Sexually dimorphic (SD) traits are important in sexual selection and species survival, yet the molecular basis remains elusive, especially in amphibians where SD traits have evolved repeatedly. We focus on the Leishan moustache toad (Leptobrachium leishanense), in which males develop nuptial spines on their maxillary skin. Here we report a 3.5 Gb genome assembly with a contig N50 of 1.93 Mb. We find a specific expansion of the intermediate filament gene family including numerous keratin genes. Within these genes, a cluster of duplicated hair keratin genes exhibits male-biased and maxillary skin-specific expression, suggesting a role in developing nuptial spines. We identify a module of coexpressed genes significantly associated with spine formation. In addition, we find several hormones likely to be involved in regulating spine development. This study not only presents a high-quality anuran genome but also provides a reference for studying skin-derived SD traits in amphibians.
Males and females of a species often exhibit different strategies to maximize fitness. This difference in optimal strategies can select for phenotypic differences between two sexes1. Sexually dimorphic (SD) modifications include male-specific morphology, such as the lion’s mane2, the sword-like tail of swordtail fishes3, and the greatly enlarged tooth of male narwhals (Monodon monoceros)4. As most of the genome is shared between sexes with the exception of genes in sex chromosomes, sexual dimorphism is primarily caused by differential, sex-biased gene expression1. To achieve sex-biased expression, gene duplication at the genomic level is a potential solution. Redundant paralogs generated by duplication events can allow one or more genes to diverge in expression levels and/or protein structure5. A wealth of evidence has proven that duplicated genes often acquire male-biased and tissue-specific expression6. Recent studies exploring genome-wide sex-biased expression patterns in different species revealed a broad variation in the percentage of sex-biased genes (ranging from 2% of transcripts in Littorina saxatilis to 90% in Drosophila melanogaster)7. Although these studies provide insights into sex-biased expression patterns, very few of them link sex-biased genes to particular traits. Furthermore, most of conclusions regarding the molecular regulation of SD traits come from studies on model species8. The main obstacle linking interesting traits to sex-biased genes outside of model organisms is that we often lack genomic information for nonmodel species, including genomic resource and knowledge of genetic networks governing trait development.
Amphibians are a group of vertebrates with abundant SD traits. In anurans (toads and frogs), >90% of the species exhibit larger body sizes in females than in males9. Another type of SD trait in anurans is skin-derived excrescences, such as the nuptial pads on the digits of hands or on the ventral surfaces of forelimbs10 and the nuptial spines on the upper jaw11. These traits are present mainly in adult males during the breeding period and exhibit a seasonal cycle. A morphological survey on nuptial pads from 26 species in Phyllomedusinae (Hylidae) found that the pads consisted of dark epidermal projections (EPs). The shape and density of EPs and the separation level between adjacent EPs differ among species12. Hormone implant experiments revealed that the pads could be induced by androgens in adult males, adult females, and even tadpoles13. However, in contrast to the results of morphological examination and hormone manipulation, the genetic basis of shaping nuptial excrescences remains elusive, which can be largely attributed to the lack of genomic information. For instance, a recent study on Leptobrachium boringii using nonreference transcriptomic analyses found several processes (such as cytosolic processes and peptidase inhibitor activity) and a list of potential genes (such as insulin-like growth factor genes and sex steroid hormone-related genes) that may be associated with the seasonal development of nuptial spines14. However, owing to the absence of a reference genome, the annotation rate of unigenes was very low (30.98%), indicating that most of the unigenes were unannotated14. This limitation will hinder our understanding of important biological processes and genes associated with SD traits.
The genomes of amphibians are exceptionally large (up to 120 Gb in salamanders) and feature high levels of repetitive sequences15, which makes both sequencing and assembly challenging. To date, six anuran genomes have been sequenced and annotated16,17,18,19,20,21,22, among which only the genomes of Xenopus laevis and Xenopus tropicalis have been assembled to chromosome level17,19. Among the sequenced anurans, X. laevis and X. tropicalis belong to the Archaeobatrachia, while the other four species (Nanorana parkeri, Rana catesbeiana, Rhinella marina, and Oophaga pumilio) belong to Neobatrachia23. However, to the best of our knowledge, no genomic data for the spadefoot toads (Pelobatoidea), a monophyletic clade sister to Neobatrachia relative to the paraphyletic Archaeobatrachia23, has been published. Within the Pelobatoidea, Megophryidae that is the most widely diversified family provides an appropriate system for studying skin-derived SD traits in amphibians. For example, in the Leishan moustache toad (Leptobrachium leishanense), males have skin excrescences that are similar to skin-derived SD traits in other anurans in terms of tissue structure and developmental cycles. During the breeding period, adult males develop four sharp and conical black spines on the maxillary skin (MS) (two on each side), whereas such a structure is absent in females. During the postbreeding period, adult males lose the conical outer casing24. Immature males have only red spots in the same area. It has been proposed that such nuptial spines may be used for male–male combat, stimulation of females, or nest construction/maintenance during the breeding season25.
Herein, to build a representative genome of Pelobatoidea and to understand the molecular basis of skin-derived SD traits in amphibians, we focus on the Leishan moustache toad. We first assemble a 3.5 Gb of chromosome-scale genome. Comparative genomic analysis reveals expansion of intermediate filament (IF) gene families (GFs) in L. leishanense, which include numerous keratin genes (krts). To identify biological processes and genes associated with the production of nuptial spines, we compare the transcriptomes of multiple tissues, including dorsal skin (DS), MS, brain, and gonad, between males and females at three developmental stages. Stage A is the subadult stage, when males have not developed spines and females have no eggs; stage B is the breeding stage, when males have developed spines and females have eggs; stage C is the postbreeding period, when males’ spines fall off and females have laid eggs. We identify a module of coexpressed genes to be significantly associated with spine formation. In addition, hormones such as androgen, thyroid hormone (TH), prolactin (PRL), and relaxin (RLN) are likely involved in the regulation of spine development. In summary, we obtain a high-quality reference genome and reveal a series of candidate genes underlying the production and regulation of nuptial spines in L. leishanense. Similar regulatory patterns can be used to guide future studies on skin-derived SD traits in amphibians.
Genome assembly and characterization
A male L. leishanense toad was selected for genome sequencing and assembly. The genome size was estimated to be 3.56 Gb based on the k-mer distribution (Supplementary Fig. 1). A total of 285.81 Gb (~80×) of PacBio long reads were assembled using Canu v1.526 combined with WTDBG v1.1.006 (https://github.com/ruanjue/wtdbg). This assembly was polished based on 174.86 Gb (~50×) of Illumina paired-end reads using Pilon v1.2227. Subsequently, we used LACHESIS (http://shendurelab.github.io/LACHESIS/) to assemble contigs on 13 pseudochromosomes28 based on 155.16 Gb (~44×) of Hi-C data. A total of 3296 contigs with a length of 3.31 Gb were assembled to chromosome-level scaffolds (Supplementary Fig. 2). We finally generated a 3.54 Gb of L. leishanense genome, with a contig N50 of 1.93 Mb and scaffold N50 of 394.69 Mb, providing the first chromosome-anchored genome among Pelobatoidea species (Table 1, Fig. 1a, Supplementary Tables 1 and 2).
To evaluate the assembly quality, we mapped the Illumina reads on the reference genome. Approximately 95% of the paired-end reads were mapped properly. Then we aligned 258,442 transcriptomic unigenes from nine L. leishanense tissues to the assembly, with >99.6% of unigenes being aligned, indicating excellent coverage of the expressed genes (Supplementary Table 3). Furthermore, we examined the completeness of the conserved core eukaryotic genes (CEGs) and universal single-copy orthologs using CEGMA v2.529 and BUSCO v330 (tetrapoda_odp9 database), respectively. The reference genome of L. leishanense includes 241 of the 248 (97.2%) complete CEGs and 3840 of the 3950 (97.2%) complete and partial BUSCO genes, indicating high completeness of the assembled genome. These metrics are higher than those of sequenced anuran genomes as evaluated by the same method (Supplementary Figs. 3 and 4 and Supplementary Tables 4 and 5).
Genome annotation and chromosome synteny
We annotated the repetitive sequences based on the de novo repeat sequence database of L. leishanense combined with Repbase 20.0131. We found that 77.1% (2.73 Gb) of the L. leishanense genome was repetitive sequences, which is higher than the values for most anurans with sequenced genomes (Table 1). Similar to other species, the most abundant transposable elements (TEs) are terminal inverted repeats (Supplementary Table 6). Notably, two major retrotransposons, long interspersed nuclear element and long terminal repeat (LTR), constitute a higher proportion of the L. leishanense genome (24.8% and 17.5%, respectively) than other anurans, suggesting the high accumulation of these TEs in L. leishanense. Such accumulation of retrotransposons provides the potential for genome size expansion via duplicative transposition. Moreover, the TE expansion pattern of L. leishanense is similar to that of R. catesbeiana and R. marina, which exhibits one major expansion wave followed by a slight burst that occurred recently (Fig. 1c). After masking repetitive sequences, we annotated a total of 23,420 genes in the reference genome with an average gene length of 30 kb (Supplementary Fig. 5), and 96.4% of these genes were annotated in public databases.
To compare structural characteristics of the genomes between L. leishanense and X. tropicalis, we analyzed chromosomal synteny based on genome-scale ortholog alignment. We found extensive chromosome synteny between two species, with a vast majority of orthologs located on one chromosome in X. tropicalis also being located on a single chromosome in L. leishanense (Fig. 1b). In L. leishanense, a total of 346 blocks with an average size of 7.68 Mb were collinear with the X. tropicalis blocks. The chromosome number in L. leishanense is 13, whereas the number in X. tropicalis is 10. This difference is reflected in the corresponding chromosomal fissions in L. leishanense. Specifically, the blocks on the X. tropicalis chromosomes (Xtr4, 7, and 8) are distributed on two separate chromosomes in the L. leishanense genome (Lle8+10, 6+11, and 9+13, respectively). In addition, chromosomal rearrangements such as inversion and translocation were detected. In the L. leishanense genome, a total of 179 out of 346 blocks were inverted (Supplementary Data 1). Seven regions in L. leishanense with sizes of 502 kb–5.93 Mb corresponding to Xtr2, 3, 6, 9, and 10 of X. tropicalis were translocated into other non-homologous chromosomes (Fig. 1b, Supplementary Figs. 6 and 7).
Expanded GFs in L. leishanense
We compared the L. leishanense genome with six other anurans (R. catesbeiana, N. parkeri, R. marina, O. pumilio, X. tropicalis, and X. laevis) and four vertebrates (Mus musculus, Homo sapiens, Anolis carolinensis, and Danio rerio) to analyze anuran divergence. A phylogenetic tree based on single-copy orthologs revealed the middle position of L. leishanense between Neobatrachia and Archaeobatrachia (Fig. 2). The most recent common ancestor (MRCA) among the seven anurans was estimated to occur at 210.06 million years ago (Ma) (176.22–248.83 Ma). L. leishanense shared an MRCA with Neobatrachia species at 185.90 Ma (154.10–221.31 Ma, Fig. 2).
It has been proposed that changes in gene copy number would support adaptive evolution32. We thus estimated the expanded GFs in L. leishanense to explore targets of adaptive evolution. A total of 107 GFs were significantly expanded in L. leishanense compared with its MRCA (Fig. 2). Genes from expanded families were mainly enriched in signal response categories, including sensory perception of smell, response to stimulus, and signal transduction (Supplementary Data 2). These categories were also significantly expanded in other anurans such as N. parkeri, R. catesbeiana, and R. marina (Supplementary Data 2). The PI3K-Akt signaling pathway, which plays important roles in responding to extracellular stimuli and regulating multiple cellular functions, such as cell proliferation, apoptosis, and survival33, was significantly overrepresented in L. leishanense (Fisher’s exact test, Benjamini–Hochberg (BH) corrected p < 0.001). In addition, biological processes associated with immune responses, including the immune system process, antigen processing and presentation, and immune response, were significantly and specifically expanded in L. leishanense (Supplementary Data 2). Further analyses and empirical data are needed to explore the relationship between habitat adaptation and the expansion of immune response-related genes in L. leishanense.
Expanded keratin genes support nuptial spine formation
We found two GFs encoding IF proteins were significantly expanded in L. leishanense (Fig. 2). These GFs include genes encoding keratins that are filament-forming proteins of epithelial cells and are involved in shaping keratinized tissues in vertebrates, such as human hairs and nails34. Considering the hardness of nuptial spines, we hypothesize that the expansion of the IF GFs may be associated with the occurrence of nuptial spines in L. leishanense. We therefore conducted a comprehensive analysis of krts in L. leishanense and compared to other vertebrates.
In tetrapod, an important component of integument rigidity is formed by two classes of α-keratins (termed type I and type II α-keratins)34. Genomic analyses have demonstrated that humans possess a total of 54 functional krts (28 type I and 26 type II), separately clustered on two chromosomes35. In L. leishanense, we identified a total of 101 complete α-krts, including 53 type I and 48 type II genes (Supplementary Data 3), which is much higher than the number observed in other species34 (Supplementary Table 7). By comparing the genomic distribution of α-krts, we found that type I and II α-krts were separately clustered on two chromosomes (Lle 12 and Lle 2) in L. leishanense and were flanked by the same genes as those in other vertebrates, suggesting conserved arrangement order of krts. There are several clusters of lineage-specific paralogous krts in mammals (human and mouse), L. leishanense, and other anurans (Fig. 3a; Supplementary Fig. 8). The most diversified and duplicated krts within L. leishanense are homologous to mammalian hair keratins (HKs). Notably, the copy number of HK genes in L. leishanense is almost double than in other anurans (Fig. 3a).
To further explore the correlation of duplicated HK genes with nuptial spines in L. leishanense, we sequenced the expressed RNAs and compared the expression patterns of HK and non-HK genes in multiple samples (Supplementary Fig. 9). We found that the duplicated HK genes (types I and II) in L. leishanense showed male-biased and MS-specific expression during the breeding stage (B), whereas non-HK genes expressed similarly in both the MS and DS in two sexes (Fig. 3b, Supplementary Data 4). These results support that the expansion of the IF GFs, especially the highly duplicated HK genes, is associated with the occurrence of nuptial spines in L. leishanense.
Biological processes and pathways in producing spines
To investigate biological processes associated with nuptial spines, we sequenced transcriptomes from the MS in two sexes and three developmental stages (Table 2). We also sequenced transcriptomes from the DS as controls for skin. For each stage, we performed three groups of independent comparisons: (1) comparison of MS between males and females; (2) comparison of MS against DS in males; and (3) comparison of MS against DS in females. We identified differentially expressed genes (DEGs) for each comparison and conducted Gene Ontology (GO) and Kyoto Encyclopedia of Genes and Genomes (KEGG) enrichments.
Stage A is the subadult period, when males and females show similar red spots on the MS. In adult males, these spot regions develop black spines during breeding seasons. Thus DEGs in males’ MS at stage A would be associated with early preparation for spine development. Differential expression analyses revealed that only 19 out of the 15,135 genes were expressed differentially between males’ and females’ MS (|log2(fold change)| > 1 and BH corrected p < 0.01) and no GO term was enriched (Fig. 4, Supplementary Table 8), which is consistent with the similar phenotypes between two sexes. DEGs between the MS and DS in males were mainly involved in transcriptional regulation processes (including positive and negative regulation of transcription from the RNA polymerase II promoter) and skeletal development-related processes (including embryonic skeletal system morphogenesis and cartilage development). Similar processes were not enriched in the comparison of females (Fig. 4). Stage B is the adult breeding period, when males have developed spines and females retain red spots in the MS. Thus DEGs in males’ MS at stage B would be directly associated with the formation of nuptial spine. We found that 83 out of 14,409 genes were differentially expressed in the MS between two sexes. These DEGs were enriched in hormone activities (such as TH transport and steroid biosynthetic process) and epithelial cell differentiation (Fig. 4). Similar processes were also enriched for DEGs in males’ MS vs DS but were absent in the same comparison for females (Fig. 4). At the postbreeding stage (C) when males’ spines fall off, we found that the DEGs of males’ MS vs DS were significantly enriched in the proteolytic process that contains multiple genes encoding trypsin and cathepsin (Fig. 4). Proteolysis can break peptide bonds and hydrolyze proteins to small polypeptides (https://www.ebi.ac.uk/QuickGO/term/GO:0006508). It was reported that trypsin could degrade human epidermal keratins in vitro36. We thus consider that the proteolytic process may be involved in triggering the falling off of the spines, probably by degrading the keratin protein complex.
Findings from amphibian endocrine research have revealed that male-biased structures such as enlarged flexor muscles and nuptial thumb pads are androgen dependent13. In amphibians, as in other vertebrates, androgens (testosterone and 5α-dihydrotestosterone) are mainly produced by testes and regulated by gonadotropins released from the pituitary gland37. Therefore, to explore the hormonal regulation of nuptial spines in L. leishanense, we sequenced the transcriptomes from brain and gonads (Table 2). DEGs from brains were significantly enriched in the neuroactive ligand–receptor interaction pathway (BM3 vs BF3 and AM3 vs BM3, Supplementary Fig. 10), which includes numerous signaling molecules, such as hormones and the associated receptors. Within this pathway, five genes encoding hormone subunits were specifically highly expressed in males’ brains at stage B (BM3 in Fig. 5a). Among these genes, the lh (luteinizing hormone) and fsh (follicle-stimulating hormone) encode peptide hormones that stimulate the synthesis of steroid hormones by traveling via the blood to the gonads38. High expression of these genes indicates the activation of the hypothalamus–pituitary–gonad axis in males during the breeding season. Consistently, the steroid hormone biosynthesis pathway was significantly enriched in males’ testes between different stages (AM4 vs BM4 and BM4 vs CM4, Supplementary Figs. 10 and 11). Within this pathway, genes encoding enzymes involved in sex steroid biosynthesis were highly expressed in male testes at stage B (Fig. 5b), suggesting a high level of androgen biosynthesis.
In addition to gonads, the skin is a major nonclassic tissue for steroidogenesis via the expression of functionally active enzymes39. In L. leishanense, DEGs between the male and female MS were also significantly enriched in the steroid biosynthesis process, which includes genes encoding 3-beta-hydroxysteroid dehydrogenase (hsd3b), 17α-hydroxylase (cyp17a1), and steroidogenic acute regulatory protein (star)40. High expression of these enzymes indicates the accumulation of C19 steroids of androgens in males’ MS. Although the level of steroidogenesis in nonclassic tissues is quite modest compared with that in gonads, this process could be very important in locally autocrine or paracrine regulation39. However, androgens alone are insufficient to support male SD traits, and other hormones play critical roles as well13. Here we found that TH, PRL, and RLN may be involved in the regulatory process. The “thyroid hormone transport” term was highly enriched in DEGs in BM2 vs BF2 and BM1 vs BM2 (Fig. 4). Furthermore, genes encoding PRL and RLN were more highly expressed in the male MS than in the female MS based on both RNA-seq and quantitative PCR (qPCR) data (false discovery rate (FDR)prl < 0.001, log2(FC)prl = −8.41; FDRrln < 0.001, log2(FC)rln = −10.59, Supplementary Figs. 12 and 13).
Gene coexpression analyses reveal the genetic networks underlying the spine production
To further identify the key genes controlling the production of nuptial spines, we analyzed the gene coexpression networks and identified genetic modules using weighted gene correlation network analysis (WGCNA)41. We sought to identify modules (groups of highly correlated genes that may exhibit the same biological activity) that were associated with specific traits. Based on the correlation of expression profiles among genes, we identified 12 modules in total, with the 13th module including genes that were not assigned to any module (Fig. 6a; Supplementary Fig. 14; Supplementary Data 5). Then we characterized the gene expression pattern of each module based on the module eigengene (ME), which represents the first principal component of the scaled module expression profiles. MEs of each module were correlated to external sample traits, with significantly correlated modules suggesting specifically high expression patterns for a particular trait. We found that ME03, representing 170 genes, was significantly correlated with MS at stage A in both males and females (rmale = 0.53, p < 0.001; rfemale = 0.32, p = 0.007, Fig. 6a), suggesting that genes in this module are mainly involved in MS specialization prior to spine development. Within this module, positive regulation of the canonical Wnt signaling pathway was significantly enriched (corrected p = 0.012, Supplementary Data 6). In addition, the Wnt signaling pathway (ko04310) was enriched from KEGG analysis (p = 0.02, corrected p = 0.27, Supplementary Data 6). In the canonical Wnt signaling pathway, genes encoding the Wnt protein, two leucine-rich repeat-containing G-protein coupled receptors (LGRs), and Wnt inhibitory factor 1 (Wif1) were included in ME03 (Fig. 6b). The coexpression pattern between the Wnt protein and LGR receptors suggests the activation of Wnt signaling in the MS prior to the formation of nuptial spines. In addition, we analyzed the positively selected genes in L. leishanense compared with six other anurans using the branch-site model of CODEML in PAML v4.942. The gene encoding adenomatous polyposis coli, which facilitates β-catenin degradation43, had experienced significantly positive selection in L. leishanense (p = 0.004; Supplementary Data 7). Signaling by the Wnt proteins is one of the fundamental mechanisms that directs cell proliferation during embryogenesis43. Moreover, the Wnt signaling pathway has been reported to facilitate the formation of skin appendages such as teeth, hair follicles, and deer antlers by mediating epithelial–mesenchymal interactions and determining cell fate44,45. Therefore, we propose that activation of the Wnt signaling pathway is involved in triggering the production of nuptial spines in L. leishanense. This result reveals the co-option of the pre-existing Wnt signaling pathway for generating novel traits.
We also found that module ME05 showed a significant correlation with the male MS at stage B (r = 0.86, p < 0.001, Fig. 6a). This module contains 43 genes, 16 of which are krt genes (Fig. 6c). Within this module, the coexpression network reveals that krt genes showed strong correlation with other genes, supporting the core roles of krt genes in the formation of SD spines. This module also contains three tgl2 genes (encoding protein-glutamine gamma-glutamyltransferase 2) that are associated with cross-linking in the keratin assembly process46. In addition, the hoxc13 gene showed a distinct correlation with other genes, although the edge weight was not as high as that of the krt genes (Fig. 6c). It has been shown that hoxc13 is involved in the regulation of human HK expression47. The gene encoding tyrosinase (tyr), which is the essential enzyme in melanin synthesis48, was also included in this module, which suggests the role of this gene in the maintenance of the black color of the spines. Thus this module provides the most potential candidate genes that are directly associated with the occurrence of nuptial spines in male L. leishanense toads.
In this study, we generated a Pelobatoidea reference genome by combining PacBio long-read sequencing and chromatin interaction scaffolding methods. We detected two recent burst events of TEs in L. leishanense genome, which is similar to the genomes of R. catesbeiana (5.8 Gb), R. marina (2.6 Gb), and O. pumilio (5.5 Gb) from Neobatrachia. In contrast, Xenopus species (sizes in 1.5–2.7 Gb) from Archaeobatrachia only presented one peak. However, the number of protein-coding genes is similar across these species, indicating that different expansion and accumulation of TEs is one of the major factors contributing to the variation in genome size in amphibians. Diverse genome sizes in amphibians may be driven by adaptive and/or nonadaptive forces. On the one hand, the genome size is strongly associated with developmental rates in Anura. Natural selection will reduce the genome size in environments that foster rapid embryonic and larval development49. On the other hand, the evolution of genome size can be affected by neutral factors, such as mutation rates and effective population size49,50. For example, the substitution rates of both mitochondrial and nuclear datasets are positively correlated with genome size in Anura49. Exploring the relative roles of adaptive and neutral factors in driving changes in genome size would be interesting when an increasing number of amphibian genomes have been sequenced.
Although vertebrate genomes differ greatly in DNA content and chromosome karyotype, extensive synteny exists across species51. Here we detected a highly conserved genomic arrangement pattern between L. leishanense and X. tropicalis, despite the different numbers of chromosomes and the long-term divergence (~210.1 Ma) between two species. Similar genomic synteny analysis has been conducted between N. parkeri and X. tropicalis18. Because of the relatively fragmented assembly of the N. parkeri genome (scaffold N50 of 1.1 Mb), most of the collinear blocks that belong to one chromosome in X. tropicalis were located on separate scaffolds in N. parkeri. For example, the orthologs of chromosome Xtr1 were mapped on scaffolds13 and 18 in N. parkeri. Thus fragmented assembly will underestimate the number of collinear blocks identified between two genomes. Despite of the fragmented assembly, whole-genome alignments between N. parkeri and X. tropicalis still revealed a large amount of synteny18. Nevertheless, collinear blocks may vary in orientation or position between species. For example, we detected inversions and translocations in L. leishanense compared with the chromosomes of X. tropicalis, suggesting the presence of structural variation in amphibian genomes.
In this study, we focused on examining the molecular basis underlying the skin-derived nuptial spines in L. leishanense by combining genomic and transcriptomic analyses. A previous study has been conducted on L. boringii by analyzing the nonreference transcriptomes of males’ MS, brain, and testes between the breeding (B) and postbreeding (C) stages14. Although a total of 1181 DEGs were identified between stages B and C in males’ MS, no nuptial spine-specific GO term was enriched, which may be largely due to the lack of a reference genome. Here we designed a comprehensive sampling strategy by collecting more tissues (added DS as a control) at three developmental stages (added subadult stage A) from two sexes (added the same samples from females). Moreover, the high-quality L. leishanense genome provided a solid reference for identifying DEGs and for GO enrichments. Transcriptomic analyses revealed meaningful biological processes associated with shaping nuptial spines, such as epithelial cell differentiation, steroid biosynthesis, and TH transport, which have not been identified previously. In addition, genomic-level examination allowed us to analyze the species-specific duplication of krts and the homologous relationship with HK genes in mammals. Based on the expression pattern of krts, we conclude that duplicated HK genes are closely associated with shaping nuptial spines. The study on L. boringii also obtained one krt (comp28513_c0) expressed differentially; however, it is difficult to extensively annotate krts based solely on transcriptomic data. Therefore, the reference genome of L. leishanense contributes greatly to the exploration of the molecular bases underlying nuptial spine formation.
Notably, the duplication of HK genes also occurs in the other four Neobatrachia anurans (N. parkeri, R. catesbeiana, R. marina, and O. pumilio). Most of these terrestrial anurans present skin-derived SD structures, such as tiny nuptial pads on the male chest (N. parkeri)52 and fingers (R. catesbeiana and R. marina)53,54. The skin of male O. pumilio exhibits specialized dorsal color55. In contrast, the aquatic Xenopus species (X. laevis and X. tropicalis) only have one type I HK gene and do not have any type II HK gene (Fig. 3a). Such differences in the number and duplication of HK genes between terrestrial and aquatic anurans suggest that HK genes may experience species-specific divergence and support the formation of diverse structures in the adaptation to terrestrial habitats in amphibians. Therefore, extensive studies on keratin GFs across amphibians are required.
However, there remain several unanswered questions. First, sex chromosomes that differ between males and females are important to understand the molecular mechanisms underlying sexual dimorphism. Insights obtained from model species, particularly fruit flies, proved that the central mechanism underlying the generation of SD traits is the integration of sex-determination system and the genetic networks governing trait development8. Sex chromosomes are usually associated with sex determination. Thus identifying sex chromosomes is crucial for screening genes that control the development of sexual dimorphism. Unfortunately, based on available data, we cannot identify sex chromosomes due to the lack of morphologically distinguishable sex chromosomes in L. leishanense28. In fact, the sex chromosome system varies considerably in amphibians, and the heterogametic sex (either XX/XY or ZZ/ZW) can vary even between populations of a single species56. Pool sequencing based on confidently sexed males and females from multiple populations and assembly to the reference genome may be helpful for identifying sex-specific molecular markers and localizing sex chromosomes. However, the success of pool sequencing depends on the size of detectable differences between sex chromosomes. Very small genetic differences may be difficult to accurately screen.
In addition to the differences in gene content on sex chromosomes, the secretion of sex steroids (androgens and estrogens) differs between males and females in vertebrates, which is another factor affecting sexual dimorphism. Here we found two pathways involved in the hypothalamus–pituitary–gonad regulation axis were significantly enriched in the brain and testes, suggesting a high level of androgen biosynthesis in male testes at stage B. In addition, three genes involved in testosterone synthesis were present at significantly high levels in the male MS during spine development, suggesting local accumulation of androgens in MS. These findings reveal the essential role of androgens in the formation of nuptial spines. However, it is difficult to directly identify the regulatory processes of androgens in triggering the expression of nuptial spine-related genes.
We also observed the involvement of other hormones, such as PRL and RLN, supported by sex-related DEGs in the MS. PRL, which was named for its role in the promotion of lactation in mammals, has been found to serve multiple roles in reproduction57. Interestingly, PRL was reported to induce the formation of nuptial pads in combination with TH in male red-spotted newts58. Thus we propose that PRL plays an essential role in regulating the production of nuptial spine in L. leishanense. RLN is another small peptide hormone that was broadly implicated in regulating the female reproductive process and improving spermatogenesis in vertebrates59,60. However, its role in the regulation of SD trait development has rarely been reported14. We found that the gene encoding RLN is highly and specifically expressed in the male MS, suggesting the crucial role in nuptial spine production in L. leishanense. Further studies on hormone regulation are needed to examine the roles of different hormones in regulating nuptial spine development in L. leishanense.
Samples used for different sequencing methods
The Leishan moustache toad samples used in this study were collected from the Leishan county, Guizhou Province, China. For genomic sequencing, we collected muscle from one adult male. Muscle and other eight tissues from the same individual (including DS, MS, brain, testis, liver, heart, kidney, and spleen) were collected for transcriptomic sequencing. For the Hi-C library preparation, we collected 4 ml of heart blood from four adult males to achieve enough sample volume. For comparative transcriptomic sequencing, RNA was extracted from four types of tissues (MS, DS, brain, and gonads) in two sexes at three developmental stages (Supplementary Fig. 9). Each sample includes three biological replicates. For the qPCR validation, we used MS and DS from adult males and adult females during the breeding season. For detailed information on sampling strategies, please see Supplementary Table 9. All experiments involving animals in this study were approved by the Animal Ethics Committee of the School of Life Sciences, Central China Normal University (CCNU-IACUC-2019–008). We have complied with all relevant ethical regulations for animal testing and research.
Illumina sequencing and genome survey
Muscle tissue from an adult male was flash frozen in liquid nitrogen. Genomic DNA was extracted using the DNeasy Blood & Tissue Kit (Qiagen, Valencia, CA, USA). Eight paired-end libraries and 12 mate-pair libraries were prepared following Illumina protocols. Library sequencing was performed on the Illumina HiSeq 4000 system (Illumina, San Diego, CA, USA). After removal of sequencing adapters, contaminant reads (mitochondrial, bacterial, and viral sequences), and low-quality reads, we finally obtained 572.38 Gb (~163× coverage) of clean reads (Supplementary Table 10). We estimated the genome size by k-mer distribution (k = 21, Supplementary Fig. 1).
Sequencing of PacBio long reads and Hi-C libraries
Genomic DNA was sheared by a g-TUBE device (Covaris, Woburn, MA, USA) setting with 20 kb. The sheared DNA was purified and concentrated with AmpureXP beads (Agencourt, Beverly, MA, USA) and further used for Single Molecular Real Time (SMRTbell) library preparation according to the PacBio 20-kb template preparation protocol. The isolated SMRTbell fractions were purified and used for primer and polymerase (P6) binding according to the manufacturer’s binding calculator (Pacific Biosciences, Menlo Park, CA, USA). Single-molecule sequencing was conducted on a PacBio RS-II platform with C4 chemistry. After adapter removal, we totally obtained 285.81 Gb of subreads (80.3× coverage), with an average length of 10.03 kb.
Hi-C libraries were created from the whole blood cells (WBCs) from adult males. According to the protocol, nuclear DNA from WBCs was cross-linked and enzymatically digested with Hind III, leaving pairs of distally located but physically interacting DNA molecules attached to each other. The sticky ends of the digested fragments were biotinylated and ligated to each other to form chimeric circles. Biotinylated circles, which are chimeras of physically associated DNA molecules from the original cross-linking, were enriched, sheared, and sequenced with the Illumina HiSeq 2500 platform. A total of 517.9 million clean Hi-C reads pairs (155.16 Gb, 44× coverage) were obtained.
Genome assembling processes
We first used Illumina reads (163×) and PacBio long reads (10×) to obtain the initial assembly (this version is referred to as Illumina-and-PacBio Assembly, IPA, Supplementary Table 1). The paired-end reads were assembled to contigs using Platanus61. The resulting contigs and PacBio reads were assembled to hybrid contigs with DBG2LOC62. Then we used SSPACE63 to assemble contigs to scaffolds based on the mate-pair reads, which resulted in the IPA version of the reference genome with a contig N50 of 408.18 kb and scaffold N50 of 754.32 kb (Supplementary Table 1). Considering the poor quality of the IPA version and the high proportion of repeat sequences estimated from the genome survey, we chose to use pure PacBio reads to assemble the genome of L. leishanense.
We used the 285.81 Gb of PacBio reads to assemble the second version of genome. Prior to assembly, we used the error correction module in Canu v1.5 to correct long subreads. Sequencing errors were corrected with a corrected error rate of 0.025. The corrected subreads were used for genome assembly using WTDBG v1.1.006. The draft genome was polished using Pilon v1.22 with 174.86 Gb of paired-end reads, which produced a L. leishanense reference genome with a contig N50 of 2.29 Mb (this version is referred to as PacBio-and-Illumina Polishing, version PIP, Supplementary Table 1).
Then we used the Hi-C data to correct the PIP version and assembled contigs to chromosome-level scaffolds. We used the Hi-C interaction signals to correct assembling errors. Contigs within the assembled genome were broken into fragments with a length of 50 kb. These fragments were used for correcting errors by reassembling based on Hi-C interaction signals. We regarded those positions that cannot be restored to the original positions on the assembled genome as candidate error regions. Within these regions, the positions with low Hi-C coverage were identified as error points. Based on the Hi-C correction, the contig N50 of the PIP version decreased slightly from 2.29 to 1.93 Mb. Then we assembled 3296 (out of 8601) contigs to 13 pseudochromosomes. We finally generated a 3.54 Gb of the L. leishanense reference genome, with a contig N50 of 1.93 Mb and scaffold N50 of 394.69 Mb (this version is referred to as PacBio-Illumina polishing and Hi-C assembly, version PIH, Supplementary Table 1), which was used for subsequent analyses.
Evaluation of genome assembly
To evaluate the quality of the L. leishanense reference genome, we aligned the Illumina reads onto the assembly. We also aligned 258,442 unigenes (≥100 bp) from 9 tissues on the assembly to assess the completeness of gene regions. In addition, we identified conserved eukaryotic core genes and single-copy orthologs in tetrapods based on the CEGMA v2.5 and BUSCO v3 databases, respectively. Based on the standard evaluation procedures, we searched 208 (out of 248, 83.9%) CEGs and 3142 (out of 3950, 79.5%) BUSCO genes in the assembly. To further check whether the missing genes exist in the assembled genome, we aligned the predicted genes in our genome against the hmmer profiles of missing genes in the CEGMA and BUSCO databases, respectively. For the CEGMA evaluation, we defined the genes with >70% of coverage as the complete CEGs based on the standard procedure. We finally found 33 complete CEGs presented in our assembly, which were missed in the standard evaluation procedure. To further explore why these CEGs cannot be detected based on the standard procedure, we examined the features of missing genes and found that these genes usually have longer introns (Supplementary Fig. 3). For the BUSCO evaluation, we also searched the missing genes based on the hmm model. We finally identified 421 previously missed BUSCO genes in L. leishanense genome, which resulted in 97.22% of complete and fragmented BUSCOs. Similar with the missing CEGs, the missing genes that omitted by original BUSCO evaluation procedure are featured by long introns compared with the complete BUSCOs. Similar phenomenon also exists in other anuran genomes (Supplementary Fig. 4).
Genome annotation and chromosome synteny analysis
We constructed the de novo repetitive sequence database of L. leishanense and combined with the Repbase database 20.01 to create the final repeat library (for detailed software list, please see Supplementary Table 6). Repeat sequences in the L. leishanense genome were identified and classified using RepeatMasker 4.0.664. The LTR family classification criterion was defined by the 5’LTR sequences of the same family that shared at least 80% identity over at least 80% of their lengths. To compare repeat sequences across different anurans, we applied the same methods for six other genomes (N. parkeri, R. catesbeiana, X. tropicalis, X. laevis, R. marina, and O. pumilio).
We integrated three approaches, namely, de novo prediction, homology search, and transcript-based assembly, to annotate protein-coding genes in a repeat-masked genome (for detailed software list, please see Supplementary Fig. 5). Consensus gene models were generated by integrating the de novo prediction and protein and transcript alignments using EVidenceModeler v1.1.165. To assign gene functions, the predicted gene sequences were searched against seven databases: NR, GO, KEGG, KOG, Pfam, SwissProt, and TrEMBL.
We analyzed the chromosome synteny between L. leishanense and X. tropicalis via all-to-all BLASTP searches of protein sequences (with an E-value cut-off of 1e−5). Collinear blocks containing at least 10 genes (-s 10) and a maximum of 25 gaps (genes) between two proximal orthologs within a block (-m 25) were identified using MCScanX66. The serial numbers of the chromosomes were manually adjusted to reflect the descending order of chromosome length (Lle1 is the longest chromosome of L. leishanense, and Lle13 is the shortest chromosome).
Comparative genomic analyses
Orthologous groups among 11 species (including D. rerio, A. carolinensis, H. sapiens, M. musculus, N. parkeri, X. tropicalis, X. laevis, O. pumilio, R. marina, R. catesbeiana, and L. leishanense) were constructed using OrthoMCL v2.0.967 based on an all-to-all BLASTP strategy (with an E-value of 1e−5). We extracted 881 single-copy genes from the 11 species and aligned proteins for each gene. All the alignments were combined to one supergene to construct a phylogenetic tree using RAxML v7.2.8 with 1000 rapid bootstraps followed by a search of the best-scoring maximum likelihood (ML) tree in one single run. Divergence time was estimated using the MCMCTree program in PAML v4.9 under the relaxed clock model. Several calibrated time points were used to date the divergence time in the unit of Ma (Supplementary Table 11).
A GF was defined as a group of similar genes that descended from a single gene in the last common ancestor. Expansion and contraction of GFs were determined using CAFÉ v3.168 based on changes in GF size. The cluster size of each branch was compared with the cluster size of the ancestral node. p value was calculated using the Viterbi method under the hidden Markov model, with p < 0.05 defining significant expansion or contraction. Genes belonging to expanded GFs were subjected to GO and KEGG enrichment. p values were calculated with two-sided Fisher’s exact test and corrected by the BH procedure.
To identify positively selected genes (PSGs) in L. leishanense, we first extracted the orthologous genes of L. leishanense and six other anurans via reciprocal best alignment. The protein sequences were aligned and removed sites with gaps. The alignments were used for PSG identification. We used the branch-site model of CODEML in PAML v4.9 by setting the L. leishanense as the foreground branch and the six other anurans as background branches. We allowed ω values (the ratio of the rate of nonsynonymous substitutions to the rate of synonymous substitutions) to vary among sites in the foreground clade following three categories (0 < ω0 < 1; ω1 = 1; ω2 > 1). The likelihood of M2a was compared with that of the null model M1a (0 <ω0 <1 and ω1 = 1) by performing a likelihood ratio test (defined as twice the log likelihood difference between the M2a and M1a) and calculating the corresponding p values. Sites showing a signature of positive selection were identified by calculating the posterior probability that a site belongs to a category with ω > 1 using the Bayes Empirical Bayes approach. Genes with p < 0.05 and containing at least one codon with a posterior probability >0.95 were defined as PSGs.
Keratin GF analyses
The structures of keratin genes in L. leishanense were determined using the software GeMoMa v1.4.269 based on the keratin gene models available from the reference organisms (H. sapiens, M. musculus, A. carolinensis, Gallus gallus, X. laevis, X. tropicalis, and N. parkeri). The predictions were handled using a GeMoMa annotation filter with default parameters except for the evidence percentage filter (e = 0.1), and then the predicted krt genes were manually checked to obtain a single high confidence transcript prediction per locus. Keratin genes in R. catesbeiana, O. pumilio, and R. marina genomes were also searched using GeMoMa v1.4.2. Then the keratin gene sequences from the 11 species were aligned and used to construct an ML tree in RAxML v7.2.8.
Samples from the DS, MS, brain, and gonads were collected at three developmental stages from males and females. Each sample included three replicates; thus we collected a total of 72 samples (Supplementary Fig. 9). Total RNA was isolated using the TRIzol reagent (Invitrogen, Carlsbad, CA, USA) followed by treatment with RNase-free DNase I (Promega, Madison, WI, USA) according to the manufacturers’ protocols. RNA quality was checked using an Agilent 2100 Bioanalyzer. Illumina RNA-seq libraries were prepared for 72 samples and sequenced on a HiSeq 2500 system with a PE150 strategy following the manufacturer’s instructions.
The expression level of predicted transcripts in each RNA-seq library was calculated as the transcripts per million (TPM) using the following formula: TPM = (CDS read count × mean read length × 106)/(CDS length × total transcript count). To identify sex-related DEGs, we compared TPM values from various tissues between males and females. Taking MS (tissue #2) as an example, we separately compared AM2 vs AF2, BM2 vs BF2, and CM2 vs CF2. Similar comparisons were conducted for the brain and gonads. In addition, to examine genes specific to the MS, we used DS (tissue #1) as a background and compared MS vs DS in males and females separately (in males: AM1 vs AM2; BM1 vs BM2; CM1 vs CM2; in females: AF1 vs AF2; BF1 vs BF2; CF1 vs CF2). For each pairwise comparison, DEGs were identified by |log2(fold change)| > 1 and BH corrected p < 0.01 in the DESeq package70. The potential functions of the DEGs were examined by GO and KEGG enrichment.
To validate the accuracy of DEGs identified from transcriptomic data, we selected five genes and tested the relative expression levels of these genes between the males DS and MS at stage B (BM1 vs BM2) and the expression levels between the males’ and females’ MS at stage B (BM2 vs BF2) using qPCR. The reactions were performed in a CFX96 Touch RealTime PCR Detection System (Bio-Rad, Richmond, CA, USA) using TransStart Tip Green qPCR SuperMix (TransGen, Beijing, China) in a 15-µl volume with 1.5 µl of cDNA. The expression levels of the tested genes were normalized to the level of gapdh (glyceraldehyde-3-phosphate dehydrogenase). Two-sided Student’s t test was used to evaluate the significance of differential expression between samples. Primers are listed in Supplementary Table 12. Three biological replicates were examined for each sample.
Gene coexpression analysis
To cluster genes with similar expression patterns across samples, we conducted coexpression analysis based on 72 samples using WGCNA v1.63. We constructed an unsupervised network for transcriptome data using the function blockwiseModules with default parameters. First, a matrix of Pearson correlations between genes was generated based on TPM values across samples. Then an adjacency matrix representing the connection strength among genes was constructed by raising the correlation matrix to a soft threshold power to achieve a scale-free topology fit index of 0.80. Next, the adjacency matrix was used to calculate the topological overlap matrix (TOM). Genes with similar coexpression patterns across samples were grouped using hierarchical clustering of dissimilarity among the topological overlap measures (1 − TOM). Coexpressed modules were determined using a dynamic tree cutting algorithm setting with a minimum module size of 30 and a cut height of 0.998. For each module, GO and KEGG enrichments were conducted to understand the enriched functions. An eigengene value (the first principal component of the scaled module expression profiles) was calculated to characterize the overall expression trend for each module. The intramodular connectivity was measured as kME values that represent the Pearson correlation between the expression level of that gene and the ME. Then the Pearson correlations between ME values and sampling trait values were calculated to measure the strength and direction of association between modules and traits. Fisher’s asymptotic p values were calculated for given correlations using the corPvalueFisher module. Significant module–trait associations were considered when p < 0.05.
Further information on research design is available in the Nature Research Reporting Summary linked to this article.
All the data have been deposited in the NCBI database under the BioProject PRJNA505224. Specifically, the assembled version IPA of the Leishan moustache toad genome has been deposited in NCBI Genbank (accession: RXON00000000). The HDF5 raw data for PacBio sequencing have been deposited in NCBI SRA database (accession numbers: SRR8897348–SRR8897543; SRR9670029–SRR9670067). The RNA-seq reads for 72 samples have been deposited in NCBI SRA database (accession numbers: SRR8736149–SRR8736220). The Hi-C library reads (including eight libraries) have been deposited in the SRA (accession numbers: SRR8784800–SRR8784807). The Illumina paired-end reads have been deposited in the SRA (accession numbers: SRR8788204–SRR8788209; SRR10019514–SRR10019515). The Illumina mate-pair reads have been deposited in the SRA (accession numbers: SRR10019502–SRR10019513). All these raw data can be downloaded under study SRP188598 [https://trace.ncbi.nlm.nih.gov/Traces/sra/?study=SRP188598]. The annotation files can be found in Figshare (https://figshare.com/; https://doi.org/10.6084/m9.figshare.8019986). Other miscellaneous information are available from the corresponding authors upon request. Published genome data used in the analyses can be found under the following accession codes: X. tropicalis (GCF_000004195.3); X. laevis (GCF_001663975.1); N. parkeri (GCF_000935625.1); R. catesbeiana (GCA_002284835.2); O. pumilio ([https://academic.oup.com/mbe/article/35/12/2913/5106668#supplementary-data]); R. marina (5524/100483); D. rerio (GCF_000002035.6); A. carolinensis (AnoCar2.0 [ftp://ftp.ensembl.org/pub/release-90/fasta/anolis_carolinensis/dna/]); M. musculus (GRCm38 [ftp://ftp.ensembl.org/pub/release-90/fasta/mus_musculus/dna/]); H. sapiens (GRCh38 [ftp://ftp.ensembl.org/pub/release-90/fasta/homo_sapiens/dna/]).
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We are thankful to Professor Patricia Wittkopp and all members of the Wittkopp laboratory from the University of Michigan, Ann Arbor (USA) for their helpful comments and suggestions for this paper. We thank Dr. Jinzhong Fu from the University of Guelph for his kind help with explaining the results and writing the manuscript. We are thankful to Dr. Rebekah Rogers and Dr. Rasmus Nielsen for sharing the genomic data of the strawberry poison frog. This work was supported by the National Natural Science Foundation of China (No.31770405) and the Biodiversity Survey, Monitoring and Assessment Project of Ministry of ecology and environment, China (2019–2023).
The authors declare no competing interests.
Peer review information Nature Communications thanks Andrew Crawford and the other anonymous reviewer(s) for their contribution to the peer review of this work.
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Li, J., Yu, H., Wang, W. et al. Genomic and transcriptomic insights into molecular basis of sexually dimorphic nuptial spines in Leptobrachium leishanense. Nat Commun 10, 5551 (2019). https://doi.org/10.1038/s41467-019-13531-5
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