Actinotrichia-independent developmental mechanisms of spiny rays facilitate the morphological diversification of Acanthomorpha fish fins

Domain: neuroscience
Published: 2026-02-14
Editorial outcome: minor_revision

Abstract
Skeletal forms in vertebrates have been regarded as good models of morphological diversification. Fish fins show great diversity in form, with their supporting skeletal structure being classified into soft rays and spiny rays. In fish evolution, spiny-ray morphologies are known to be sometimes extremely modified; however, it remains unknown how the developmental mechanisms of spiny rays have contributed to their morphological diversification. By using the rainbowfish Melanotaenia praecox for examination of the extracellular matrix (ECM) and cell dynamics of spiny-ray development, we demonstrate that spiny-ray development is independent of the actinotrichia (needle-shaped collagen polymers at the tip of fins), which are known as an important ECM in soft-ray morphogenesis. Furthermore, we found that in the thorny spiny ray of the filefish Stephanolepis cirrhifer, the lateral protrusions are associated with BMP-positive osteoblast condensation, as in the spiny-ray tips in M. praecox and S. cirrhifer. Taken together, our findings reveal that osteoblast distribution and signaling-molecule intensity would contribute to spiny-ray modification. In comparison to soft ray development, the independence from actinotrichia in spiny rays would facilitate growth direction change, leading to their morphological diversification. This suggests that variation in cell distribution and ECM usage may be important contributors to morphological diversification, not only in Acanthomorpha, but also in other animal taxa.

Introduction
Understanding the fundamental mechanisms of morphological diversification is an ultimate goal of evolutionary biology. Skeletal forms in vertebrates are highly divergent1–3, and the molecular and genetic mechanisms of skeletal development have been extensively studied. Although skeletal forms serve as a good model of morphological diversification, the contribution of underlying developmental mechanisms to morphological diversification remains unknown.

Fish fin bones are among the most diversified organs among vertebrate skeletal forms and have acquired various functions and facilitate adaptations to various habitats4,5. In teleosts, the fins are commonly supported by bone structures called fin rays1,3,6–9, which are further classified into soft rays and spiny rays8,10–12 (Fig. 1a). Soft rays are flexible, segmented, and typically branched, whereas spiny rays are stiff, unsegmented, and terminate in an acute point5,11,13–15. Spiny rays have evolved independently in several lineages, such as the order Siluriformes and the clade Acanthomorpha11,15–17 (Fig. 1b). The Acanthomorpha is one of the largest lineages of vertebrates18–21, and their spiny rays are called “true spines”5,7,11,18,22. Interestingly, true spines are known to be extremely modified to form novel structures that do not work as fin components but as species-specific apparatuses. For example, pectinated lamellae, which form the sucking disc of remoras23,24, and illicium, which is the fishing apparatus of goosefishes and anglerfishes8,25, originated from the spiny rays of ancestral fish. Spiny rays could be regarded as an easily diversified skeletal feature, and have long fascinated scientists26. In contrast, soft rays sometimes evolved to be elongated in several lineages (e.g., in the pectoral fins of flying fish, and the dorsal fin of sailfish), but tend not to have been modified into non-rod-like shapes in their evolution5. Thus, the spiny-ray morphology is considered highly diversified5. Presently, little is known about factors that lead to this difference in the morphological diversity between spiny and soft rays. Developmental mechanisms of spiny rays have been largely unknown, although the development and regeneration process of soft rays have been well studied27–34. While it is possible that the differences in the developmental mechanisms between soft- and spiny-ray morphologies might be a candidate cause, we have not yet understood the contribution of spiny-ray developmental mechanisms to their morphological diversification.

Therefore, to uncover the developmental mechanisms of spiny rays, we sought to examine their morphogenesis at the cellular and molecular levels using laboratory animals. However, a critical problem was that popular model fish, such as zebrafish and medaka, could not be used for spiny-ray research. This is because zebrafish are not an Acanthomorpha and lack this structure11 and medaka have degenerated their spiny rays in the dorsal fin and have only one small spiny ray in their anal fin35. Consequently, for this study, we chose the dwarf neon rainbowfish Melanotaenia praecox, a small freshwater fish in the order Atheriniformes (Fig. 1c), which has spiny rays in the dorsal, anal, pectoral, and pelvic fins12, as a new model fish. Furthermore, we have already established a staging system for its postembryonic development based on fin development12, as well as a genome editing system for M. praecox36.

Here, we examined the extracellular matrix (ECM) and cell dynamics of spiny-ray development in M. praecox and compared them with those of soft-ray development. Our data reveal several differences in developmental mechanisms between spiny and soft rays, such as their dependence on or independence from the actinotrichia, which are known as an important ECM structure in zebrafish fin-ray growth and regeneration31,32,37,38. To gain insight into spiny-ray modification in Acanthomorpha evolution, we also investigated the dynamics of premature osteoblasts and bone morphogenetic protein (BMP) signaling in these cells in the filefish Stephanolepis cirrhifer, which has extremely modified spiny rays on its back, as well as in M. praecox. Based on our observations, we propose that the developmental mechanisms of spiny rays may have facilitated their morphological diversification.

Results
Growth dynamics of the spiny ray
Soft rays grow by adding bone matrix to the distal tip34. However, the growth patterns of spiny rays are largely unknown. To identify differences in morphogenesis between soft and spiny rays, we first visualized their developmental changes using two different staining methods12. We first used alizarin red for staining calcified bone in M. praecox dorsal-fin stage 4 (DF-st4) larvae12, of which some spiny rays were observed in their first dorsal fin (Fig. 1d); we then re-stained the bones with calcein, a green-fluorescent molecule, 5 days after the first staining with alizarin red (Fig. 1e). Hence, this two-color staining of the larvae (DF-st4 larvae, n=10) showed the morphology of their spiny rays at two time points (Fig. 1e–g). By comparing the morphology at different time points, we found that in each spiny ray, the calcein-stained area extended distally from the alizarin red/calcein double-positive area (Fig. 1f, g). This result suggests that the spiny rays mainly grow by adding bone matrix to their distal tips. Furthermore, the spiny rays also grow radially by adding bone matrix, since the calcein single-positive area surrounded the alizarin red/calcein double-positive area. The dorsal view showed that the spiny rays were composed of left and right elements at the time of alizarin red staining, and a thick single bone element at the time of calcein staining (Fig. 1f, g). The right and left elements seem fused by the radial growth of each element, which is consistent with previous morphological descriptions of the spiny-ray character5,15.

Actinotrichia do not play a crucial role in spiny-ray growth
Bone staining of M. praecox larvae with two different-colored staining molecules revealed that the spiny rays grow by adding bone matrix to their distal tips, which is similar to the growth process observed in soft-ray growth32,34,39. Thus, spiny rays may share some growth mechanisms with soft rays. In soft rays, actinotrichia (needle-shaped collagen polymers) are bundled at their distal tips and serve as scaffolds for mesenchymal cell migration during bone formation in fins29,31–33,37–42. To understand the function of spiny-ray development, we focused on the actinotrichia distribution in M. praecox fin fold and fins by using DAFFM (diaminofluorescein-FM) staining32,43,44 at the early stages of soft- and spiny-ray growth (Fig. 2a). DAFFM is a small fluorescent molecule that has been broadly used for NO detection, and recent research has shown that DAFFM can bind to collagenous structures like actinotrichia32,43–46. At the tips of soft rays in the second dorsal fin of larvae and juveniles (DF-st3 larvae, n=3; DF-st4 larvae, n=4; juveniles, n=4), we found that actinotrichia were arranged and bundled during soft-ray development (Fig. 2a). Before the emergence of spiny rays (i.e., in DF-st3 larvae), relatively small and less developed actinotrichia, compared with those at the tips of the soft rays, were observed in the median fin fold, including regions that later give rise to the spiny rays. At the initial stage of spiny-ray development (at DF-st4), fine actinotrichia were aligned in parallel with spiny-ray primordia, but these were not distributed at the tip of each spiny ray, as seen in the soft rays. At the juvenile stage, actinotrichia were not detected around the spiny rays (Fig. 2a). DAFFM staining visualizes collagenous structures, including actinotrichia, bone collagen, and tendon of the segments in the soft rays32,44. Therefore, the bright signals in Fig. 2a correspond mainly to actinotrichia, bone collagen, and tendon of the segments in the soft rays.

We further confirmed the production of actinotrichia around spiny rays in M. praecox by determining the expression of the gene actinodin1 (and1) and actinodin2 (and2), which encodes proteins known to be essential structural components of actinotrichia28,37,38. At DF-st4 (and1, DF-st4 larvae, n=6; and2, DF-st4 larvae, n=4) and the juvenile stage (and1, juvenile, n=4; and2, juvenile, n=4), and1 and and2 were strongly expressed at the tip of each soft ray in the second dorsal fin (Supplementary Fig. 1). However, at both stages, there was no and1 expression and no or weak and2 expression around spiny rays in the first and second dorsal fins (Supplementary Fig. 1). These results indicate that, at the tips of spiny rays, actinotrichia are neither actively formed nor maintained.

It is reported that actinotrichia in the fin folds were absent in zebrafish and1 and and2 morphants38. Therefore, we investigated this in M. praecox by first making a double knockout (KO) of genes and1/and2 using the CRISPR/Cas system (Supplementary Figs. 9 and 10). We isolated some types of mutant fish with several deletions in the and1/and2 genes, each of which causes a frame-shift mutation and is expected to encode a non-functional truncated protein (Supplementary Figs. 2 and 3). Next, to assess the role of the actinotrichia in spiny-ray growth, we generated mutant M. praecox with the genotype and1–/–/and2+/– (DF-st4 or juveniles with an SL<1 cm, n=6) (Supplementary Figs. 2 and 3), for comparison of the actinotrichia condition and soft/spiny-ray morphologies in wild-type fish (DF-st4 or juveniles with a SL<1 cm, n=8) (Fig. 2b) and the mutant fish (Fig. 2c). In mutant fish images of the DAFFM and alizarin red staining, we observed that the actinotrichia were not radially arranged as in wild-type fish (Fig. 2b), and that the soft rays were distorted and curled in the mutant fish (white arrowheads in Fig. 2c and Supplementary Fig. 4). We observed that the intervals of the segments of the soft rays were uncertain (yellow arrowheads in Fig. 2c and Supplementary Fig. 2). The restriction of fin ray growth may result in the shortening of each soft ray. The curled phenotypes were not always observed in the mutant fish, with this phenomenon being caused by the edge of the caudal part of the fin fold failing to grow. Rather, the mutants had a normal, straight spiny-ray bone morphology similar to that of the wild-type fish (Fig. 2b, c). Furthermore, we generated M. praecox mutants with the genotype and1+/–; and2–/– (DF-st4 or juveniles with SL<1 cm, n=2) (Supplementary Figs. 5 and 10). In these mutants, we observed the actinotrichia were radially arranged and that the soft rays were slightly distorted (white arrowhead in Supplementary Fig. 5). Although weak and2 expression was detected around the spiny rays in some in situ hybridization specimens (Supplementary Fig. 1), the spiny rays of the and1+/–; and2–/– mutants were not affected by the and2 knockout (Supplementary Fig. 5). Regarding the function of the actinotrichia, our observations strongly suggest that actinotrichia do not play a role in the growth and morphogenesis of spiny rays (Fig. 2d). The actinotrichia seen early in the first dorsal fin in DF-st4 larvae could be what remains from supporting the median fin fold before that stage, as seen in the larval median fin fold of zebrafish, where actinotrichia are present and surmised to be essential for fin-fold development and maintenance32,38,40,47. Considering this together, the presence of actinotrichia around the spiny rays in the DF-st4 larvae would be consistent with the condition in the mutant fish. Therefore, we next asked how spiny rays could achieve distal growth independently of the actinotrichia.

Mesenchymal cell condensation at the spiny-ray tips
To explore the growth mechanisms of spiny rays, we observed the cellular conditions at the tips of spiny rays in comparison with those of soft rays. First, we examined the 3D cell distribution at the tips of spiny rays (DF-st4 larvae, n=3) and soft rays (DF-st4 larvae, n=3) by staining the nucleus and collagen structures simultaneously. DAR-4M is a small fluorescent molecule similar to DAF-FM that is widely used for NO detection; however, the two molecules exhibit different fluorescence emission wavelengths32,44. Recent studies have shown that DAR-4M can bind to collagenous structures such as actinotrichia32,46. We found that mesenchymal cells are multilayered at the tips of spiny rays (Fig. 3a-left), whereas such condensed mesenchymal cells were not detected at the tips of soft rays (Fig. 3a-right).

We next observed the detailed conditions of the mesenchymal cells and ECM. Transmission electron microscopy (TEM) images of transverse sections of soft rays (DF-st4 larvae, n=3) revealed several thick actinotrichia (magenta coloration) aligned between the two hemirays (light blue coloration), with a few mesenchymal cells in the space between the hemirays, as previously reported in zebrafish48 (Supplementary Fig. 6). Furthermore, the hemirays closely adjoined the basement membrane, and no mesenchymal cells were distributed between the hemirays and the basement membrane. TEM images of transverse sections of spiny rays (DF-st4 larvae, n=3) (Fig. 3b) showed very few or no actinotrichia, consistent with our staining observations indicating the absence of actinotrichia (Fig. 2a). Mesenchymal cells (green coloration) were highly condensed surrounding the spiny ray (orange coloration), and the layer composed of basement membrane (yellow coloration) and basal lamella49,50 (hot pink coloration) surrounded this cell condensation (Fig. 3b). Furthermore, the basement membrane around spiny rays was thicker than the basement membrane seen next to soft rays (Supplementary Fig. 6). We then focused on the distribution of laminin proteins, a major component of the basement membrane51,52. Three-dimensional reconstructions of immunohistochemistry analysis at the tip of a spiny rays showed that the signal for laminin specifically accumulated beneath the TP63-positive epidermal basal cells (DF-st4 larvae, n=9). The distribution of laminin strongly suggests that the laminin layer surrounding the mesenchymal condensation at the tips of spiny rays in TEM images (Fig. 3b) is the basement membrane, with basal lamella forming a cap-like structure on the spiny-ray tips (Fig. 3c). This structure was not detected at the tips of soft rays (Supplementary Fig. 6), nor has it been reported in developing and regenerating soft rays where the actinotrichia are distributed at the tip.

The differences in the ECM and cellular conditions of developing spiny and soft rays are shown in Fig. 4 and demonstrate that the developmental mechanism of the spiny rays is independent of actinotrichia, which are a vital factor for the straight growth of soft rays. Rather, it is possible that the condensed mesenchymal cells at the tip of developing spiny rays may play a pivotal role in spiny-ray bone formation.

BMP signaling for proper osteogenesis of spiny rays
Given that mesenchymal cells surround the tips of spiny rays, we considered that they may contribute to the morphogenesis of the spiny ray through their osteogenesis. Therefore, we examined the osteoblast distribution and osteogenic signaling pathways in these cells. The upstream osteogenic gene runx253 is known to be expressed in pre-osteoblasts at the distal tip of regenerative blastema in the caudal fin of zebrafish54,55. Moreover, BMP signaling promotes osteoblast differentiation56. Accordingly, we next observed the distribution of premature osteoblasts and BMP signaling in the mesenchymal cell condensation around the tip of the spiny ray in M. praecox larvae using immunohistochemistry (DF-st4 larvae, n=9) (Fig. 5). Runx2 signals were specifically detected in the cell condensations of the distal tips of the spiny rays, and active Smad1, 5, and 9 (pSmad1/5/9) signals were detected in some cells of the Runx2-positive cells (Fig. 5a). This suggests that the cell condensation at the distal tips of spiny rays was composed of pre-osteoblasts, and that BMP signaling may contribute to the osteogenesis of spiny rays.

To verify the role of BMP signaling in spiny-ray osteogenesis, we administrated a BMP-receptor inhibitor, DMH1. In DMH1-treated fish (DF-st4 larvae, n=8), pSmad 1/5/9 signals were lost in the cell condensation, although they were clearly detected in control fish (DF-st4 larvae, n=7) (Fig. 5b), thereby indicating that DMH1 properly inhibited BMP signaling at the spiny-ray tip (Fig. 5b). In the immunohistochemistry images for pSmad1/5/9 (leftmost panels of Fig. 5b), several punctate signals are visible in both control and DMH1-treated fish. In previous studies using the same antibody, pSmad1/5/9 immunoreactivity was prominent in nuclei57,58. Therefore, the punctate signals observed in our images may represent non-specific staining. Surprisingly, the tips of spiny rays in the DMH1-treated fish were largely deformed and much thicker than in the control fish, where several cells were buried in the deformed spiny-ray bone in some of the treated specimens (DF-st4 larvae, n=5/8) (blue arrowheads in Fig. 5b), which we did not see in the spiny rays of the controls (DF-st4 larvae, n=0/7) (Fig. 5b). Despite BMP signaling inhibition by DMH1, the spiny rays retained their characteristic unsegmented and rigid morphology. No actinotrichia-like collagenous structures or segmentation were observed at the tips of the spiny rays in treated specimens (Fig. 5b). Taken together, these results suggest that premature osteoblasts condense at the spiny-ray tip, which is the most readily growing part of this bone, and that BMP signaling in these cells is required for proper spiny-ray morphogenesis.

Density of mesenchymal cells in modified spiny rays
Developmental mechanisms of spiny rays in M. praecox imply that the distribution of premature osteoblasts and working BMP signaling should be a key factor for regulating the spiny ray morphology. Based on these findings, we hypothesized that alternation of the osteoblast distribution and BMP signaling in these cells have resulted in morphologically modified spiny rays in the course of Acanthomorpha fish diversification (Fig. 6). To test the hypothesis, we observed the premature osteoblast cell distribution of spiny rays in the filefish Stephanolepis cirrhifer (Fig. 6a), a member of the order Tetraodontiformes (Fig. 1b). Larva of S. cirrhifer has a thorny dorsal spine, which is a modified spiny ray (Fig. 6a, b, CT-image, 0.98 cm and 1.3 cm SL, n=2). Runx2-positive premature osteoblasts were condensed at the tip of the dorsal spine in S. cirrhifer (Fig. 6c and Supplementary Fig. 7, 1.2–1.35 cm SL, n = 3), and some of these cells had pSmad1/5/9 signals (yellow arrowhead in Fig. 6c and Supplementary Fig. 7), as seen in the spiny-ray tips in M. praecox larvae (Fig. 5a). Furthermore, in protrusions (thorn primordia) that branch from the stem of a dorsal spine, premature osteoblasts (Fig. 6c and Supplementary Fig. 7) specifically form a thin layer, and pSmad1/5/9 positive signals (yellow arrowhead in Fig. 6c and Supplementary Fig. 7) were detected in these cells. The cellular conditions of the S. cirrhifer suggest that the mechanisms adding bone at the tip of the stem of that spiny ray also act on the tip of the protrusions in the thorny dorsal spine, consistent with our hypothesis.

Methods
Animal husbandry and embryo culture
Melanotaenia praecox were hatched and maintained in a laboratory at Tohoku University, as previously described12,36. Adults were held in 1-L or 3-L tanks, at ~28°C, a pH slightly greater than 7.0, and under a light:dark cycle of 14:10 h. Hatched larvae were transferred to a 250-mL rearing tank, at a density of 1–15 individuals per tank. Adults were fed live brine shrimp once or twice daily. Depending on the progeny size, larvae were fed live Paramecium at least once every 2 days, and/or brine shrimp at least once daily. Removal of dead larvae and excreta was performed as required. After the juvenile stage, the progenies were moved to larger tanks (≥250 mL). When observing the dorsal fin bone morphologies of young male and female fish (male, n=3; female, n=3), whose sexes are distinguishable by their body coloration36, we did not find any significant differences in the fin bone morphologies between males and females at this stage (Supplementary Fig. 8). Furthermore, we did not have a reliable technique to distinguish the sex of larval and juvenile M. praecox using PCR-based methods or other investigations, and thus did not distinguish the sex of the larval and juveniles specimens in our experiments.

Larvae of Stephanolepis cirrhifer were captured at Shoubutahama Fishing Harbor in Miyagi, Japan (38°16′55″N, 141°03′41″E). Fish identification followed the work of Okiyama64. Although sex chromosome system and sexual dimorphisms in the adult second dorsal fin have been reported in this species65,66, we did not distinguish the sex of our specimens because a reliable PCR-based genotyping method has not yet been established. According to Kwon et al.67, sexual maturation in this species occurs at a total length of approximately 11.7 cm. The specimens used in our experiments were all significantly smaller than this size and would thus not have been expected to have developed sexual dimorphisms. Furthermore, sexual dimorphisms of the dorsal spine have not been reported. Taken together, we concluded that distinguishing the sex of our S. cirrhifer specimens was unnecessary.

Animal care experimental procedures were conducted in accordance with institutional and national guidelines and regulations, and were approved by the Tohoku University Animal Research Committee (permit number 2022LsA-002-10). The study was carried out in compliance with ARRIVE guidelines.

X-ray micro-CT scanning
The skeletal structures of M. praecox adults (Male, n=3) and S. cirrhifer larvae (0.98 cm SL & 1.2 cm SL, n=2) were observed using X-ray micro-computed tomography (CT) scanning, as previously described in Akama et al.68. Samples of M. praecox adults were fixed with 10% formaldehyde at room temperature overnight, followed by dehydration using an ethanol series (50% EtOH, 70% EtOH, 80% EtOH, 90% EtOH, 95% EtOH, 100% EtOH, 95% EtOH, 90% EtOH, 80% EtOH, 70% EtOH). These samples were then stored in 70% ethanol. Samples of S. cirrhifer larvae were fixed with 4% paraformaldehyde (PFA) at 4°C overnight, and then transferred to 100% methanol. Using an X-ray micro-CT scanner (ScanXmate-D180RSS270, modified by Voxel Works Co., Tokyo, Japan and VOXIA room VR150V-HH-TAVE, Voxel Works Co., Tokyo, Japan), the fixed specimens were scanned at conditions shown in Supplementary Table 1. The micro-CT data were reconstructed using coneCTexpress (White Rabbit Corp.). Three-dimensional image analysis was performed with Molcer v.1.8.5.1 (White Rabbit Corp.).

Small molecule staining
Live larvae were stained to examine bone12 (double staining, DF-st4, n=10; only calcein staining, young male, n=3; only calcein staining, young female, n=3) and actinotrichia32 (WT, DF-st4 or juveniles with a SL<1 cm, n=8; and1–/–/and2+/–, DF-st4 or juveniles with an SL<1 cm, n=6; and1+/–/and2–/–, DF-st4 or juveniles with an SL<1 cm, n=2; and1–/–/and2+/+, DF-st4 or juveniles with an SL<1 cm, n=1) as described previously, with minor modifications. To observe the actinotrichia, vital staining with 5 µM of DAFFM DA (Goryo Chemical, #SK1004-01) was used. Larvae of M. praecox were immersed in the DAFFM DA solution overnight at room temperature. After staining of the actinotrichia and/or bone, the larvae were washed twice with system water. Thereafter, staining of bone with alizarin red was conducted. For observations, larvae were anesthetized with 0.025% MS222/E3, placed on a 1% agarose gel/E3, transferred immediately to a small case filled with system water, and awakened with water. The stained live larvae were observed and photographed under a microscope (Leica M205FA, with Leica DFC360FX attachment). The images were analyzed using Leica LAS-AF software (Leica Microsystems Inc.), LAS-X software (Leica Microsystems Inc.), and Adobe Photoshop CS6.

To observe actinotrichia in fixed specimens, M. praecox larvae (DF-st3, n=3; DF-st4, n=4; juvenile, n=4) were fixed in 4% PFA overnight at 4°C. After washing two times in phosphate-buffered saline (PBS) with Tween 20 (PBT: 0.1% Tween 20 in PBS), specimens were stained with 10 µM of DAFFM (Goryo Chemical, #SK1003-01) overnight at room temperature, and then washed two times in PBT. To further clear and remove the pigment, the stained samples were immersed in 0.5% KOH/3% H2O2/PBT solution overnight at 4°C, and then washed three times in PBT. The head and abdomen were dissected from the samples to prepare them for observation. The specimens were placed on a glass slide, covered with a coverslip, lightly pressed, and observed under a confocal microscope system (Leica TCS SP5 II).

DAR4M and SYTO13 staining
To observe cells, actinotrichia, and the bone condition, M. praecox larvae (spiny ray, DF-st4, n=3; soft ray, DF-st4, n=3) were fixed in 4% PFA overnight at 4°C, and then dehydrated, as described previously9. Following rehydration, specimens were immersed in 0.5% KOH/3% H2O2 solution for 30 min to 1 h at room temperature. Next, the specimens were washed three times with PBT, and stained with SYTO13 green-fluorescent nucleic acid stain (Invitrogen, #S7575) for 1 h at room temperature. After washing three times with PBT, specimens were immersed in 10 µM of DAR4M (Goryo Chemical, #SKSK1005-01) overnight at 4°C. The next day, the specimens were again washed three times with PBT. Finally, each specimen was placed in a 1-well cell culture plate with PrimeGel Agarose LMT 1-20K GAT (Takara Bio, 5806A) and observed under a light-sheet microscope (QuVi SPIM, Luxendo).

Whole-mount in situ hybridization
Whole-mount in situ hybridization for gene expression analysis was performed as previously described69,70, with minor modifications. Samples of M. praecox (and1, DF-st4 larvae, n=6; and1, juvenile, n=4; and2, DF-st4 larvae, n=4; and2, juvenile, n=4) were fixed with 4% PFA/PBS at 4°C overnight, dehydrated in methanol/PBT, and stored at −30°C in methanol. The samples were then rehydrated progressively with 75% methanol/H2O, 50% methanol/H2O, 25% methanol/Tris-buffered saline with Tween 20 (TBST), and TBST, for 10 min each. Next, these samples were bleached with 3% H2O2 and 0.5% KOH in TBST for 1 h. After washing five times with TBST for 5 min, the samples were incubated in Proteinase K (Pro K, Invitrogen, #25530049) in TBST (20 µg/ml) for 1 h, rinsed once briefly with TBST, and washed once for 5 min with TBST. Pro K-treated samples were placed in 0.1 M triethanolamine (TEA, pH 7–8) and incubated for 10 min after adding 10 µl of acetic acid per 1 ml TEA. After washing five times with TBST for 5 min, samples were refixed with 4% PFA/PBS solution for 20 min, and washed five times with TBST for 5 min. The samples were placed in hybridization buffer (containing 50% formamide, 5× SSC, 0.1% Tween 20, 5 mg/ml of tRNA, 50 µg/ml of heparin, 2.5% blocking regent, and 10 mM of EDTA) at 65°C for 5 h, and then incubated in hybridization buffer containing 5–10 µg/ml of probe, overnight, at 65°C. Probe-hybridized samples were washed twice with 50% formamide/5× SSC/0.1% Tween 20/10 mM of EDTA at 65°C for 30 min, three times with 2× SSC/0.1% Tween 20 at 65°C for 20 min each, and three times with 0.2× SSC/0.1% Tween 20 at 65°C for 20 min each. After washing twice with maleic acid buffer for 5 min, and blocking with blocking buffer (1% blocking regent/10% heat-inactivated goat serum in maleic acid buffer) for 1.5 h, the samples were incubated in blocking buffer containing 1:3000 anti-digoxigenin-AP Fab fragments (Roche #11093274910) for 5 h. These samples were washed with maleic acid buffer as follows: twice for 5 min, three times for 20 min, once overnight, four times for 2 h, and once overnight. After washing three times with TBST for 5 min, the samples were incubated in AP buffer (0.1 M Tris base at pH 9.5, 0.1 M NaCl, 0.1% Tween 20). The samples were stained with BM-Purple (Roche #11442074001) in the dark, at room temperature, for 1–7 days. The staining reaction was stopped by washing the samples three times for 10 min with PBT. All steps were performed on a shaking platform.

For in vitro transcription of the digoxigenin (DIG)-labeled RNA probe, the polymerase SP6 (Roche #10810274001) or T7 RNA (Invitrogen #18033019) was used, along with a DIG RNA labeling mix (Roche #11277073910), followed by a deoxyribonuclease step and ethanol precipitation. The labeled in situ RNA probes were resuspended in 50% formamide and stored at −30°C.

Molecular cloning and sequencing
Molecular cloning and sequencing were performed following the protocol outlined in a previous study36, with slight modifications. A primer table is provided in Supplementary Table 2. Melanotaenia praecox sequences were identified by BLAST (Basic Local Alignment Search Tool) against genome data for Boeseman’s rainbowfish, Melanotaenia boesemani.

Generation of CRISPR/Cas9 and1/and2 mutant M. praecox lines
To investigate the functions of the actinotrichia in vivo, CRISPR/Cas-induced knockout fish were generated. In accordance with a previously described method36, four types of sgRNAs (Supplementary Figs. 2 and 3, and Supplementary Table 3) for CRISPR/Cas were prepared, and microinjection into M. praecox fertilized eggs was performed (Supplementary Fig. 9a). We prepared a 5-µl solution containing 100 ng of each sgRNA, 1,250 ng of Cas9 protein, and 0.5 µl of phenol red in nuclease-free water, and then 2–3 nL of this solution were injected into the single cell of the embryo.

The injected fish (G0) with fin bone phenotype were in-crossed to produce a generation of homozygous and1 knockout (and1–/–) and heterozygous and2 knockout (and2+/–) fish (F1) (Supplementary Fig. 9). Although the homozygous and1/and2 knockout fish (and1–/–; and2–/–) was the most ideal for this study, this genotype could not be reared, probably because this knockout mutation is lethal or causes a high mortality rate. To determine the sequences of mutant alleles, the genomic region, including the target site of sgRNAs was amplified with KOD One® PCR Master Mix -Blue- polymerase (Toyobo #KMM-201) using the primer pairs listed in Supplementary Table 2. The PCR conditions were as follows: one cycle at 94°C for 2 min, followed by 35 cycles of 98°C for 10 sec, 60°C for 5 sec, and 68°C for 15 sec. Obtained fragments were cloned using a TOPO Cloning Kit (Invitrogen #K460001) or TArget Clone™ -Plus- (TAK-201) and sequenced by Sanger sequencing. Following the genotyping conducted by the above sequence analysis, after observation and genotyping of the F1 fish, we in-crossed selected single pairs of F1 fish that had fin phenotype and and1–/–/and2+/– genotypes, and we obtained and1–/–/and2+/– fish (F2) (Supplementary Fig. 9a). Some of the F2 fish obtained from single pairs of F1 fish had a few short actinotrichia in their fins (Supplementary Fig. 9b) and we confirmed that one of these fish genotypes was homozygous and1 knockout only (and1–/–/and2+/+) (Supplementary Fig. 9c). This analysis suggests that homozygous and1 knockout alone does not cause the actinotrichia loss. Then, we only used fish that lost radially arranged actinotrichia as in wild type fish and had and1–/–/and2+/– genotypes for our analysis (Fig. 2b, c). The sequences of mutant alleles of F2 fish using observation were also determined by the above sequence analysis. Detailed information on the and1/and2 mutations in the F1 and F2 fish is summarized in Supplementary Figs. 2 and 3. To further analyze the and1+/–; and2–/– fish, we first outcrossed F1 individuals with the genotype and1–/–; and2+/– to wild-type fish, obtaining F2 fish with the genotype and1+/–; and2+/– (Supplementary Fig. 10). Next, we crossed two types of F2 individuals—those with and1–/–; and2+/– genotypes, and those with and1+/–; and2+/– genotypes—to generate F3 offspring. For our analyses, we used only F3 fish that had the and1+/–; and2–/– genotype (Supplementary Fig. 10). The mutant alleles of the analyzed F3 fish were confirmed by sequence analysis, and detailed information on the and1 and and2 mutations is summarized in Supplementary Fig. 5b, c. In this study, because some alleles with missense mutations are not expected to encode a non-functional truncated protein, we denote those alleles with the symbol “+”.

TEM analysis
Preparation of epoxy blocks was performed as described previously71, with minor modifications. Larvae of M. praecox (spiny ray, DF-st4, n=3; soft ray, DF-st4, n=3) were fixed in 2% glutaraldehyde and 2% paraformaldehyde in 0.01 M PBS at 4°C overnight. The fixed specimens were post-fixed in 1% OsO4 in 0.01 M PBS at 4°C overnight, after decalcification in 10% ethylenediaminetetraacetic acid in 0.01 M PBS at 4°C for 4 days. The specimens were then embedded in epoxy resin (Epon 812; TAAB Laboratories Equipment Ltd) by polymerizing overnight at 37°C, and again overnight at 60°C after dehydration through a graded ethanol series followed by a transition in propylene oxide.

The epoxy blocks were trimmed, and serial sections (70 nm in thickness) were cut with an ultramicrotome (Leica, EM UC7) using a Histo Diamond Knife (Syntek, SYM3045 Ultra). The sections were picked up and mounted on a grid (Veco, Square Mesh Handle 150 mesh Nickel Grid). Next, the sections were stained with 4% uranyl acetate for 15 min and then washed three times with Mili-Q water. After drying, they were stained with lead citrate staining solution (2% w/v trisodium citrate dihydrate, 1% w/v lead (II) citrate trihydrate, 1% w/v lead (II) acetate, 1% w/v lead (II) nitrate in 0.18 mol/L sodium hydroxide solution) and then washed three times with Mili-Q water. Finally, the specimens were observed under a transmission electron microscope (JEOL, JEM 1400).

Immunohistochemistry
A whole-mount immunohistochemistry protocol was used to detect laminin (Sigma-Aldrich #L9393), TP63 (Abcam, #ab735), Runx2 (Santa Cruz Biotechnology, #sc-101145), and pSmad1/5/9 (Cell Signaling Technology, #13820). Whole-mount immunohistochemistry was performed as described previously9, with minor modifications, until primary antibody incubation steps were different between staining laminin and TP63, and staining Runx2 and/or pSmad1/5/9.

To stain laminin and TP63, M. praecox larvae (DF-st4, n=9) were fixed with absolute methanol. The samples were rehydrated with methanol/PBT, blocked with blocking buffer (1% BSA, 1% goat serum, and 1% DMSO in PBS) and stained with a 1:50 dilution of anti-laminin antibody and 1:50 dilution of anti-TP63 antibody in blocking buffer overnight. Samples were then washed five times in PBT, blocked with blocking buffer, and incubated with a 1:500 dilution of Alexa Fluor 488 goat anti-rabbit (Invitrogen, #A-11008) and Alexa Fluor 594 goat anti-mouse (Invitrogen, #A-11032). After washing five times in PBT, samples were stained with a 0.001% w/v DAPI (4′,6-Diamidino-2-phenylindole dihydrochloride; Dojindo, #342-07431) for 1 h, and then washed three times in PBT. The head and abdomen were dissected from the samples to prepare them for observations. The samples were placed on a glass slide, covered with a coverslip, lightly pressed, and observed under a confocal microscope system (Leica TCS SP5 II).

To stain Runx2 and/or pSmad1/5/9, M. praecox (DF-st4, n=9) and S. cirrhifer larvae (1.2–1.35 cm SL, n=3) were fixed with 4% PFA/PBS, dehydrated with methanol/PBT, and stocked in absolute methanol at −20°C. Following incubation in absolute acetone for 20 min, the samples were rehydrated with methanol/PBT, permeabilized with 1% Triton X-100, blocked with blocking buffer (1% BSA, 1% goat serum, and 1% DMSO in PBT) and stained with a 1:50 dilution of anti-Runx2 antibody and/or 1:100 dilution of anti-pSmad1/5/9 antibody in blocking buffer overnight. Secondary antibody and DAPI staining procedures are shown above, and Alexa Fluor 488 goat anti-rabbit (Invitrogen, #A-11008), Alexa Fluor 488 goat anti-mouse (Invitrogen, #A-11001), and Alexa Fluor 594 goat anti-rabbit (Invitrogen, #A-11012) were used as the secondary antibodies. DMH1-treated specimens (DMH1 treated, DF-st4, n=7; control, DF-st4, n=8) were additionally stained with DAR4M after the DAPI staining.

DMH1 treatment experiments
Inhibition of BMP signaling was performed with 5 µM of DMH1 (Selleckchem #S7146, dissolved in DMSO to 20 mM). DMH1 stock solution was dissolved in E3 water72. The M. praecox larvae (DF-st4, n=7) were treated with DMH1 in the dark at 28°C for 10 days, subsequently washed with system water three times, and fixed with 4% PFA for phenotype analysis. Mock treatments (DF-st4, n=8) were performed using 0.025% DMSO, which does not result in phenotypic changes.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "In this study, Miyamoto et al. demonstrated that the developmental mechanisms of spiny rays in Acanthomorpha fish are independent of actinotrichia, in contrast to those of soft rays, by introducing the rainbowfish Melanotaenia praecox as a new model organism. Through transmission electron microscopy (TEM) analysis and immunostaining, the authors identified the condensation of mesenchymal cells and the presence of premature Runx2/pSmad-positive osteoblasts at the tips of spiny rays. Additionally, they found that Runx2/pSmad-positive cells were similarly distributed in the tips and lateral protrusions of spiny rays in Stephanolepis cirrhifer. Regarding the authors' primary claim presented in the title—that actinotrichia-independent developmental mechanisms “facilitate” morphological diversification in Acanthomorpha fish—I feel that the supporting results appear to provide correlative evidence rather than strong proof. However, this study offers valuable insights into the developmental mechanisms of spiny rays, which contribute to the highly diversified morphology observed in some species; nevertheless, the following points should be addressed further.\n\nIn situ hybridization reveals slight expression of actinodin2 but no detectable expression of actinodin1 in the developing spiny rays of Melanotaenia praecox (Extended Fig 1). The and1-/- and2+/- phenotype is presented in this manuscript; however, there is no description of homozygous mutants of and2. Including the and2-/- or and1+/- and2-/- phenotypes would further clarify the independence of actinotrichia, which is one of the main findings of the paper.\n\nThe phenotype of the and1-/- and and2+/- mutants is intriguing. The authors briefly noted that the soft rays are deformed in the mutants; however, could a more detailed analysis be conducted? In the mutants, the soft rays do not appear to be segmented. Additionally, is the elongation of the soft rays affected in the mutants? Why do they curl up in the caudal fin? I understand this is somewhat tangential to the main topic of the paper, but a more detailed analysis could yield new insights into soft ray formation by actinotrichia.\n\nRegarding Figure 5b, a previous study by Hoch et al. (PNAS 2021) demonstrated that DMH1 treatment causes spiny rays in cichlid fish to transform into soft rays. Perhaps the DMH1 treatments were carried out after the initial development of the spiny ray, but that should be properly described. In this study, larvae were incubated for 10 days, which is a considerable duration. Can the authors rule out the possibility that DMH1 treatments caused the spiny ray to develop a soft ray-like structure, resulting in the observed deformation? Do DMH1-treated spiny rays retain the properties of spiny rays? Additionally, I believe the study by Hoch et al. is highly relevant to this paper and influences the authors' work. While their paper is cited in this manuscript (11), a more detailed description is needed. Their findings, which illustrate the establishment of spiny and soft-rayed fin domains in acanthomorph fishes, provide essential molecular insights into the differences between spiny and soft-rayed fins.\n\nIn Figure 6, the Runx2/pSmad signals largely overlap in the regions surrounding the tips of the laterally protruding spiny ray in filefish. In contrast, Figures 5a and 5b, which utilize Melanotaenia praecox, show less overlap in the Runx2/pSmad signals, particularly at the tips of the spiny fins. Could this discrepancy be attributed to the presentation of optical sections? The images should be updated to display the z-stack images to clearly show the distributions of the pSmad signals.\n\nHow many larvae of Stephanolepis cirrhifer were examined using immunostaining? The staining results from other individuals should be presented as supplementary data to confirm reproducibility.\n\nRegarding the evolution of spiny rays, do the authors think they evolved from soft rays by losing actinotrichia, or were they acquired independently?\n\n(Minor points)\nIn Fig. 1a, spiny and soft rays stained with Alizarin red are shown. Please specify which fin rays belong to which fish.\nLines 530-532: Was 5 µl injected into each embryo? That seems like a large volume; please confirm.\nFigure 5b: Are the green signals on several dots non-specific? Please clarify this point.\nLine 210: While both DAFFM and DAR4M are claimed to detect actinotrichia, I am unclear about the differences between them.", "reviewer_2": "The manuscript entitled “Actinotrichia-independent developmental mechanisms of spiny rays facilitate the morphological diversification of Acanthomorpha fish fin”, submitted to Nature Communications, investigates developmental differences between spiny rays and soft rays in rainbowfish (Melanotaenia preacox), with additional comparative studies from the highly modified spiny rays of the filefish (Stephanolepis cirrhifer).\n\nTo address their objectives, the authors employ a comprehensive suite of techniques, including micro-CT, electron microscopy, immunofluorescence, and in-situ hybridization, to demonstrate key distinctions in fin ray development and anatomy at different scales (from the entire spine to the cellular level). Specifically, they show that spiny rays are formed by an accumulation of mesenchymal cells where BMP signaling plays an important role, which differs from soft rays that rely on actinotrichia as a structural scaffold. Furthermore, the authors identify similar spiny-ray developmental mechanisms in the filefish, suggesting that this actinotrichia-independent mode of growth may have facilitated the morphological diversification of fin rays within Acanthomorpha.\n\nOverall, the manuscript is well-executed and clearly written, with well-organized flow of ideas. The experimental design is robust to address the question proposed (although see general comments about sample size and sexes used), and the integration of the multiple methods provides a holistic and convincing analysis. The figures clearly show the results and are easy to follow. The research question is original, particularly because little is known about the skeletal development of acanthomorphs. Therefore, this research is of interests across multiple fields, including fish biology, physiology, skeletal development, and evolution. However, I suggest the authors to revise my general comments regarding this topic.\n\nMy main concern is the lack of discussion and consideration of the bone type the authors are studying (acellular bone) as well as comparing to (cellular bone), which I consider that is crucial to understand the results and do an accurate comparative analysis. I hope my review will help further improve the clarity and impact of this work.\n\nGeneral comments\nAcellular bone skeletogenesis.\nOne aspect that stood out to me is that the authors do not mention, or consider, the type of bone that forms the skeleton of both species of fishes that are studied here. The majority of Neoteleostei, and thus acanthomorps (except some species like tunas) have acellular bone (i.e. a bone without osteocytes). This type of bone is also found in some Euteleostei. In contrast, more basal groups of fishes are characterized by cellular bone (i.e. a bone with osteocytes), including Ostariophysi where cypriniformes (zebrafishes) and siluriformes (catfishes) are part of.\n\nThis difference in bone type could be one of the reasons why the development of spiny rays in zebrafish and catfish (species that authors use to compare their results) may differ from that in the species studied here. Therefore, the spiny ray development described in the manuscript may probably be related with the periosteal osteogenesis of acellular bone. Check also the anatomical terminology used (e.g. basement membranes), as some of the structures may have been already described in Moss (1961). This point can be a very interesting topic to include and discuss, since the formation of acellular bone has not been largely studied.\n\nFor these reasons, I suggest that the authors consider and discuss the acellular bone formation in the manuscript, as well as to be cautions in describing this developmental process as “unique” to Acanthomorph spiny rays, as what the authors may be describing reflect general features of acellular bone skeletogenesis rather than a distinct or novel pathway of spiny rays. Additionally, the importance of BMP signaling, while certainly relevant, is not surprising as this superfamily of proteins is well known to have a crucial role during bone development across vertebrates.\n\nBelow I included several papers that discuss the differences between acellular or cellular bone that the authors may find useful.\n\nSample size.\nThe authors specify sizes in each section of the Results, but not in Methods. I recommend also including a detailed breakdown in the Methods section, indicating the number of animals used at each developmental stage and for each technique (e.g. How many S. cirrhifer and M. praecox where µCT-scanned?, this is not specifically clarified anywhere). Additionally, in some instances, it is unclear which ontogenetic stage is used for a given specific technique (larvae, juveniles, adults?). Lastly, in some cases it is unknown whether the reported sample size in Results refers to the number of rays examined or the number of individual animals. Since multiple rays can be studied from a single specimen, it would be helpful to clarify the number of animals studied similar to how it is done in L130.\n\nAnimal sex.\nThe authors do not specify whether the specimens used were males or females. Several teleosts, including M. preacox, are sexual dimorphic, with males exhibiting enlarged caudal spines.\n\nSpecific comments:\nAbstract\n- L22. The hyphenation of “spiny rays” and “soft rays” varies throughout the manuscript. In some instances, the terms are hyphenated (e.g., lines 23, 24, 27), while in others they are not (e.g., lines 22, 48, 49, 56). Please check for consistency and revise accordingly.\n- L22. “their bone structure” is a vague descriptor in this context. What about “their supporting skeletal structure”.\n- L30. “the lateral protrusions equip the BMP positive osteoblast condensation…”. The verb “equip” here is a bit confusing. What about: “the lateral protrusions are associated with BMP-positive osteoblast condensation”\n- L36. “… ECM usage would be major factors driving the morphological diversification in animals”. I recommend focusing this sentence specifically on acanthomorphs, rather than using the broader term “animals”.\n\nIntroduction\n- L48. “…and mostly branching,…”. Maybe replace “mostly branching” to “typically branched”.\n- L54 – 55. Are the sucking disc of remoras or the illicium considered “spiny rays”? or they have been so modified that they are no longer considered as such?\n- L72. “…there anal fin.” There is a typo here, it should be “their anal fin”.\n- L78. “…with that of soft-ray development.” Those of soft-ray development instead, to be consistent with the plural subject (“matrix and cell dynamics”).\n\nResults\n- L92 – 94. This sentence reads awkwardly and may be either missing some information or be poorly structured. I recommend revising it for clarity, as its current form makes the intended meaning difficult to follow.\n- L114, Figure 1. It is “yellow” instead of “white”.\n- L128. “DAFFM is a small fluorescent molecule that broadly used for..”. Instead: “DAFFM is a small fluorescent molecule that has been broadly used for…”.\n- L131. Delete “obviously”.\n- L132. Regarding the statement “fine actinotrichia were present at the future position of spiny rays in the median fin fold”. This statement is not clearly supported by the figure. In the image (Fig 2a), the actinotrichia do not appear to be specifically localized at the future position of the spiny rays. In fact, it doesn’t look like they are found in a specific position. I recommend clarifying the figure or revising the interpretation in the text to better reflect what is shown.\n- L135 – 136. In the spiny and soft rays, the DAFFM only stains actinotrichia? If not, which molecules form the brightest bulk of the rays? See image below:\n- L209: “n=3”. The number of spiny tails (sample size) sampled are from 3 different animals?\n- L213: “…such condensed mesenchyme cells”. How the authors know that these are mesenchyme cells if they did not use any markers, only the staining of the nuclei? If the authors are defining mesenchyme cells based on their rounder, larger morphology, they could be also osteoblasts or other precursor cell?\n- Figure 3: There is a typo in SYTO13 (nucleus).\n\nDiscussion\n- L334: using the “unique” adjective here is unnecessary. The authors do not know if this mechanism is unique to spiny rays or to acanthomorps, since this process can be happening in other skeletal elements or in other clades (perhaps is how the acellular bone is formed?).\n- L344: Secondarily?\n- L361 – L363: The fact that the developmental process that forms the spiny rays described in this manuscript is different of the process of the formation of spiny rays in Siluriformes may be because catfishes have cellular bone (see my comments above).\n- L406 – L411: this text looks like doesn’t belong here?\n\nMethods\n- Add sample sizes for each technique (see my comment above).\n- L445: after fixation, where the samples directly transferred to 70% ETOH? Or the authors dehydrated them using increasing concentration steps?\n\nReferences\n- Davesne, D., Meunier, F. J., Schmitt, A. D., Friedman, M., Otero, Otero, O., & Benson, R. B. (2019). The phylogenetic origin and evolution of acellular bone in teleost fishes: insights into osteocyte function in bone metabolism. Biological Reviews, 94(4), 1338-1363.\n- Shahar, R., & Dean, M. N. (2013). The enigmas of bone without osteocytes. BoneKEy reports, 2, 343.\n- Moss, M. L. (1961). Osteogenesis of acellular teleost fish bone. American Journal of Anatomy, 108(1), 99-109.\n- Weiss, R. E., & Watabe, N. (1979). Studies on the biology of fish bone. III. Ultrastructure of osteogenesis and resorption in osteocytic (cellular) and anosteocytic (acellular) bones. Calcified tissue international, 28(1), 43-56.\n- Moss, M. L. (1963). The biology of acellular teleost bone. Annals of the New York Academy of Sciences, 109(1), 337-350.\n- Dean, M. N., & Shahar, R. (2012). The structure‐mechanics relationship and the response to load of the acellular bone of neoteleost fish: a review. Journal of Applied Ichthyology, 28(3), 320-329."}, "author_response": "Point-by-point responses\nWe thank you very much for reviewing our manuscript. We have revised it following careful\nexamination of your suggestions. Your comments helped us to greatly improve the quality of our paper,\nand we express our gratitude to both of you for your constructive suggestions.\nTo improve the overall readability, we have had the entire document professionally proofread.\nIn addition to responding to your comments, we found some mistakes in the explanation of the\nphylogenetic tree in the original manuscript. Therefore, we have revised the legend of Figure 1 and\nFigure 2 as below:\nLine 112-113, “b Simplified phylogenetic tree of the Teleostei (modified from Figs. 1 and 2 in\nGhezelayagh et al.19 and Fig. 2 in Hughes et al.39”\nLine 191-192, “All panels except the top-left panel represent magnified views of the area enclosed\nby the yellow dashed box in the top-left panel.”\nIn addition, we have also revised the product number of the staining molecule in our Methods section,\nas below:\nLine 572-573, “To observe the actinotrichia, vital staining with 5 µM of DAFFM DA (Goryo\nChemical, #SK1004-01) was used.”\nPlease find our responses to each comment from the reviewers below.\nReviewer #1 (Remarks to the Author):\nReviewer comment 1-1:\nIn this study, Miyamoto et al. demonstrated that the developmental mechanisms of spiny rays in\nAcanthomorpha fish are independent of actinotrichia, in contrast to those of soft rays, by introducing\nthe rainbowfish Melanotaenia praecox as a new model organism. Through transmission electron\nmicroscopy (TEM) analysis and immunostaining, the authors identified the condensation of\nmesenchymal cells and the presence of premature Runx2/pSmad-positive osteoblasts at the tips of spiny\nrays. Additionally, they found that Runx2/pSmad-positive cells were similarly distributed in the tips and\nlateral protrusions of spiny rays in Stephanolepis cirrhifer. Regarding the authors' primary claim\npresented in the title—that actinotrichia-independent developmental mechanisms “facilitate”\nmorphological diversification in Acanthomorpha fish—I feel that the supporting results appear to\nprovide correlative evidence rather than strong proof. However, this study offers valuable insights into\nthe developmental mechanisms of spiny rays, which contribute to the highly diversified morphology\nobserved in some species; nevertheless, the following points should be addressed further.\nResponse 1-1:\nWe thank the referee for the positive comments. We agree that our results provide correlative evidence\nrather than strong proof. Therefore, we have added a sentence to address this point.\nLine 402-405, “Although our observations of S. cirrhifer provide correlative rather than definitive\nevidence, the results presented here nevertheless suggest that modifications of the ancestral\ndevelopmental mechanisms of spiny rays may have triggered morphological diversification of\nthese structures in specific acanthomorph lineages (Fig. 7).”\nReviewer comment 1-2:\nIn situ hybridization reveals slight expression of actinodin2 but no detectable expression of actinodin1\nin the developing spiny rays of Melanotaenia praecox (Extended Fig 1). The and1-/- and2+/- phenotype\nis presented in this manuscript; however, there is no description of homozygous mutants of and2.\nIncluding the and2-/- or and1+/- and2-/- phenotypes would further clarify the independence of\nactinotrichia, which is one of the main findings of the paper.\nResponse 1-2:\nWe thank you for these helpful suggestions. To address these concerns, we generated and1⁺/⁻; and2⁻/⁻\nmutant fish and examined their fin bone morphology. We did not observe any differences in the spiny\nrays between and1⁺/⁻; and2⁻/⁻ mutants and wild-type fish. Our and1⁺/⁻; and2⁻/⁻ mutant data suggest that\nactinodin2 is unlikely to play a crucial role in spiny-ray morphogenesis. We have revised the manuscript\nto include this description and discussion.\nLine 169-172, “Furthermore, we generated M. praecox mutants with the genotype and1⁺/⁻; and2⁻/⁻\n(DF-st4 or juveniles with SL < 1 cm, n = 2) (Extended Data Fig. 5 and Fig. 10). In these mutants,\nwe observed the actinotrichia were radially arranged and that the soft rays were slightly distorted\n(white arrowhead in Extended Data Fig. 5).”\nExtended Data Figure 5 Actinotrichia distribution and spiny- and soft-ray bone morphology in\nthe actinodin1+/-/actinodin2-/- knockout Melanotaenia praecox.\nWhite arrowheads indicate abnormal bending of the soft rays in the knockout fish. Actinotrichia\nwere labeled with DAFFM DA (green), and the spiny- and soft-ray bones were labeled with\nalizarin red (magenta). All panels except the top-left panel represent magnified views of the area\nenclosed by the yellow dashed box in the top-left panel.\nLine 671-678, “To further analyze the and1⁺/⁻; and2⁻/⁻ fish, we first outcrossed F individuals with\n1\nthe genotype and1⁻/⁻; and2⁺/⁻ to wild-type fish, obtaining F fish with the genotype and1⁺/⁻; and2⁺/⁻\n2\n(Extended Data Fig. 10). Next, we crossed two types of F individuals—those with and1⁻/⁻; and2⁺/⁻\n2\ngenotypes, and those with and1⁺/⁻; and2⁺/⁻ genotypes—to generate F3 offspring. For our analyses,\nwe used only F3 fish that had the and1⁺/⁻; and2⁻/⁻ genotype (Extended Data Fig. 5). The mutant\nalleles of the analyzed F3 fish were confirmed by sequence analysis, and detailed information on\nthe and1 and and2 mutations is summarized in Extended Data Figs. 5b and 5c.”\nExtended Data Figure 10. Schematic illustration of the obtaining the actinodin1+/-/actinodin2-/-\nknockout fish.\nReviewer comment 1-3:\nThe phenotype of the and1-/- and and2+/- mutants is intriguing. The authors briefly noted that the soft\nrays are deformed in the mutants; however, could a more detailed analysis be conducted? In the mutants,\nthe soft rays do not appear to be segmented. Additionally, is the elongation of the soft rays affected in\nthe mutants? Why do they curl up in the caudal fin? I understand this is somewhat tangential to the\nmain topic of the paper, but a more detailed analysis could yield new insights into soft ray formation\nby actinotrichia.\nResponse 1-3:\nWe apologize for the insufficient explanation of the phenotypes of the and1⁻/⁻ and and2⁺/⁻ mutants. We\nare currently examining the function of the actinodin genes in fin-ray morphogenesis, and we plan to\nsubmit these findings as a separate article. Thus, in this article, we further added a brief description of\nthe fin-ray phenotype as follows.\nIn the mutant fish, we observed segmentation in the soft rays, although the spacing between segments\nwas uncertain. Restricted fin-ray growth may also contribute to the shortening of individual soft rays.\nThe curled phenotype was not consistently observed among mutant individuals, and appears to result\nfrom the posterior finfold margin failing to extend properly.\nTo more clearly describe these phenotypes, we have revised Figure 2 by adding yellow arrowheads\nindicating examples of soft-ray segmentation in the knockout fish. In addition, we have added\nexplanatory text to the manuscript and included new extended figures (Extended Data Fig. 4) showing\nmutant individuals without the curled caudal-fin phenotype.\nLine 164-167, “We observed that the intervals of the segments of the soft rays were uncertain\n(yellow arrowheads in Fig. 2c and Extended Data Fig. 2). The restriction of fin ray growth may\nresult in the shortening of each soft ray. The curled phenotypes were not always observed in the\nmutant fish, with this phenomenon being caused by the edge of the caudal part of the fin fold\nfailing to grow.”\nExtended Data Figure 4 Actinotrichia distribution and spiny- and soft-ray bone morphology in\nthe actinodin1-/-/actinodin2+/- knockout Melanotaenia praecox.\nWhite arrowheads indicate abnormal bending of the soft rays in the knockout fish. Yellow arrow\nheads show aberrantly positioned segmental structures in the soft rays of the knockout fish.\nActinotrichia were labeled with DAFFM DA (green), and the spiny- and soft-ray bones were\nlabeled with alizarin red (magenta).\nReviewer comment 1-4:\nRegarding Figure 5b, a previous study by Hoch et al. (PNAS 2021) demonstrated that DMH1 treatment\ncauses spiny rays in cichlid fish to transform into soft rays. Perhaps the DMH1 treatments were carried\nout after the initial development of the spiny ray, but that should be properly described. In this study,\nlarvae were incubated for 10 days, which is a considerable duration. Can the authors rule out the\npossibility that DMH1 treatments caused the spiny ray to develop a soft ray-like structure, resulting in\nthe observed deformation? Do DMH1-treated spiny rays retain the properties of spiny rays?\nResponse 1-4:\nThis is an important point, and we thank the reviewer for raising it. We carefully considered the\npossibility that DMH1 treatment caused the spiny rays to acquire soft ray-like characteristics. However,\nseveral lines of evidence suggest that this is unlikely.\nFirst, in the previous study by Hoch et al. (2021), cichlid fish were treated with DMH1 before the\nappearance of fin rays. In their experiment, inhibition of BMP signaling altered the future fate of fin\nmesenchymal cells. In contrast, in our study, we treated M. praecox individuals after the spiny and soft\nrays had already formed, thereby perturbing fin mesenchymal cells only after osteoblast differentiation\nhad occurred. Thus, the timing of BMP inhibition differs markedly between the two studies.\nFurthermore, if DMH1 treatment had induced a transition in osteoblast identity from a spiny-ray type\nto a soft-ray type, we would expect the transformed spiny rays to exhibit certain features characteristic\nof soft rays. For example, we would have detected actinotrichia-like collagenous structures or\nsegmentation within the transformed spiny rays. However, our analyses confirmed that none of these\nfeatures appeared in DMH1-treated spiny rays.\nWhile we cannot completely exclude the possibility of subtle changes in osteoblast identity, our results\ndo not support the hypothesis that DMH1 treatment induced a major transformation from spiny rays to\nsoft rays. We have revised the manuscript to incorporate this discussion.\nLine 336-339, “Despite BMP signaling inhibition by DMH1, the spiny rays retained their\ncharacteristic unsegmented and rigid morphology. No actinotrichia-like collagenous structures\nor segmentation were observed at the tips of the spiny rays in treated specimens (Fig. 5b).”\nLine 425-434, “Hoch et al. (2021)11 reported that inhibition of BMP signaling prior to fin-ray\nformation induced homeotic transformations of prospective spiny rays into soft rays. In contrast,\nour DMH1 treatment, applied after the spiny rays had already formed, did not result in any\napparent transformation of spiny rays into soft ray-like structures. Although we cannot entirely\nrule out minor shifts in osteoblast identity, the absence of segmentation or actinotrichia-like\ncollagenous structures in treated specimens indicates that BMP inhibition did not induce a major\ntransformation from spiny to soft rays once the spiny rays had formed. Taken together, these\nfindings suggest that BMP signaling primarily contributes to establishing the anterior–posterior\ndivision of the dorsal and anal fins into spiny- and soft-ray domains, but may not drive a\nqualitative switch in osteoblast identity after cell differentiation has occurred.”\nReviewer comment 1-5:\nAdditionally, I believe the study by Hoch et al. is highly relevant to this paper and influences the authors'\nwork. While their paper is cited in this manuscript (11), a more detailed description is needed. Their\nfindings, which illustrate the establishment of spiny and soft-rayed fin domains in acanthomorph fishes,\nprovide essential molecular insights into the differences between spiny and soft-rayed fins.\nResponse 1-5:\nWe agree that the study by Hoch et al. (2021) provides important molecular insights into the distinction\nbetween spiny- and soft-rayed fins. In the revised manuscript, we have expanded our description of this\nstudy to clarify how their findings relate to our work, as shown below:\nLine 416-424, “A previous study reported that distinct transcription factor signatures\ncharacterize the spiny- and soft-ray domains11. The spiny-ray domain specifically expresses\nalx4a/b and tbx2b, whereas the soft-ray domain expresses hoxa13a/b. Our study revealed\ndifferences in developmental mechanisms between spiny and soft rays at both cellular and ECM\nlevels. These differences may be regulated by transcription factors that exhibit mutually exclusive\nexpression patterns in the two domains. Consistent with this idea, ablation of hoxa13a/hoxd13a-\nexpressing fin fold mesenchymal cells reduced the formation of actinotrichia. Taking these\nfindings together with their expression specificities, alx4a/b and tbx2b may promote the\nmesenchymal cell aggregation characteristics of spiny rays, whereas hoxa13a/b may enhance the\nactinotrichia synthesis essential for soft rays.”\nReviewer comment 1-6:\nIn Figure 6, the Runx2/pSmad signals largely overlap in the regions surrounding the tips of the laterally\nprotruding spiny ray in filefish. In contrast, Figures 5a and 5b, which utilize Melanotaenia praecox,\nshow less overlap in the Runx2/pSmad signals, particularly at the tips of the spiny fins. Could this\ndiscrepancy be attributed to the presentation of optical sections? The images should be updated to\ndisplay the z-stack images to clearly show the distributions of the pSmad signals.\nResponse 1-6:\nWe sincerely apologize for our incomplete explanation, which led to this confusion. You have raised\nan important point. The colocalization patterns do indeed appear to be different between the two species.\nRegarding the technical suggestion to use z-stack images, we apologize for not clarifying our\nmethodology. All images in Figure 5a and the right-hand three columns of Figure 6c are optical section\nimages. For the M. praecox samples, optical sections were captured using a pinhole size of 200 µm,\nwhereas for the S. cirrhifer samples the pinhole was adjusted to 202.1 µm. In confocal microscopy, the\npinhole is the key component that gives the system its ability to produce thin optical sections. The\ndifference in pinhole size between the images of the two species is minimal, and it does not affect the\ndegree of overlap between Runx2 and pSmad signals.\nWe intentionally presented optical sections because they are the most rigorous and accurate method for\nassessing true colocalization within individual cells. As you no doubt know, z-stack imaging can create\nfalse signal overlaps from cells at different depths because of the collapsing of the 3D data. This would\nmake it more difficult to determine the true colocalization status of Runx2 and pSmad within individual\ncells. Therefore, we believe the optical sections that we present are the most scientifically accurate data\nfor this analysis.\nAs the reviewer pointed out, the colocalization patterns of Runx2 and pSmad differ between the lateral\nprotrusions of the dorsal spine in S. cirrhifer and the tips of the spiny rays in M. praecox. We are very\ngrateful to you for highlighting this important finding. We interpret these differences as reflecting\ngenuine variation in BMP signaling intensity, which may in turn contribute to the distinct\nmorphogenetic processes in these structures. To make this biological difference clearer (as requested),\nwe have added arrowheads in Figures 5 and 6. We have also added this important comparative point to\nthe Discussion, because we agree that the differences reflect genuine variation in BMP signaling\nintensity, which may in turn contribute to the distinct morphogenetic processes. We have revised the\nmanuscript to include this explanation.\nLine 462-467, “Our pharmacological inhibition assay in M. praecox demonstrated that BMP\nsignaling in spiny rays regulates the thickness of the accumulating bony matrix (Fig. 5b). In\naddition, the colocalization patterns of Runx2 and pSmad1/5/9 differed between the spiny rays of\nM. praecox and the thorny spine of S. cirrhifer. Together, these data suggest that in the thorny\nspine of S. cirrhifer, position-dependent modifications in BMP signaling intensity may regulate\nthorn formation.”\nReviewer comment 1-7:\nHow many larvae of Stephanolepis cirrhifer were examined using immunostaining? The staining results\nfrom other individuals should be presented as supplementary data to confirm reproducibility.\nResponse 1-7:\nWe sincerely apologize for our oversight regarding the sample size of the immunostaining data using\nStephanolepis cirrhifer larvae. We have added the sample size at Line 356 and immunostaining images\nfrom other individuals in Extended Data Figure 7.\nExtended data figure 7. Premature osteoblasts with BMP signaling are spatially modified from\nthe tip to the lateral protrusions of the first dorsal-fin spiny ray in Stephanolepis cirrhifer.\na, b Distributions of Runx2-positive cells (green) and pSmad1/5/9-positive cells (magenta) in the\ndorsal spine of two S. cirrhifer specimens, stained by immunohistochemistry with DAPI (white).\nYellow arrow heads show examples of pSmad1/5/9 and Runx2-positive cells. Dorsal-spine bones\nare outlined in orange. The left-most picture is a confocal Z-stack image, and the others are\noptical sections obtained by confocal microscopy.\nReviewer comment 1-8:\nRegarding the evolution of spiny rays, do the authors think they evolved from soft rays by losing\nactinotrichia, or were they acquired independently?\nResponse 1-8:\nThank you very much for this comment. To answer this question, we have added text to the Discussion,\nas below.\nLine 435-442, “Building on previous studies, our findings allow us to speculate on the evolutionary\norigin of spiny rays. One simple hypothesis is that spiny rays evolved from soft rays through the\nloss of their actinotrichia in the anterior part of each fin. However, the soft-ray morphology of\nthe and1⁻/⁻; and2⁺/⁻ mutants did not resemble that of spiny rays because the soft rays lacked their\nstraight morphology. Therefore, this hypothesis is not supported by the data. To definitively test\nthe evolutionary hypothesis regarding the evolutionary acquisition of spiny rays, further studies\nare required on developmental mechanisms in species belonging to lineages of the teleost\nphylogeny that diverged earlier than the Acanthomorpha did.”\n(Minor points)\nReviewer comment 1-9:\nIn Fig. 1a, spiny and soft rays stained with Alizarin red are shown. Please specify which fin rays belong\nto which fish.\nResponse 1-9:\nAs suggested, we have added information about the fin-ray samples stained with alizarin red in the\nlegend of Figure 1, as below.\nLine 111-112, “The spiny- and soft-ray samples stained with alizarin red are the dorsal fins of\nMelanotaenia praecox.”\nReviewer comment 1-10:\nLines 530-532: Was 5 µl injected into each embryo? That seems like a large volume; please confirm.\nResponse 1-10:\nThank you for pointing this out. We have corrected the error below:\nLine 648-650, “We prepared a 5-µl solution containing 100 ng of each sgRNA, 1,250 ng of Cas9\nprotein, and 0.5 µl of phenol red in nuclease-free water, and then 2–3 nL of this solution were\ninjected into the single cell of the embryo.”\nReviewer comment 1-11:\nFigure 5b: Are the green signals on several dots non-specific? Please clarify this point.\nResponse 1-11:\nWe apologize for being unclear on this part. We added an explanation of these dots as below:\nLine 329-332, “In the immunohistochemistry images for pSmad1/5/9 (leftmost panels of Fig. 5b),\nseveral punctate signals are visible in both control and DMH1-treated fish. In previous studies\nusing the same antibody, pSmad1/5/9 immunoreactivity was prominent in nuclei58,59. Therefore,\nthe punctate signals observed in our images may represent non-specific staining.”\nReviewer comment 1-12:\nLine 210: While both DAFFM and DAR4M are claimed to detect actinotrichia, I am unclear about the\ndifferences between them.\nResponse 1-12:\nWe apologize for the poor explanation. We added a brief explanation of the difference of these two\nsmall molecules as below.\nLine 244-247, “DAR-4M is a small fluorescent molecule similar to DAF-FM that is widely used\nfor NO detection; however, the two molecules exhibit different fluorescence emission\nwavelengths32,45. Recent studies have shown that DAR-4M can bind to collagenous structures such\nas actinotrichia32,47.”\nReviewer #2 (Remarks to the Author):\nReviewer comment 2-1:\nThe manuscript entitled “Actinotrichia-independent developmental mechanisms of spiny rays facilitate\nthe morphological diversification of Acanthomorpha fish fin”, submitted to Nature Communications,\ninvestigates developmental differences between spiny rays and soft rays in rainbowfish (Melanotaenia\npreacox), with additional comparative studies from the highly modified spiny rays of the filefish\n(Stephanolepis cirrhifer).\nTo address their objectives, the authors employ a comprehensive suite of techniques, including micro-\nCT, electron microscopy, immunofluorescence, and in-situ hybridization, to demonstrate key\ndistinctions in fin ray development and anatomy at different scales (from the entire spine to the cellular\nlevel). Specifically, they show that spiny rays are formed by an accumulation of mesenchymal cells\nwhere BMP signaling plays an important role, which differs from soft rays that rely on actinotrichia as\na structural scaffold. Furthermore, the authors identify similar spiny-ray developmental mechanisms\nin the filefish, suggesting that this actinotrichia-independent mode of growth may have facilitated the\nmorphological diversification of fin rays within Acanthomorpha.\nOverall, the manuscript is well-executed and clearly written, with well-organized flow of ideas. The\nexperimental design is robust to address the question proposed (although see general comments about\nsample size and sexes used), and the integration of the multiple methods provides a holistic and\nconvincing analysis. The figures clearly show the results and are easy to follow. The research question\nis original, particularly because little is known about the skeletal development of acanthomorphs.\nTherefore, this research is of interests across multiple fields, including fish biology, physiology, skeletal\ndevelopment, and evolution. However, I suggest the authors to revise my general comments regarding\nthis topic.\nMy main concern is the lack of discussion and consideration of the bone type the authors are studying\n(acellular bone) as well as comparing to (cellular bone), which I consider that is crucial to understand\nthe results and do an accurate comparative analysis. I hope my review will help further improve the\nclarity and impact of this work.\nResponse 2-1:\nWe fully appreciate these comments. We have made revisions to the manuscript in line with your\ncomments and suggestions.\nGeneral comments\nReviewer comment 2-2:\nAcellular bone skeletogenesis.\nOne aspect that stood out to me is that the authors do not mention, or consider, the type of bone that\nforms the skeleton of both species of fishes that are studied here. The majority of Neoteleostei, and thus\nacanthomorps (except some species like tunas) have acellular bone (i.e. a bone without osteocytes).\nThis type of bone is also found in some Euteleostei. In contrast, more basal groups of fishes are\ncharacterized by cellular bone (i.e. a bone with osteocytes), including Ostariophysi where\ncypriniformes (zebrafishes) and siluriformes (catfishes) are part of.\nThis difference in bone type could be one of the reasons why the development of spiny rays in zebrafish\nand catfish (species that authors use to compare their results) may differ from that in the species studied\nhere. Therefore, the spiny ray development described in the manuscript may probably be related with\nthe periosteal osteogenesis of acellular bone. Check also the anatomical terminology used (e.g.\nbasement membranes), as some of the structures may have been already described in Moss (1961). This\npoint can be a very interesting topic to include and discuss, since the formation of acellular bone has\nnot been largely studied.\nFor these reasons, I suggest that the authors consider and discuss the acellular bone formation in the\nmanuscript, as well as to be cautions in describing this developmental process as “unique” to\nAcanthomorph spiny rays, as what the authors may be describing reflect general features of acellular\nbone skeletogenesis rather than a distinct or novel pathway of spiny rays. Additionally, the importance\nof BMP signaling, while certainly relevant, is not surprising as this superfamily of proteins is well\nknown to have a crucial role during bone development across vertebrates.\nBelow I included several papers that discuss the differences between acellular or cellular bone that the\nauthors may find useful.\nReferences\n- Davesne, D., Meunier, F. J., Schmitt, A. D., Friedman, M., Otero, O., & Benson, R. B. (2019). The\nphylogenetic origin and evolution of acellular bone in teleost fishes: insights into osteocyte function in\nbone metabolism. Biological Reviews, 94(4), 1338-1363.\n- Shahar, R., & Dean, M. N. (2013). The enigmas of bone without osteocytes. BoneKEy reports, 2, 343.\n- Moss, M. L. (1961). Osteogenesis of acellular teleost fish bone. American Journal of Anatomy, 108(1),\n99-109.\n- Weiss, R. E., & Watabe, N. (1979). Studies on the biology of fish bone. III. Ultrastructure of\nosteogenesis and resorption in osteocytic (cellular) and anosteocytic (acellular) bones. Calcified tissue\ninternational, 28(1), 43-56.\n- Moss, M. L. (1963). The biology of acellular teleost bone. Annals of the New York Academy of Sciences,\n109(1), 337-350.\n- Dean, M. N., & Shahar, R. (2012). The structure‐mechanics relationship and the response to load of\nthe acellular bone of neoteleost fish: a review. Journal of Applied Ichthyology, 28(3), 320-329.\nResponse 2-2:\nWe thank you for raising this very interesting and crucial point. Your comment has prompted us to\nclarify our argument regarding the evolution of spiny rays and the context of acellular bone.\nThe acquisition of spiny rays in Cypriniformes and Siluriformes is likely independent of that in\nAcanthomorpha. Hoch et al. (2021) showed that the early developmental phase of spiny rays in African\ncatfish resembles that of typical soft rays. In addition, Kubicek et al. (2019) demonstrated that soft rays\npossess actinotrichia at their distal tips. These observations suggest that the spiny rays found in non-\nacanthomorph fishes are homologous to soft rays, rather than to the acanthomorph spiny rays. For this\nreason, we consider that the differences in spiny-ray morphology between acanthomorph and non-\nacanthomorph fishes are primarily caused by differences in osteoblast distribution and physical\nconstraints associated with the presence or absence of actinotrichia.\nWe agree with your point about acellular bone being a critical distinction. However, we propose that\nthe acellular bone type alone cannot explain the developmental differences we observed, with this being\nbased on two key lines of evidence. First, the phylogenetic timing of the bone type transition is\naccording to the emergence of the Euteleost, and this shows a mismatch with the timing of the\nemergence of spiny rays. Second, our study's primary comparison is between spiny rays and soft rays\nwithin the same species. In these fish, both structures are acellular, meaning the bone type itself cannot\nbe the variable that explains the difference in their developmental mechanisms (i.e., actinotrichia-\ndependence vs. independence).\nWe agree with your hypothesis that the evolutionary acquisition of acellular bone may have facilitated\nthe subsequent emergence of acanthomorph spiny rays. This is an interesting point that enriches our\ndiscussion. Following your suggestion, we have now added this important speculation to the manuscript,\nas shown below:\nLine 442-449, “Furthermore, the evolution of bone types might have contributed to the\nacquisition of certain features of spiny rays. The majority of Euteleostei, including\nAcanthomorpha (except for some species such as tunas), possess acellular bone, which lacks\nosteocytes60. In contrast, more basal groups of fishes are characterized by cellular bone, which\ncontains osteocytes60. Although the phylogenetic timing of the transition from cellular to acellular\nbone does not coincide with the emergence of spiny rays, the prior establishment of acellular bone\nmay have facilitated the acquisition of features such as the fusion of the two hemirays and the stiff\nstructure of spiny rays.”\nIn light of your comments, we have carefully reviewed the anatomical terminology in our manuscript.\nWe appreciate this prompt to ensure precision, and although we have confirmed our existing\nterminology is appropriate, we are grateful for the advice.\nWe agree that “unique” was a strong term because some aspects of spiny-ray osteogenesis may be\nshared with other types of bones. We have removed this word and revised the sentence as follows:\nLine 87-88, “Based on our observations, we propose that the developmental mechanisms of spiny\nrays may have facilitated their morphological diversification.”\nReviewer comment 2-3:\nSample size.\nThe authors specify sizes in each section of the Results, but not in Methods. I recommend including\na detailed breakdown in the Methods section, indicating the number of animals used at each\ndevelopmental stage and for each technique (e.g. How many S. cirrhifer and M. praecox where µCT-\nscanned?, this is not specifically clarified anywhere). Additionally, in some instances, it is unclear\nwhich ontogenetic stage is used for a given specific technique (larvae, juveniles, adults?). Lastly, in\nsome cases it is unknown whether the reported sample size in Results refers to the number of rays\nexamined or the number of individual animals. Since multiple rays can be studied from a single\nspecimen, it would be helpful to clarify the number of animals studied similar to how it is done in L130.\nResponse 2-3:\nThank you for this important comment. We apologize for the lack of detail in the Methods section. To\naddress these concerns, we have now included a detailed breakdown of the number of individuals used\nfor each technique in the Methods section. Furthermore, we have clarified that all reported sample sizes\n(n) refer to the number of individuals, and we now explicitly describe the developmental stages of the\nsamples in both the Results and Methods sections.\nIn the original manuscript, we used one individual each of S. cirrhifer and M. praecox for the µCT\nscanning. To increase the robustness of our data, we have performed additional µCT scans on both S.\ncirrhifer and M. praecox individuals. The corresponding data and additional methodological details\nhave been added to the manuscript. Additionally, we have made the CT data publicly available on\nfigshare.\nReviewer comment 2-4:\nAnimal sex.\nThe authors do not specify whether the specimens used were males or females. Several teleosts,\nincluding M. preacox, are sexual dimorphic, with males exhibiting enlarged caudal spines.\nResponse 2-4:\nYou have raised important points. For M. praecox, we did not distinguish the sex for most experiments\n(only for µCT scanning of adults) because the sex of larval and juvenile M. praecox cannot be\ndistinguished; the sex determination mechanisms remain unclear for this species. Consequently, it is\nnot currently feasible to distinguish the sex of our experimental samples using PCR-based methods or\nother investigations. To address the potential issue you have raised, we conducted an additional\nobservation on young adult M. praecox (Extended Data Figure 8), whose sexes are distinguishable by\ntheir body coloration. We found no significant differences in fin bone morphologies between males and\nfemales of this stage. This result indicates that skeletal sexual dimorphism in this species likely appears\nafter sexual maturation. Therefore, we concluded that it was not necessary to distinguish the sex of the\nlarval and juvenile specimens used in our experiments.\nRegarding S. cirrhifer, although sex chromosome system and sexual dimorphisms in the adult second\ndorsal fin have been reported in Murofushi et al. (1980) and Fujita (1955), we did not distinguish the\nsex of our specimens. According to Kwon et al. (2021), sexual maturation in this species occurs at a\ntotal length of approximately 11.7 cm. The specimens used for experiments (0.9–1.4 cm Standard\nLength) were all substantially smaller than this size, and thus would not be expected to have developed\nsexual dimorphisms. Furthermore, sexual dimorphisms of the dorsal spine have not been reported.\nTaken together, we concluded that distinguishing the sex of our S. cirrhifer specimens was unnecessary.\nTo clarify our rationale for not distinguishing sexes in the manuscript, we have added the\naforementioned explanations to the Methods section and new supporting data as an extended figure\n(Extended Figure 5).\nLine 523-529, “When observing the dorsal fin bone morphologies of young male and female fish\n(male, n = 3; female, n = 3), whose sexes are distinguishable by their body coloration36, we did not\nfind any significant differences in the fin bone morphologies between males and females at this\nstage (Extended Data Fig. 5). Furthermore, we did not have a reliable technique to distinguish the\nsex of larval and juvenile M. praecox using PCR-based methods or other investigations, and thus\ndid not distinguish the sex of the larval and juvenile specimens in our experiments.”\nLine 531-539: “Although sex chromosome system and sexual dimorphisms in the adult second\ndorsal fin have been reported in this species66,67, we did not distinguish the sex of our specimens\nbecause a reliable PCR-based genotyping method has not yet been established. According to\nKwon et al. (2021)68, sexual maturation in this species occurs at a total length of approximately\n11.7 cm. The specimens used in our experiments were all significantly smaller than this size and\nwould thus not have been expected to have developed sexual dimorphisms. Furthermore, sexual\ndimorphisms of the dorsal spine have not been reported. Taken together, we concluded that\ndistinguishing the sex of our S. chirrifer specimens was unnecessary.”\nExtended Data Figure 8. Fin bone morphology in the dorsal fin of male and female Melanotaenia\npraecox.\nCalcein staining in young adult M. praecox showing bone morphology in the dorsal fins. Yellow\nasterisks indicate the tungsten needles that were used to spread their dorsal fins.\nSpecific comments:\nAbstract\nReviewer comment 2-5:\n- L22. The hyphenation of “spiny rays” and “soft rays” varies throughout the manuscript. In some\ninstances, the terms are hyphenated (e.g., lines 23, 24, 27), while in others they are not (e.g., lines 22,\n48, 49, 56). Please check for consistency and revise accordingly.\nResponse 2-5:\nThank you for pointing out the inconsistency in the hyphenation of “spiny ray” and “soft ray.”\nWe have carefully revised the manuscript to ensure consistency. Specifically, we use “spiny-ray” and\n“soft-ray” when these terms modify another noun (e.g., spiny-ray development), and “spiny ray” and\n“soft ray” otherwise (e.g., the spiny rays). This usage conforms to standard English grammar\nconventions for compound adjectives.\nReviewer comment 2-6:\n- L22. “their bone structure” is a vague descriptor in this context. What about “their supporting skeletal\nstructure”.\nResponse 2-6:\nWe revised the manuscript according to your suggestions.\nReviewer comment 2-7:\n- L30. “the lateral protrusions equip the BMP positive osteoblast condensation…”. The verb “equip”\nhere is a bit confusing. What about: “the lateral protrusions are associated with BMP-positive\nosteoblast condensation”\nResponse 2-7:\nWe revised the manuscript according to your suggestions.\nReviewer comment 2-8:\n- L36. “… ECM usage would be major factors driving the morphological diversification in animals”. I\nrecommend focusing this sentence specifically on acanthomorphs, rather than using the broader term\n“animals”.\nResponse 2-8:\nWe apologize for the insufficient explanation. We hypothesize that the evolutionary mechanisms\nunderlying diversification of the acanthomorph spiny rays may represent a general process occurring in\nvarious cases of animal evolution. Thus, we suggest that cell distribution and ECM usage could be\nregarded as factors driving morphological diversification not only in Acanthomorpha, but also across a\nbroad range of animal taxa. However, this hypothesis has not yet been tested, and direct evidence\nsupporting this possibility is still lacking. We have revised the manuscript to take these points into\naccount.\nLine 37-39, “This suggests that variation in cell distribution and ECM usage may be important\ncontributors to morphological diversification, not only in Acanthomorpha, but also in other\nanimal taxa.”\nTo clarify our suggestion, we have also revised text in the Discussion section.\nLine 501-505, “We suggest that, in addition to genome duplications61, acquisition of regulatory\ncis-elements62, and the innovation of cell types63 and molecular interactions64, variation in cell\ndistribution and the usage of ECM, such as in the actinotrichia in our fish models, may also be\nmajor factors determining the extent of morphological diversification in a broad range of animal\nevolution.”\nIntroduction\nReviewer comment 2-9:\n- L48. “…and mostly branching,…”. Maybe replace “mostly branching” to “typically branched”.\nResponse 2-9:\nWe revised the manuscript according to your suggestions.\nReviewer comment 2-10:\n- L54 – 55. Are the sucking disc of remoras or the illicium considered “spiny rays”? or they have been\nso modified that they are no longer considered as such?\nResponse 2-10:\nWe apologize for the unclear explanation. The sucking discs of remoras and the illicium are considered\nto be homologous structures to spiny rays. We refrain from proposing a single definitive name here\nbecause various naming schemes are possible and each emphasizes different aspects. Consistent with\nour original manuscript, we avoid making a determination about whether these structures should be\nregarded as spiny rays.\nReviewer comment 2-11:\n- L72. “…there anal fin.” There is a typo here, it should be “their anal fin”.\nResponse 2-11:\nAs suggested, we corrected this error.\nReviewer comment 2-12:\n- L78. “…with that of soft-ray development.” Those of soft-ray development instead, to be consistent\nwith the plural subject (“matrix and cell dynamics”).\nResponse 2-12:\nAs suggested, we corrected this error.\nResults\nReviewer comment 2-13:\n- L92 – 94. This sentence reads awkwardly and may be either missing some information or be poorly\nstructured. I recommend revising it for clarity, as its current form makes the intended meaning difficult\nto follow.\nResponse 2-13:\nWe appreciate your suggestion. The sentence has been revised for clarity and readability. The revised\nversion now reads as follows:\nLine93-94, “To identify differences in morphogenesis between soft and spiny rays, we first\nvisualized their developmental changes using two different staining methods12.”\nReviewer comment 2-14:\n- L114, Figure 1. It is “yellow” instead of “white”.\nResponse 2-14:\nAs suggested, we corrected this error.\nReviewer comment 2-15:\n- L128. “DAFFM is a small fluorescent molecule that broadly used for..”. Instead: “DAFFM is a small\nfluorescent molecule that has been broadly used for…”.\nResponse 2-15:\nWe revised the manuscript according to your suggestions.\nReviewer comment 2-16:\n- L131. Delete “obviously”.\nResponse 2-16:\nWe revised the manuscript according to your suggestions.\nReviewer comment 2-17:\n- L132. Regarding the statement “fine actinotrichia were present at the future position of spiny rays in\nthe median fin fold”. This statement is not clearly supported by the figure. In the image (Fig 2a), the\nactinotrichia do not appear to be specifically localized at the future position of the spiny rays. In fact,\nit doesn’t look like they are found in a specific position. I recommend clarifying the figure or revising\nthe interpretation in the text to better reflect what is shown.\nResponse 2-17:\nWe appreciate your helpful comment. We agree that the actinotrichia are not specifically localized at\nthe future positions of spiny rays. Accordingly, we have revised the sentence to clarify the description\nas follows:\nLine134-136, “Before the emergence of spiny rays (i.e., in DF-st3 larvae), relatively small and less\ndeveloped actinotrichia, compared with those at the tips of the soft rays, were observed in the\nmedian fin fold, including regions that later give rise to the spiny rays.”\nReviewer comment 2-18:\n- L135 – 136. In the spiny and soft rays, the DAFFM only stains actinotrichia? If not, which molecules\nform the brightest bulk of the rays? See image below:\nResponse 2-18:\nWe apologize for the unclear explanation. DAFFM staining visualizes collagenous components,\nincluding both actinotrichia, bone collagen, and tendon of the segments in the soft rays. We have\nclarified this point in the added text as follows:\nLine139-140, “DAFFM staining visualizes collagenous structures, including actinotrichia, bone\ncollagen, and tendon of the segments in the soft rays32,45.”\nReviewer comment 2-19:\n- L209: “n=3”. The number of spiny tails (sample size) sampled are from 3 different animals?\nResponse 2-19:\nWe revised the explanation about sample sizes as you suggested in Reviewer comment 2-3.\nLine242-244, “First, we examined the 3D cell distribution at the tips of spiny rays (DF-st4 larvae,\nn = 3) and soft rays (DF-st4 larvae, n = 3) by staining the nucleus and collagen structures\nsimultaneously.”\nReviewer comment 2-20:\n- L213: “…such condensed mesenchyme cells”. How the authors know that these are mesenchyme cells\nif they did not use any markers, only the staining of the nuclei? If the authors are defining mesenchyme\ncells based on their rounder, larger morphology, they could be also osteoblasts or other precursor cell?\nResponse 2-20:\nWe apologize for the mistake. In this context, our intention was to indicate that the non-epithelial cells\ncondensed at the tip of the spiny rays, but not at the tip of the soft rays. Therefore, we have replaced the\nword “mesenchyme” with “mesenchymal cells” in the relevant sentence.\nReviewer comment 2-21:\n- Figure 3: There is a typo in SYTO13 (nucleus).\nResponse 2-21:\nAs suggested, we corrected this error.\nDiscussion\nReviewer comment 2-22:\n- L334: using the “unique” adjective here is unnecessary. The authors do not know if this mechanism\nis unique to spiny rays or to acanthomorphs, since this process can be happening in other skeletal\nelements or in other clades (perhaps is how the acellular bone is formed?).\nResponse 2-22:\nThank you for this suggestion. We omitted the “unique” adjective and revised the sentence as follows:\nLine391-392, “Our study confirms that some features of the morphogenetic mechanisms\nunderlying spiny-ray development in acanthomorph fish differ from those of soft-ray\ndevelopment.”\nReviewer comment 2-23:\n- L344: Secondarily?\nResponse 2-23:\nWe apologize for the insufficient explanation. We used the word “secondarily” to indicate that the\ndevelopmental mechanisms occurring at the tips of spiny rays were also utilized at the lateral protrusions\nof the dorsal spine in S. cirrhifer. To avoid confusion, we have removed this term and revised the\nmanuscript as follows.\nLine 373-374, “Figure 6. Premature osteoblasts with BMP signaling are spatially modified from\nthe tip to the lateral protrusions of the first dorsal-fin spiny ray in Stephanolepis cirrhifer.”\nLine 400-402, “The acquisition of the thorny spine in S. cirrhifer may have resulted from a spatial\nmodification of an ancestral developmental mechanism that was originally established at the tips\nof spiny rays.”\nLine 494-496, “The pre-established independence from actinotrichia and the autonomous\nosteogenic mechanisms at the tips of spiny rays may have been spatially modified and redeployed\nat alternative positions.”\nReviewer comment 2-24:\n- L361 – L363: The fact that the developmental process that forms the spiny rays described in this\nmanuscript is different of the process of the formation of spiny rays in Siluriformes may be because\ncatfishes have cellular bone (see my comments above).\nResponse 2-24:\nThank you for bringing this to our attention. We have carefully considered this point, and our detailed\nexplanation can be found in Response 2-2.\nIn brief, recent studies (e.g., Hoch et al., 2021) suggest that the “spiny rays” in Siluriformes (catfish)\nare developmentally homologous to soft rays (i.e., they are actinotrichia-dependent). As this point is\nintegral to the main discussion on bone types, our full and detailed explanation can be found in Response\n2-2.\nReviewer comment 2-25:\n- L406 – L411: this text looks like doesn’t belong here?\nResponse 2-25:\nThe text in lines L406–L411 is correctly placed. We apologize for the confusion caused by a simple\nformatting error that resulted in this section being displayed in a smaller font size.\nMethods\nReviewer comment 2-26:\n- Add sample sizes for each technique (see my comment above).\nResponse 2-26:\nAs suggested, We revised the manuscript according to your suggestions.\nReviewer comment 2-27:\n- L445: after fixation, where the samples directly transferred to 70% ETOH? Or the authors dehydrated\nthem using increasing concentration steps?\nResponse 2-27:\nWe apologize for our unclear explanation. We revised the methodology as follows:\nLine 554-557, “Samples of M. praecox adults were fixed with 10% formaldehyde at room\ntemperature overnight, followed by dehydration using an ethanol series (50% EtOH, 70% EtOH,\n80% EtOH, 90% EtOH, 95%EtOH, 100% EtOH, 95% EtOH, 90%EtOH, 80%EtOH,\n70%EtOH). These samples were then stored in 70% ethanol.”\nReference\n- Höch, R., Schneider, R. F., Kickuth, A., Meyer, A., & Woltering, J. M. (2021). Spiny and soft-rayed\nfin domains in acanthomorph fish are established through a BMP-gremlin-shh signaling network.\nProceedings of the National Academy of Sciences of the United States of America, 118(29).\n- Kubicek, K. M., Britz, R., & Conway, K. W. (2019). Ontogeny of the catfish pectoral-fin spine\n(Teleostei: Siluriformes). Journal of Morphology, 280(3), 339–359.\n-Murofushi, M., Oikawa, S. Nishikawa, S. Yoshida, T. H. (1980) Cytogenetical Studies on Fishes, III.\nMultiple Sex Chromosome Mechanism in the Filefish, Stephanolepis cirrhifer, The Japanese Journal\nof Genetics, 1980, Vol. 55, No. 2, 127-131,\n- Fujita, S. (1955) On the development and prelarval stages of the file-fish, Monacanthus cirrhifer\nTemminck et Schlegel. Sci. Bull. Fac. Agr., Kyushu Univ., 15: 229-234. (In Japanese with English\nsummary.)\n- Kwon, H. C., Lee, J. B., Zhang, C. I., Lee, D. W., Choi, Y. M. (2011) Maturation and Spawning of\nFilefish, Stephanolepis cirrhifer in the East Sea of Korea. KOREAN JOURNAL OF ICHTHYOLOGY,\nVol. 23, No. 2, 111-118."}, {"round": 2, "reviewer_comments": {"reviewer_1": "The authors have responded to my comments thoroughly and satisfactorily. This is an excellent study that deserves publication in Nature Communications, and I commend the authors for their sustained efforts over the past several months.", "reviewer_2": "This is the second time that I have reviewed the manuscript entitled “Actinotrichia-independent developmental mechanisms of spiny rays facilitate the morphological diversification of Acanthomorpha fish fins”, submitted to Nature Communications.\nThe authors have adequately addressed my previous comments. In particular, they have responded to my suggestions regarding the discussion of cellular versus acellular bone, and they have clarified the sex of the specimens used whenever possible.\n\nOverall, I think the authors present a robust and carefully conducted study. This work offers valuable insights into the developmental mechanisms of spiny and soft rays which, in case of spiny rays, likely contribute to the highly diversified morphologies observed in some acanthomorph species.\n\nI have noted a few minor grammatical errors throughout the manuscript, but aside from these minor issues, the manuscript reads well and is scientifically sound.\n- L50: “Branched” instead of “branching”\n- L248: “We found that mesenchymal cells condensed at the tips of spiny rays, whereas such condensed mesenchymal cells were not detected at the tips of soft rays (Fig. 3a)”. Personally, I don’t see the condensation of mesenchymal cells in Figure 3a.\n- L254: “Zebrafish” instead of “Zebrafiah”\n- L280: “a Cell, ECM, and bone condition”. The ECM is not particularly visible in a.\n- Small yellor and pink numbers, present in figure 3 and Extended Data figure 6, that indicate regions located at the top-left corner. These are very difficult to see, maybe adding a black background may help.\n- L360: “Larva” is singular, therefore it sould be “has”.\n- L411: “Alternative to” instead of “Alternative of\""}, "author_response": "Point-by-point responses\nWe thank you very much for reviewing our manuscript. We have revised it following careful\nexamination of your suggestions. Your comments helped us to greatly improve the quality of our paper,\nand we express our gratitude to both of you for your constructive suggestions.\nReviewer #2 (Remarks to the Author):\nReviewer comment 2-1:\nThis is the second time that I have reviewed the manuscript entitled “Actinotrichia-independent\ndevelopmental mechanisms of spiny rays facilitate the morphological diversification of Acanthomorpha\nfish fins”, submitted to Nature Communications.\nThe authors have adequately addressed my previous comments. In particular, they have responded to\nmy suggestions regarding the discussion of cellular versus acellular bone, and they have clarified the\nsex of the specimens used whenever possible.\nOverall, I think the authors present a robust and carefully conducted study. This work offers valuable\ninsights into the developmental mechanisms of spiny and soft rays which, in case of spiny rays, likely\ncontribute to the highly diversified morphologies observed in some acanthomorph species.\nI have noted a few minor grammatical errors throughout the manuscript, but aside from these minor\nissues, the manuscript reads well and is scientifically sound.\nResponse 2-1:\nWe thank the reviewer for their positive evaluation of our revised manuscript and for taking the time to\nre-review it. We are pleased that our responses to the previous comments were found to be adequate.\nWe have carefully corrected the minor grammatical errors noted by the reviewer. We sincerely\nappreciate the reviewer’s constructive feedback and support.\nReviewer comment 2-2:\n- L50: “Branched” instead of “branching”\nResponse 2-2:\nAs suggested, we corrected this error.\nReviewer comment 2-3:\n- L248: “We found that mesenchymal cells condensed at the tips of spiny rays, whereas such condensed\nmesenchymal cells were not detected at the tips of soft rays (Fig. 3a)”. Personally, I don’t see the\ncondensation of mesenchymal cells in Figure 3a.\nResponse 2-3:\nWe appreciate your helpful comment. Accordingly, we have revised the sentence to clarify the\ndescription as follows:\nLine247-249, “We found that mesenchymal cells are multilayered at the tips of spiny rays (Fig.\n3a-left), whereas such condensed mesenchymal cells were not detected at the tips of soft rays (Fig.\n3a-right).”\nReviewer comment 2-4:\n- L254: “Zebrafish” instead of “Zebrafiah”\nResponse 2-4:\nAs suggested, we corrected this error.\nReviewer comment 2-5:\n- L280: “a Cell, ECM, and bone condition”. The ECM is not particularly visible in a.\nResponse 2-5:\nWe apologize for the unclear explanation. DAR4M staining visualizes only collagenous components.\nThen, we have clarified this point in the added text as follows:\nLine280, “a Cell, collagen, and bone conditions of the dorsal fin in a larva at DFst-4.”\nReviewer comment 2-6:\n- Small yellor and pink numbers, present in figure 3 and Extended Data figure 6, that indicate regions\nlocated at the top-left corner. These are very difficult to see, maybe adding a black background may\nhelp.\nResponse 2-6:\nWe revised the figures according to your suggestions.\nFigure3\nExtended Figure6\nReviewer comment 2-7:\n- L360: “Larva” is singular, therefore it sould be “has”.\nResponse 2-7:\nAs suggested, we corrected this error.\nReviewer comment 2-8:\n- L411: “Alternative to” instead of “Alternative of\"\nResponse 2-8:\nAs suggested, we corrected this error."}]

Decision letter