpisua Belmonte JC, 2013, Gallus gallus ALX homeobox 4 (ALX4), mRNA, http://www.ncbi.nlm.nih.gov/nuccore/NM_204162, Publicly available at the NCBI Nucleotide (Accession no: NM_204162).

Bollag RJ, Siegfried Z, Cebra-Thomas J, Garvey N, Davison EM, Silver LM, 1994, Mus musculus T-box 2 (Tbx2), mRNA, http://www.ncbi.nlm.nih.gov/nuccore/120407038, Publicly available at the NCBI Nucleotide (Accession no: NM_009324).

Zhang G, Cohn MJ, 2009, Scyliorhinus canicula Sox9 (Sox9) mRNA, complete cds, http://www.ncbi.nlm.nih.gov/nuccore/EU241880, Publicly available at the NCBI Nucleotide (Accession no: EU241880).

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eLife. 2015 Aug 18;4:e07048. doi: 10.7554/eLife.07048.012

Decision letter

Editor: Marianne E Bronner1

eLife posts the editorial decision letter and author response on a selection of the published articles (subject to the approval of the authors). An edited version of the letter sent to the authors after peer review is shown, indicating the substantive concerns or comments; minor concerns are not usually shown. Reviewers have the opportunity to discuss the decision before the letter is sent (see review process). Similarly, the author response typically shows only responses to the major concerns raised by the reviewers.

Thank you for sending your work entitled “A shift in anterior-posterior positional information underlies the fin-to-limb evolution” for consideration at eLife. Your article has been evaluated by Diethard Tautz (Senior editor), Marianne Bronner (Reviewing editor), and three reviewers.

The Reviewing editor and the other reviewers discussed their comments before we reached this decision, and the Reviewing editor has assembled the following comments to help you prepare a revised submission.

In this paper, the authors analyze fin development in Chondricthian fish from the catshark, Scyliorhinus canicula. Specifically they show a lack of an elaboration of genes in the catfish that are known to impart polarity along the anterior posterior axis of mice and chicken. As this polarity has been tied to the specification and patterning of the tetrapod limb, the authors ask if this shift is genetically regulated within the catshark through Gli3 transcriptional regulation and further if perturbation of general AP signaling is sufficient for altering chondricthian fin development in a manner consistent with AP polarity of early tetrapod lineages. Of note they discover a functional enhancer for Gli3 that is retained in tetrapods but not seen in chondricthians that they argue has been a key development in the evolution of limbs.

This is an issue that has received intense interest from both paleontologists and developmental biologists and thus should be of general interest to the readership of eLife. Although the paper is interesting and elegantly done, there are several major issues that cloud their conclusions and therefore need to be addressed.

1) The idea that there is a broadening of the AP axis in the evolution of tetrapod limbs is not new. Shubin and Alberg (1986) and Oster, et al. (1988) outlined a metapterygial arch hypothesis that outlined a shift in AP axis as a key mechanism leading to evolution of the tertrapod limb. They supported this hypothesis with developmental and comparative analysis. Thus, finding molecular correlates of this pattern is not altogether unexpected, although the data in the present manuscript are the most extensive analysis of the patterning changes in chondricthians. Surprisingly, these papers are neither cited nor discussed, and must be included.

2) The data on “posteriorizing” of the Scyliorhinus fin through retinoic acid is not convincing. The authors present quite elegant experiments that uncover a limb-specific Gli3 regulator domain. However the cartilage staining of the treated fins does not look anything similar to the author's interpretation – and does not phenocopy an early tetrapod such as Tiktaalik. Even if the remnant posterior fragment can be argued to be stylopodial, the authors must explain how this occurred as they have shown the lack of a key functional Gli3 regulatory domain in Scyliorhinus that is necessary for the regulation of polarity in mouse and chick – presumably even after RA treatment. Unlike the elaborate elements formed in the miniskate (Dahn et al. 2007), the data here make it seem like cartilage development has been delayed or inhibited. Even though dHand expansion is expanded after RA treatment the changes in domains are small and this may be due to developmental delay or independent effects of RA such as on proliferation (note the fin size is quite different than controls at the same stage), rather changes in polarity.

3) The expression patterns in the fins are quite late in comparison with comparable sized limbs such that these signaling events may pattern more distal structures. Changes in patterning of proximal bones such as the humerus would be expected to occur earlier, such as HH stage 20 in the chick. Could the authors comment on how such late expression changes would be expected to alter early developmental patterning events in the shark?

4) Correlation of positional value shift (expression boundaries) in fin/limb bud to resultant adult morphology isn't very tight. The one example that we're shown of an experimentally RA-dosed cat shark pectoral fin is juxtaposed with the pectoral fin of Tiktaalic. What we're not shown is the adult fin morphology of Callorhinchus, which is also dominated by the metapterygium (and its various, more distal, radials). Callorhinchus pectoral fin skeletons are morphologically like those of Tiktaalik and its various finned relatives.

5) ‘Partial posteriorisation’ (the authors' term) is a repeated phenomenon in the broad spectrum of paired fin evolution. It looks like this results in a ‘necessary but insufficient’ item of the fin-to-limb agenda, and likewise it might or might not be part of the history of chimaeroid fins. For example, how does Callorhinchus have a metapterygium-dominated pectoral fin, but seems not to have employed/evolved the positional value shift at the heart of this study?

6) The connection between the part of the paper that paper that documents the changes in Gli3 expression and function are not clearly linked to the final part of the paper that uses RA treatment to shift AP patterning. The authors need to make a clearer statement about exactly how they link together these observations using the experimental manipulations they have done. It seems that the RA treatment is a very indirect way to try and link together the observations on Gli3 to the actual effects of shifting AP patterning. The link seems very indirect and RA treatment is bound to have many effects that are independent of the changes in Gli3. Would the authors expect that the reduction of Shh signaling in a tetrapod increase the number of bones in the anterior part of the limb, or better yet what would happen if they expressed Gli3 in a more posterior region? At the very least, the authors need to make a stronger link here. Without this, the work would not be particularly compelling of broad interest.

[Editors' note: further revisions were requested prior to acceptance, as described below.]

Thank you for resubmitting your work entitled “A shift in anterior-posterior positional information underlies the fin-to-limb evolution” for peer review at eLife. Your revised submission has been favorably evaluated by Diethard Tautz (Senior editor), Marianne Bronner (Reviewing editor), and one of the original reviewers.

Summary:

The authors have addressed the majority of the previous concerns, but one comment should be addressed more directly.

1) The reviewers remain unconvinced of the interpretation of the RA work and identity of the resulting elements. As this is a key component of their conclusions that alteration of RA/Shh signaling can cause reduction of the pro/mesopterygium, similar to the fin-limb morphological transition, it is important that this is clear to the reader to be able to interpret their findings. For example the severe RA treated fin shown has a strongly stained element (comparable to the proximal elements of the DMSO treated fin) on the anterior side – not compatible with a reduction of a pro/meso component of the fin. Given the lightly staining of the treated fins compared with the control, it would be helpful if the authors could make a schematic of their interpretation of the resulting pattern and explain why there are patterning/staining differences.

2) The title for Figure 4 should reflect that it is RA mediated signaling that was tested not Shh. Shh levels or activity were not directly tested by gene over-expression or like means.

3) Results and discussion, sixth paragraph, the conclusions that “loss of the anterior proximal elements during evolution appears partially ‘driven’ by cis regulatory changes” has not been directly shown. What is shown is that changes are associated with morphological transitions and may have been a component of the changes leading to the evolution of these forms. They also may have been secondary and not directly involved.

eLife. 2015 Aug 18;4:e07048. doi: 10.7554/eLife.07048.013

Author response

1) The idea that there is a broadening of the AP axis in the evolution of tetrapod limbs is not new. Shubin and Alberg (1986) and Oster, et al. (1988) outlined a metapterygial arch hypothesis that outlined a shift in AP axis as a key mechanism leading to evolution of the tertrapod limb. They supported this hypothesis with developmental and comparative analysis. Thus, finding molecular correlates of this pattern is not altogether unexpected, although the data in the present manuscript are the most extensive analysis of the patterning changes in chondricthians. Surprisingly, these papers are neither cited nor discussed, and must be included.

As suggested, we have added discussion about the digital arch model in the seventh paragraph of the Results and discussion section.

2) The data on “posteriorizing” of the Scyliorhinus fin through retinoic acid is not convincing. The authors present quite elegant experiments that uncover a limb-specific Gli3 regulator domain. However the cartilage staining of the treated fins does not look anything similar to the author's interpretation – and does not phenocopy an early tetrapod such as Tiktaalik.

We agree that the RA-treated skeletons are not a direct phenocopy of Tikitaalik and we have therefore removed that panel from Figure 4, and also replaced the scheme of evolutionary process into a comparison between specific species (S. canicula and mice). Nevertheless, our important observation remains the striking correlation between the anterior shift of gene expression patterns, and the fusion of basal bones into a single metapterygium. In the context of the fin-to-limb transition, this correlation is a very important scientific observation.

Even if the remnant posterior fragment can be argued to be stylopodial, the authors must explain how this occurred as they have shown the lack of a key functional Gli3 regulatory domain in Scyliorhinus that is necessary for the regulation of polarity in mouse and chick – presumably even after RA treatment.

We agree that this part may have been confusing – in particular we may not have distinguished clearly enough between regulation at the transcriptional level versus the protein activity level. On the one hand, we show that Gli3 in S. canicula is lacking a relevant cis-regulatory element to restrict its expression anteriorly. On the other hand, we also show that RA is indeed able to cause a shift in AP patterning for Ptch1, Hand2 and Pax9. However, there is no contradiction here: The knowledge that RA treatment increases Shh signalling in fin and limb buds is well-known from the literature (Riddle et al., 1993 Cell 75: 1401-1416; Hoffman et al., 2002 Int. J. Dev. Biol. 46: 949-956; Dahn et al., 2007 Nature 445: 311-314). In turn, the effect of Shh signalling on Gli3 is not an alteration in transcriptional regulation, but rather a modification of the protein, specifically inhibiting formation of the repressor form (which is responsible for repressing Hand2 in the anterior tissue; Vokes et al. 2008 Genes Dev. 22: 2651-2663). This reduced repressive activity allows Hand2 to be upregulated, causing the general AP shift. Thus we propose that the effect seen in the RA-treated embryos is a post-translational regulation of Gli3 – not transcriptional. In other words, the AP shift during evolution was partially driven by cis-regulatory changes, while by contrast in our RA experiments it was driven by a Shh-mediated effect on the Gli3 protein itself, but in both cases achieving similar phenotypic changes. We have made this much clearer in the main manuscript now (Results and discussion, fifth and sixth paragraphs).

Unlike the elaborate elements formed in the miniskate (Dahn et al. 2007), the data here make it seem like cartilage development has been delayed or inhibited. Even though dHand expansion is expanded after RA treatment the changes in domains are small and this may be due to developmental delay or independent effects of RA such as on proliferation (note the fin size is quite different than controls at the same stage), rather changes in polarity.

We agree it is important to address the potential non-specific effect of RA experiments, for example on general growth and development of the bud (even in this non-model system which is non-trivial to work with). We have therefore performed new experimental works to address this issue (Figure 4 and Results and discussion, fifth paragraph).

Firstly, we addressed whether the treated fin buds appear to have retarded patterning, by examining another molecular marker which reveals the progress of proximo-distal patterning – Hoxa13. We detected slightly weaker expression in RA treated embryos than that in control embryos. However, the expression domain of Hoxa13 was not obviously shifted (Figure 4E), showing that this dosage and timing of RA treatment is having a clear effect on AP patterning (the shifts of Ptch1, Hand2 and Pax9, Figure 4A-C), while causing no obvious effects on PD patterning or general growth.

Secondly, we re-evaluated the sizes of fin buds from the RA-treated experiments, and also repeated the experiment to increase the replicates of Hand2 analysis. The image in the original panel of this figure was slightly distorted due to the in situ hybridisation process. From further analysis of all RA-treated results, it is clear that we observed the anterior expansion of Hand2 expression in buds of a similar size to the controls (in Figure 4B).

Since the reviewer considered that the change in Hand2 expression was small, we have also chosen to boost the result by analysis of another AP marker – Pax9 which is expressed in the anterior tissue (Figure 4C). As with Hand2 and Ptch1, this marker also displays a “posteriorisation” (or anterior shift of positional information) as it’s graded levels are reduced after RA treatment. Also important to note is that again the size of the fin bud is not reduced.

3) The expression patterns in the fins are quite late in comparison with comparable sized limbs such that these signaling events may pattern more distal structures. Changes in patterning of proximal bones such as the humerus would be expected to occur earlier, such as HH stage 20 in the chick. Could the authors comment on how such late expression changes would be expected to alter early developmental patterning events in the shark?

Indeed, we agree that the timing of these patterning processes is important point to address, and so we have added new gene expression data to clarify the time point at which the proximal skeletal elements begin to be patterned. Figure 1–figure supplement 2E now shows that Sox9 expression starts in the proximal elements between stage 28 and late stage 29. This is the same period for which we show the more posteriorly-biased gene expression patterns in S. canicula (Figure 1–figure supplement A-D) and also the RA-treatment experiments (Figure 4).

4) Correlation of positional value shift (expression boundaries) in fin/limb bud to resultant adult morphology isn't very tight. The one example that we're shown of an experimentally RA-dosed cat shark pectoral fin is juxtaposed with the pectoral fin of Tiktaalic. What we're not shown is the adult fin morphology of Callorhinchus, which is also dominated by the metapterygium (and its various, more distal, radials). Callorhinchus pectoral fin skeletons are morphologically like those of Tiktaalik and its various finned relatives.

Although the fin skeleton of Callorhinchus has a strong metapterygium, it also clearly has a large propterygium, which is in contrast to Tiktaalik, which only has a single proximal element (metapterygium/ stylopod). To address this comment we have therefore followed the reviewers’ suggestion, and generated a new cartilage staining of Callorhinchus and included it in Figure 2B.

5) ‘Partial posteriorisation’ (the authors' term) is a repeated phenomenon in the broad spectrum of paired fin evolution. It looks like this results in a ‘necessary but insufficient’ item of the fin-to-limb agenda, and likewise it might or might not be part of the history of chimaeroid fins. For example, how does Callorhinchus have a metapterygium-dominated pectoral fin, but seems not to have employed/evolved the positional value shift at the heart of this study?

We agree that the loss of anterior elements is a repeated phenomenon. However, we do not believe this has happened in the Callorhinchus milii pectoral fin skeleton (which we now show in Figure 2B), because its skeleton still has a large proximal propterygium (annotated as “pro”). We therefore see no clear evidence against our hypothesis that an AP shift was involved in the fin-to-limb transition.

6) The connection between the part of the paper that paper that documents the changes in Gli3 expression and function are not clearly linked to the final part of the paper that uses RA treatment to shift AP patterning. The authors need to make a clearer statement about exactly how they link together these observations using the experimental manipulations they have done. It seems that the RA treatment is a very indirect way to try and link together the observations on Gli3 to the actual effects of shifting AP patterning. The link seems very indirect and RA treatment is bound to have many effects that are independent of the changes in Gli3.

We agree that RA treatment is an indirect way to perturb AP patterning, although we must point out (a) that experimental embryology in a non-model species like this is challenging, and also (b) that precisely this method for boosting Shh signalling has been used in important previous papers about fin patterning in the mini-skate (Dahn et al., 2007 Nature 445: 311-314) although here we are drawing novel and distinct conclusions about the shift in AP patterning (which nevertheless do not contradict this previous paper).

To further address this general point, we have now strengthened and clarified the manuscript in the following ways (Results and discussion, fifth and sixth paragraphs):

Firstly, we realised that our proposal may have been confusing (the link between the results on Gli3 regulation and the RA experiments) because in Figure 3 we show that S. caniculus lacks the cis-regulatory elements to be inhibited in the posterior tissue, and yet in the RA experiments we report that an AP shift is induced. This point is discussed above (point 2). Essentially, we did not explain the known relation between Gli3 and RA, and we did not distinguish strongly enough in the text between transcriptional regulation and post-translational regulation. We have now re-written this to make it clear: At the genetic/evolutionary level, our results suggest that acquisition of element 1586 was a way to shift Gli3 expression anteriorly. However, in the experiments extensive Shh signalling triggered by RA causes a similar AP shift, but through a different mechanism inhibiting production of Gli3 repressor form (rather than altered regulation of Gli3 transcription). The important result is that we have two cases of an anterior shift: (a) the comparison between sharks and tetrapods, (b) the RA-treated shark buds, and in both cases two important observations are correlated: (i) the anterior shifting of genes, (ii) a reduction or loss of anterior basal bones, leaving a metapterygium-dominated arrangement.

Secondly, we have addressed the problem of possible non-specific effects of RA (as described above for point 2). We have examined Hoxa13 expression in RA treated embryos to assess possible effects on proximo-distal (PD) patterning, which is another major role of RA in tetrapod limb buds. Although expression of Hoxa13 in RA treated embryos was slightly weaker than in control embryos, nevertheless PD patterning appears hardly affected compared to the clear AP shifts in Hand2 and Pax9 expression (as the PD boundary of Hoxa13 expression is the same as controls, Figure 4E).

Would the authors expect that the reduction of Shh signaling in a tetrapod increase the number of bones in the anterior part of the limb, or better yet what would happen if they expressed Gli3 in a more posterior region? At the very least, the authors need to make a stronger link here. Without this, the work would not be particularly compelling of broad interest.

Gli3 overexpression in a posterior limb bud may not be enough to increase the number of the proximal elements (stylopod), because the phenotype in the stylopod always appears in combination with Gli3 and other gene knockouts. Nevertheless, it is clear that in these cases the expected result (of element reduction) is indeed seen: Gli3-/-;Plzf-/- mice lack the femur (Barna, Pandolfi, and Niswander, 2005 Nature 436, 277-281), Gli3-/;Alx4-/- mice show reduction of humerus (Panman et al., 2005 Int. J. Dev. Biol. 49, 443-448). Therefore, to increase the number of stylopod elements (i.e. to get limbs back to the ancestral condition), Gli3 and additional genes are likely required. Since Alx4 and Hand2 are already expressed in the S. canicula pectoral fin bud, and Plzf is involved only in the hindlimb bud, we do not have any candidates that might be involved in the fin-to-limb evolution. Although S. canicula genome is not sequenced yet, systematic studies at whole genome level such as ChIP-seq analysis in S. canicula fin bud would provide a more complete picture of evolutionary mechanism of the loss of the anterior elements in the future (Results and discussion, eighth paragraph).

[Editors' note: further revisions were requested prior to acceptance, as described below.]

1) The reviewers remain unconvinced of the interpretation of the RA work and identity of the resulting elements. As this is a key component of their conclusions that alteration of RA/Shh signaling can cause reduction of the pro/mesopterygium, similar to the fin-limb morphological transition, it is important that this is clear to the reader to be able to interpret their findings. For example the severe RA treated fin shown has a strongly stained element (comparable to the proximal elements of the DMSO treated fin) on the anterior side – not compatible with a reduction of a pro/meso component of the fin. Given the lightly staining of the treated fins compared with the control, it would be helpful if the authors could make a schematic of their interpretation of the resulting pattern and explain why there are patterning/staining differences.

As suggested, we have added a schematic of our interpretation to Figure 4F. In particular, since the element on the anterior side of metapterygium in the severe phenotype is not directly attached to the pectoral girdle, we interpret it as a fused radial attached to the metapterygium (indicated by ** in Figure 4F). Therefore, we believe that the skeletal pattern of the severe phenotype represents a loss of anterior proximal elements. We have added the details of our interpretation to the main text as well (Results and discussion, fifth paragraph).

The different levels of staining between controls and RA treated fins seem to reflect individual differences in exposure time to Alcian Blue, as the other control sample show a lighter staining. We have therefore replaced the control figure with the lighter stained sample (Figure 4F).

2) The title for Figure 4 should reflect that it is RA mediated signaling that was tested not Shh. Shh levels or activity were not directly tested by gene over-expression or like means.

We have changed the title of Figure 4 into “RA treatment causes ectopic activation of Shh signaling and loss of anterior skeletal elements.”

3) Results and discussion, sixth paragraph, the conclusions that “loss of the anterior proximal elements during evolution appears partially ‘driven’ by cis regulatory changes” has not been directly shown. What is shown is that changes are associated with morphological transitions and may have been a component of the changes leading to the evolution of these forms. They also may have been secondary and not directly involved.

As suggested, we have changed the conclusion into “while the loss of the anterior proximal elements during evolution was associated with cis-regulatory changes of Gli3…” (Results and discussion, sixth paragraph).

Associated Data

    This section collects any data citations, data availability statements, or supplementary materials included in this article.

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