Desert Hedgehog mediates stem Leydig cell differentiation through Ptch2/Gli1/Sf1 signaling axis

Abstract

Desert Hedgehog (Dhh) mutations cause Leydig cell dysfunction, yet the mechanisms governing Leydig lineage commitment through Dhh-mediated receptor selectivity, transcriptional effector specificity, and steroidogenic coupling remain elusive. In this study, using CRISPR/Cas9-mediated gene knockout and stem Leydig cells (SLCs) transplantation, we identified a critical Dhh/Patched 2 (Ptch2)/Glioma-associated oncogene homolog 1 (Gli1)/steroidogenic factor 1 (Sf1) signaling axis essential for SLC differentiation in Nile tilapia (Oreochromis niloticus). Dhh deficiency resulted in defective adult Leydig cells and androgen insufficiency. Rescue experiments involving 11-ketotestosterone administration and a Dhh agonist treatment, combined with SLCs transplantation, demonstrated that Dhh regulates SLC differentiation, not survival. In vitro knockout of ptch1 and ptch2 in SLCs revealed that Ptch2 likely acts as the functional receptor for Dhh. This was further supported by in vivo genetic rescue experiments, where ptch2 mutation did not impair testicular development, yet completely rescued the testicular defects in dhh mutants—consistent with Ptch2 acting as an inhibitory receptor whose loss alleviates Dhh pathway suppression. Luciferase assays in Gli-knockout SLCs demonstrated that Gli1 acts as the primary transcriptional effector and transactivates sf1 expression. Additionally, functional transplantation assays confirmed that Sf1 is indispensable for SLC differentiation, as Sf1-overexpressing SLCs rescued differentiation, whereas sf1-mutant SLCs failed. Overall, our work delineates the Dhh-Ptch2-Gli1-Sf1 axis and provides fundamental insights into the endocrine regulation of Leydig cell lineage development.

Introduction

Leydig cells are the primary source of androgens, producing approximately 95% of circulating testosterone to sustain spermatogenesis and male fertility (Inoue et al., 2018; Jiang et al., 2014). Leydig cells originate from stem Leydig cells (SLCs), which differentiate into steroidogenic adult Leydig cells (ALCs) under paracrine regulation (Chen et al., 2020; Yao et al., 2022). Steroidogenic factor 1 (Sf1), an orphan nuclear receptor transcription factor, serves as a master regulator of this process by activating steroidogenic enzymes (e.g. cytochrome P450, family 11, subfamily C, polypeptide 1 [Cyp11c1], 3β-hydroxysteroid dehydrogenase [3β-HSD]) to drive ALC maturation (Parker et al., 2002). In mice, mutations of Sf1 exhibit arrested testicular development and abolished steroidogenesis (Houzelstein et al., 2024; Park et al., 2007; Smith et al., 2023), while human SF1 mutations are associated with 46,XY disorders of sexual development (DSD), such as Swyer syndrome (Yu et al., 2023). Similarly, in Nile tilapia (Oreochromis niloticus), our previous work reveals that sf1 mutation disrupts testicular morphogenesis and downregulates the expression of key steroidogenic enzymes such as cyp11c1 (Xie et al., 2016). Despite its established central role, the upstream signaling pathways that regulate Sf1 to commit the Leydig lineage remain poorly defined.

Hedgehog (Hh) signaling pathway is an evolutionarily conserved developmental regulator (Zhang and Beachy, 2023). Canonical Hh signaling involves the binding of ligands to Patched (Ptch) receptors, which relieves the suppression on Smoothened (Smo) and ultimately activates Glioma-associated oncogene homolog (Gli) transcription factors to regulate target genes (Briscoe and Thérond, 2013; Finco et al., 2015; Ingham et al., 2011). In vertebrates, there are three Hh ligands (i.e. Sonic Hh, Indian Hh, and Desert Hh [Dhh]), two Ptch receptors (Ptch1 and Ptch2), and three Gli homologs (Gli1–3) (Belgacem and Borodinsky, 2015; Carpenter et al., 1998; Cervantes et al., 2010; Kothandapani et al., 2020). A large number of studies have well demonstrated that the Dhh signaling pathway is critical in gonadal development (Finco et al., 2015; Umehara et al., 2000). Dhh, secreted by Sertoli cells, is critical for Leydig cell commitment in mammals (Mehta et al., 2021; Pachernegg et al., 2022). In mice, mutations of Dhh lead to Leydig cell dysfunction, testosterone insufficiency, and infertility (Pierucci-Alves et al., 2001), while human DHH mutations are associated with 46,XY DSD (Wei et al., 2022). Strikingly, these phenotypes mirror those caused by Sf1 deficiencies, suggesting a potential regulatory nexus between Dhh and Sf1 signaling (Park et al., 2007). However, the molecular circuitry connecting Dhh signaling to Sf1 activation remains elusive.

Studies demonstrate functional divergence between Ptch1 and Ptch2. For instance, Ptch1 knockout causes embryonic lethality due to constitutive Hh activation, whereas Ptch2 deficiency yields no overt defects (Cong et al., 2025; Nieuwenhuis et al., 2006). During testicular development, Ptch1 displays high expression at an early stage critical for gonadal primordium formation, while Ptch2 shows testis-enriched dynamic regulation (Kim et al., 2020; Yao et al., 2002). However, the distinct roles of Ptch1 and Ptch2 in Leydig lineage commitment remain unknown to date. Among Gli transcription factors, Gli1 acts as a primary activator, while Gli2 and Gli3 exhibit dual regulatory functions (Hui and Angers, 2011; Ruiz i Altaba et al., 2002). In murine testes, co-expression of Gli1 and Gli2 in Leydig cells illustrates functional redundancy—single knockouts show normal development, but combined pharmacological inhibition (GANT61) abolishes Leydig cells and spermatogenesis (Barsoum and Yao, 2011). These findings underscore the complexity of Gli-mediated transcription in the testis (Kothandapani et al., 2020) and highlight the need to delineate the specific roles of individual Gli in SLC differentiation.

Nile tilapia (O. niloticus) is not only an important farm fish in global aquaculture but also a distinguished animal model for investigating gene function and endocrine regulation. This model organism possesses multiple advantageous attributes, including an XX/XY sex-determination system, early sexual maturity (approximately 3 months after hatching for males and 6 months after hatching for females), a short spawning cycle of approximately 14 days, year-round reproductive capacity under laboratory conditions, well-characterized sex-linked genetic markers, and a body size amenable to physiological sampling (Li et al., 2024; Sun et al., 2014). Furthermore, an SLC line (TSL) has been established from the testis of a 3-month-old Nile tilapia (Huang et al., 2020). TSL expresses platelet-derived growth factor receptor α (pdgfrα), nestin, and chicken ovalbumin upstream promoter transcription factor II (coup-flla), which are usually considered as SLC-related markers in several other species (Chen et al., 2017; Ge et al., 2006; Jiang et al., 2014). Notably, this cell line exhibits the capacity to differentiate into 11-ketotestosterone (11-KT)-producing Leydig cells both in vitro and in vivo (Huang et al., 2020). When cultured in a defined induction medium, TSL cells differentiate into a steroidogenic phenotype, expressing key steroidogenic genes, including star1, star2, and cyp11c1, and producing 11-KT; upon transplantation into recipient testes, TSL cells successfully colonize the interstitial compartment, activate the expression of steroidogenic genes, and restore 11-KT production. In this study, by combining CRISPR/Cas9-mediated gene knockout with TSL transplantation technology, we unveil a Dhh-Ptch2-Gli1-Sf1 axis that is essential for Leydig cell lineage differentiation. Our findings provide fundamental insights into the endocrine regulation of Leydig cell development.

Results

Mutation of dhh disrupts testicular organization and androgen synthesis

The Nile tilapia Dhh open reading frame (GenBank ID: 100707022, 1377 bp) was cloned and validated. The encoded 458-amino acid protein shares over 56% identity with vertebrate homologs (Figure 1—figure supplement 1A) and clusters phylogenetically with mammalian DHH orthologs (Figure 1—figure supplement 1B). Transcriptomic analyses conducted on various tissues and testes across different developmental stages revealed that dhh expression peaked in the testes, particularly at 90 days after hatching (dah) (Figure 1—figure supplement 1C, D).

CRISPR/Cas9 targeting of dhh exon 1 from Nile tilapia generated homozygous mutants (dhh^-/-) harboring a 13 bp (CAGGGATGCGGAC) frameshift deletion (Figure 1—figure supplement 2A–D), resulting in premature termination and loss of the conserved Hedge domain (Figure 1—figure supplement 2E). dhh mRNA levels in dhh^-/- fish were significantly reduced compared to WT (Figure 1—figure supplement 2F). At 90 dah, dhh^-/- testes exhibited marked atrophy (Figure 1A and B) and disorganized histology, with sparse spermatogonia, rare spermatids, and collapsed interstitial architecture (Figure 1C–D’ and K). Immunofluorescence (IF) confirmed significantly reduced germ cell (Vasa⁺), meiotic cell (Sycp3⁺), and Leydig cell (Cyp11c1⁺) populations (Figure 1E–J’ and L–N). Serum 11-KT levels in dhh^-/- fish were significantly lower than WT (Figure 1O), indicating impaired androgen synthesis.

Figure 1. Mutation of dhh resulted in testicular developmental disorders.

(A–B) Morphological analysis of the testes from wild-type (WT) (dhh^+/+) and dhh^-/- XY fish at 90 days after hatching (dah). Arrows indicate the location of the testes. (C–D’) Histological analysis of testicular sections from WT and dhh^-/- XY fish at 90 dah. Sg, spermatogonia; Sc, spermatocyte. (E–J’) Representative immunofluorescence images showing the expression of germ cell marker Vasa (E–F’), meiosis cell marker Sycp3 (G–H’), and Leydig cell marker Cyp11c1 (I–J’). (K) Gonadosomatic index (GSI) of the testes from WT and dhh^-/- XY fish at 90 dah (n=5 fish/genotype). (L–N) Quantification of the percentage of cells positive for Vasa (E–F’), Sycp3 (G–H’), and Cyp11c1 (I–J’) among all DAPI-positive cells in the WT and dhh^-/- testes (n=5 fish/genotype). (O) Serum 11-KT level of WT and dhh^-/- XY fish at 90 dah (n=6 fish/genotype). Values were presented as mean ± SD. Differences were determined by two-tailed independent Student’s t-test. **, p<0.01. Scale bars: (A–B), 1 cm; (C–J), 50 μm.

Figure 1—figure supplement 1. Sequence analysis and mRNA expression profile of Nile tilapia dhh.

(A) Dhh amino acid sequence alignment. Amino acids are numbered in the right margin. Deletions are indicated by dashes; shaded areas indicate shared sequences. The amino-terminal hedge domain is underlined by a dotted line, with the signal peptide region in the front and the autocatalytic carboxy-terminal domain (hog) in the back. The box indicates the autocatalytic site of an absolutely conserved Gly-Cys-Phe tripeptide. At the end of the alignment are percentage identity values of the full-length and hedge domain of tilapia Dhh to the orthologue from other species. (B) Phylogenetic analysis of Dhh. The phylogenetic tree was constructed using the neighbor-joining method within the MEGA7.0 program. Node values represent percent bootstrap confidence derived from 1000 replicates. (C–D) The RPKM (reads per kb per million reads) values of tilapia dhh in various adult tissues (C) and XY and XX gonads at 5, 7, 20, 30, 90, 180, 300 dah (days after hatching) (D) in transcriptome sequencing data which were sequenced using Illumina 2000 HiSeq technology in our previous study (Tao et al., 2013).

Figure 1—figure supplement 2. Establishment of the Nile tilapia dhh mutant line by CRISPR/Cas9.

(A) Schematic representation of guide RNA (gRNA) targeting the Nile tilapia dhh locus. The gRNA was designed to target exon 1. The translation start codon ATG and stop codon TAA were indicated by arrows. The PAM (protospacer adjacent motif) site was marked by a green box. DNA sequence alignment of the three mutant lines with wild-type (WT). The added sequences were marked by red letters, and the deleted sequences were indicated in red dotted lines. One had a 4 bp addition, one had an 8 bp deletion, and the other had a 13 bp deletion. (B) Schematic diagram showing the breeding plans of dhh F0 to F2 fish. (C) Homozygous mutants of F2 fish were screened by polyacrylamide gel electrophoresis (PAGE). The first lane is DNA marker, the second lane is dhh^+/+ fish, the third lane is dhh ^+/- fish, and the fourth lane is dhh^-/- fish. (D) Sanger sequencing results of dhh genes from WT and the 13 bp deletion homozygous mutant fishes. (E) The dhh ORF sequence in WT was indicated in black and the frameshift-altered sequence for the 13 bp deletion dhh ORF was indicated in red. (F) The relative mRNA expression of dhh in WT and the 13 bp deletion homozygous mutant fishes by RT-qPCR. Values were presented as mean ± SD (n=3). Differences were determined by two-tailed independent Student’s t-test. **, p<0.01.

The differentiation of SLCs cannot be rescued by 11-KT, but by SAG

Given the established role of 11-KT in teleost testicular development and spermatogenesis (Zheng et al., 2020), we first asked whether exogenous 11-KT could ameliorate the testicular defects in dhh^-/- mutants. Compared to untreated mutants, dhh^-/- individuals exposed to 11-KT exhibited a more organized testicular architecture, with significantly increased populations of Vasa⁺ and Sycp3⁺, as evidenced by both hematoxylin and eosin (H&E) staining and IF (Figure 2A–C’, E–G’, I–K’, R, and S). This improvement was further supported by a recovery in gonadosomatic index (GSI) (Figure 2Q). However, the population of Cyp11c1-positive Leydig cells remained profoundly depleted in dhh^-/-+11-KT testes (Figure 2M–O’ and T). Taken together, these data indicate that 11-KT treatment partially restored the development of germ cells but not Leydig cell lineage, implying that the Leydig cell defect is androgen-independent.

Figure 2. Rescue of testicular development in dhh^-/- XY Nile tilapia by 11-ketotestosterone (11-KT) and SAG.

dhh^-/- XY Nile tilapias at 30 days after hatching (dah) were treated either with 11-KT via water immersion (with water changed every 2 days) or with 10 mg/kg SAG via intraperitoneal injection (with supplemental injection every 7 days). WT controls received 11-KT, and dhh^-/- controls received an equivalent volume of the DMSO. Then, at 90 dah, morphological and histological experiments were conducted. (A–D’) Histological analysis of testicular sections from 90 dah XY fish subjected to different treatments as indicated. Sg, spermatogonia; Sc, spermatocyte. Scale bars: 50 μm. (E–P’) Representative immunofluorescence images showing the expression of Vasa (E–H’), Sycp3 (I–L’), and Cyp11c1 (M–P’). Scale bars: 50 μm. (Q) Gonadosomatic index (GSI) of testes from the different treatment groups at 90 dah (n=5 fish per group). (R–T) Quantification of the percentage of cells positive for Vasa (E–H’), Sycp3 (I–L’), and Cyp11c1 (M–P’) among all DAPI-positive cells in the testes (n=3–5 fish per group). Values were presented as mean ± SD. Different letters above the error bar indicate statistical differences at p<0.05 as determined by one-way ANOVA followed by Tukey’s test.

Figure 2—figure supplement 1. The expression levels of Cyp11c1 in the testis and the levels of 11-ketotestosterone (11-KT) in tissue fluid from wild-type (WT) and dhh^-/- XY fish at 5, 10, 20, and 30 days after hatching (dah).

(A–E) Representative immunofluorescence images showing Cyp11c1 expression (green) in testis sections from WT fish at 5, 10, 20, 30 dah and dhh^-/- mutants at 30 dah. Scale bars: 4 μm. (F) Tissue fluid 11-KT levels in WT and dhh^-/- XY fish at indicated time points (n=6 fish per group). Values were presented as mean ± SD. Differences were determined by two-tailed independent Student’s t-test. NS, no significant difference.

We next investigated whether direct activation of the Dhh pathway could rescue the dhh^-/- phenotype. Treatment of dhh^-/- mutants with SAG, an Hh agonist, significantly rescued testicular development. This was marked by a significant recovery of germ cells (Vasa⁺), spermatocytes (Sycp3⁺), and critically, Leydig cells (Cyp11c1⁺), alongside a restoration of testicular histology (Figure 2D–D’, H–H’, L–L’, P–P’, and Q–T). These results demonstrate that dhh mutation does not disrupt survival of endogenous Leydig cell lineage, but directly impairs their differentiation, leading to secondary impairment in androgen production and germ cell development.

Mutation of dhh blocks SLC differentiation in vivo

To dissect Leydig cell lineage impairment in dhh^-/- testes, we transplanted the TSL labeled with PKH26 (a fluorescent red hydrophobic membrane dye that enables tracking of transplanted cells) into WT and dhh^-/- testes (Figure 3A). In WT recipients, TSL cells colonized the interstitium and differentiated into Cyp11c1-positive steroidogenic cells (Figure 3B1–B3). In dhh^-/- testes, TSL engrafted equivalently but failed to express Cyp11c1 (Figure 3C1–C3), indicating a differentiation-specific block. Rescue experiments showed that the TSL overexpressing tilapia Dhh (TSL-OnDhh) or treated with SAG (TSL+SAG) restored Cyp11c1 expression in dhh^-/- testes (Figure 3D1–E3 and F) and a significant increase in endogenous 11-KT levels (Figure 3G). These results suggest that SLC differentiation is inhibited, whereas the survival and engraftment of PKH26-labeled TSL cells were not affected in dhh^-/- XY tilapia testes. Collectively, these results demonstrate that Dhh signaling is specifically required for the differentiation of SLCs into steroidogenic cells, but is dispensable for their survival or initial recruitment into the testicular niche.

Figure 3. Dhh is required for stem Leydig cell (SLC) differentiation in vivo.

TSL, Dhh-overexpressing TSL (TSL-OnDhh), or 0.5 μM SAG-treated TSL (TSL+SAG) cells were labeled with PKH26 and transplanted into the testes of 90 days after hatching (dah) wild-type (WT) or dhh^-/- recipient fish. Analyses were performed 10 days post-transplantation. (A) Schematic diagram of the experimental design for SLCs transplantation and analysis. (B1–E3) Representative immunofluorescence images of testicular sections from recipient fish, showing the localization of transplanted PKH26-labeled SLCs (red) and the expression of Cyp11c1 (green). Nuclei are stained with DAPI (blue). Scale bars: 4 μm. (F) Quantification of the percentage of Cyp11c1-positive cells among the transplanted PKH26-positive SLCs for each treatment group (n=5 fish per group). (G) Serum 11-KT level in recipient fish following SLC transplantation (n=5 fish per group). Values were presented as mean ± SD. Different letters above the error bar indicate statistical differences at p<0.05 as determined by one-way ANOVA followed by Tukey’s test.

Ptch2 is the functional receptor for Dhh signaling in SLCs

RT-PCR results showed that both ptch1 and ptch2 were expressed in TSL (Figure 4—figure supplement 1). Fluorescence in situ hybridization (FISH) and IF revealed co-expression of ptch1 and ptch2 in Cyp11c1-positive Leydig cells (Figure 4A). To identify the Dhh-driving receptor selectivity in SLCs, we established ptch1^-/- and ptch2^-/- TSL cell lines (Figure 4—figure supplement 2A, B) and evaluated pathway activity using an 8×GLI luciferase reporter system (Figure 4B). In WT TSL cells, the 8×GLI reporter demonstrated substantial luciferase activity that was markedly enhanced by OnDhh overexpression (Figure 4C), confirming Dhh responsiveness in this system. Intriguingly, ptch1^-/- cells maintained Dhh-induced luciferase activation comparable to WT controls, whereas ptch2^-/- cells completely lost this responsiveness (Figure 4D). This differential receptor requirement implies that Ptch2 likely acts as the functional receptor for transducing Dhh signals in TSL cells.

Figure 4. Ptch2 mediates Dhh signaling in stem Leydig cells (SLCs).

(A) Co-localization of Cyp11c1 (green, by immunofluorescence) and ptch1 or ptch2 mRNA (red, by RNA-FISH) in adult (90 days after hatching [dah]) testis sections. Dashed lines outline representative Leydig cells. Scale bars: 4 μm. (B) Schematic illustration of the experimental setup for the luciferase reporter assays shown in panels C and D. (C) Luciferase activity in TSL cells co-transfected with a Gli-responsive reporter (8×GLI) and overexpression of tilapia Dhh (OnDhh). A plasmid lacking Gli-binding sites (pGL4.23) served as a negative control (n=3). (D) Luciferase activity in TSL-WT, TSL-ptch1^-/-, and TSL-ptch2^-/- cells transfected with the 8×GLI reporter with or without OnDhh (n=4). (E–G’) Histological analysis of testis sections from the indicated genotypes at 90 dah. Sg, spermatogonia; Sc, spermatocyte. Scale bars: 50 μm. (H–P’) Immunofluorescence analysis of Vasa (H–J’), Sycp3 (K–M’), and Cyp11c1 (N–P’) in testis sections from the indicated genotypes. Scale bars: 50 μm. (Q) Gonadosomatic index (GSI) of the testes from the indicated genotypes at 90 dah (n=4 fish per group). (R–T) Quantification of the percentage of Vasa (H–J’), Sycp3 (K–M’), and Cyp11c1 (N–P’) positive cells among DAPI-positive cells (n=6 fish per group). (U) Serum 11-ketotestosterone (11-KT) levels in fish of the indicated genotypes at 90 dah (n=6 fish per group). Statistical significance was determined by one-way ANOVA followed by Tukey’s test (C, Q–U, different letters above the error bar indicate statistical differences at p<0.05) or Student’s t-test (D) (*, p<0.05; **, p<0.01; NS, no significant difference).

Figure 4—figure supplement 1. RT-PCR analyses of Hh pathway genes in TSL cells.

β-aActin served as a loading control, and RNAs from TSL cells were used as templates for negative controls. The numbers in parentheses indicate the PCR cycles.

Figure 4—figure supplement 2. Establishment of the TSL ptch1, ptch2, gli1, gli2, gli3, and sf1 mutant lines.

(A–F) Schematic representation of guide RNAs (gRNAs) targeting the ptch1, ptch2, gli1, gli2, gli3, and sf1 locus. The gRNAs were designed to target ORF (open reading frame). The PAM (protospacer adjacent motif) sites were marked by red letters. Sanger sequencing of ptch1, ptch2, gli1, gli2, gli3, and sf1 single mutant alleles in single-cell clones. PCR amplicons from DNA templates of the six cell clones were directly used for sequencing.

Figure 4—figure supplement 3. Establishment of the Nile tilapia ptch2 mutant line by CRISPR/Cas9 system.

(A) Schematic representation of guide RNA (gRNA) targeting the Nile tilapia ptch2 locus. The gRNA was designed to target exon 1. The translation start codon ATG and stop codon TGA were indicated by arrows. The PAM site was marked by a green box. DNA sequence alignment of the five mutant lines with wild-type (WT). The added sequences were marked by red letters, and the deleted sequences were indicated in red dotted lines. (B) Schematic diagram showing the breeding plans of ptch2 F0 to F2 fish. (C) DNA sequencing showed the 25 bp deletion within the ptch2 ORF in the homozygous mutant compared to the WT. (D) Schematic of the prediction of the intact Ptch2 protein in the WT and the truncated Ptch2 protein in the homozygous mutants. (E) Homozygous mutants of F2 fish were screened by polyacrylamide gel electrophoresis (PAGE). The first lane is DNA marker, the second lane is ptch2^+/+ fish, the third lane is ptch2^+/-fish, and the fourth lane is ptch2^-/- fish. ‘*’ indicates heteroduplex, and arrows indicate homoduplex.

Figure 4—figure supplement 4. Expression analysis of ptch1 in wild-type (WT) and mutant XY testes.

Relative mRNA expression levels of ptch1 in XY testes from WT, ptch2^-/-, and dhh^-/-;ptch2^-/- fish at 90 days after hatching (dah) by RT-qPCR. Values were presented as mean ± SD (n=3 fish per group). Statistical significance was determined by one-way ANOVA followed by Tukey’s test. NS, no significant difference.

To investigate the function of Ptch2 in vivo, we generated ptch2^-/- and dhh^-/-;ptch2^-/- mutant Nile tilapias (Figure 4—figure supplement 3). Compared to WT fish at 90 dah, ptch2^-/- testes exhibited no significant differences in testicular histology (Figure 4E–F’), GSI (Figure 4Q), germ cell (Vasa⁺, Sycp3⁺), and Leydig cell (Cyp11c1⁺) populations (Figure 4H–I’, K–L’, N–O’, and R–T), and serum 11-KT levels (Figure 4U). Strikingly, the loss of ptch2 in the dhh^-/- background (dhh^-/-;ptch2^-/-) fully rescued the testicular defects, resulting in normal histology (Figure 4G–G’), Leydig cell differentiation (Figure 4P–P’), and serum 11-KT levels (Figure 4U). This genetic rescue is consistent with Ptch2 functioning as the inhibitory receptor for Dhh; its removal alleviates the suppression on the pathway, thereby bypassing the requirement for the Dhh ligand.

Gli1 is the principal transcriptional effector of Dhh signaling in SLCs

In the Hh pathway, Gli transcription factors (Gli1/2/3) mediate diverse developmental processes (Niewiadomski et al., 2019). RT-PCR results showed that gli1, gli2, and gli3 were all expressed in TSL (Figure 4—figure supplement 1). FISH and IF revealed co-expression of gli1, gli2, and gli3 in Cyp11c1-positive Leydig cells (Figure 5A). To identify Dhh-specific effectors, we generated gli1^-/-, gli2^-/-, and gli3^-/- TSL lines (Figure 4—figure supplement 2C–E) and assessed pathway activity using the 8×GLI luciferase reporter. While TSL-gli2^-/- and TSL-gli3^-/- cell lines maintained Dhh-induced luciferase activation comparable to WT controls, the TSL-gli1^-/- cell line completely lost this responsiveness (Figure 5B), implying that Gli1 is the principal transcriptional effector for Dhh signaling transduction in SLCs.

Figure 5. Gli1 transactivates sf1 to drive stem Leydig cell (SLC) differentiation.

(A) Co-localization of Cyp11c1 (green) and gli1, gli2, or gli3 mRNA (red) in adult (90 days after hatching [dah]) testis sections by immunofluorescence and RNA-FISH. Dashed lines outline representative Leydig cells. Scale bars: 4 μm. (B) Luciferase activity in TSL-WT, TSL-gli1^-/-, TSL-gli2^-/-, and TSL-gli3^-/- cells transfected with the 8×GLI reporter with or without OnDhh (n=4). (C) Volcano plots of transcriptomic changes in TSL-OnDhh, TSL-OnGli1, and TSL+SAG, compared to TSL-WT. Red and blue dots represent significantly up- and downregulated genes, respectively. (D) Schematic of the tilapia sf1 gene promoter, indicating the two predicted Gli1 binding sites (B1 and B2). (E) Transcriptional activation of the sf1 promoter by Gli1. HEK293 cells were co-transfected with pRL-TK, pGL3, pcDNA3.1, pGL3-sf1, pcDNA3.1-OnGli1, and the indicated cold probe constructs, and luciferase activity was measured 48 hr post-transfection. Competition assays were performed using unlabeled cold probe (Cold probe, GACCACCCA, 10/100/250 ng/mL) or mutant unlabeled cold probe (mutant Cold probe, TTAATTAAA, 10/100/250 ng/mL) (n=4). ‘+’ indicates the addition of the corresponding substance, while ‘-’ indicates no addition, and the number represents the amount added (ng/mL). (F–G) Representative immunofluorescence images of testicular sections from recipient fish transplanted with sf1-deficient (TSL-sf1^-/-, F) or Sf1-overexpressing (TSL-OnSf1, G) SLCs, stained for Cyp11c1 (green) and PKH26 (red). Nuclei are stained with DAPI (blue). Scale bars: 4 μm. (H) Quantification of the percentage of Cyp11c1-positive cells among the transplanted PKH26-positive SLCs in the two transplantation groups (n=5 fish per group). Statistical significance was determined by one-way ANOVA followed by Tukey’s test (E, different letters above the error bar indicate statistical differences at p<0.05) or Student’s t-test (B, H) (*, p<0.05; **, p<0.01; NS, no significant difference).

Figure 5—figure supplement 1. Transcriptome data analyses and verification by RT-qPCR.

(A) Hierarchical clustering analysis of global gene expression patterns in TSL-WT, TSL-OnDhh, TSL-OnGli1, and TSL+0.5 μM SAG. Blue indicates decreased expression, and red indicates increased expression. (B) Heatmap showing the expression patterns of upregulated differentially expressed genes (DEGs) identified in Figure 5C. The FPKM value for each gene in each sample is indicated within the squares. The color gradient from blue to red reflects low to high expression levels per row (gene). (C) qRT-PCR validation. Values were presented as mean ± SD (n=3).

Sf1 is the critical downstream effector of Gli1 in SLCs differentiation

To identify key targets of the Dhh-Gli1 axis, we performed transcriptomic profiling of TSL cells under conditions of pathway activation: Dhh overexpression (TSL-OnDhh), Gli1 overexpression (TSL-OnGli1), and SAG treatment (TSL+SAG). Comparative RNA-seq analysis identified a core set of 33 genes consistently upregulated across all three conditions (Figure 5, Figure 5—figure supplement 1). Among these, sf1 emerged as a prominently upregulated candidate. Functional annotation of its promoter region identified two conserved Gli1-binding motifs, B1 (AACCACCCA) and B2 (GAGCCACCCA) (Figure 5D). A dual-luciferase reporter assay confirmed that Gli1 potently transactivates the sf1 promoter (Figure 5E). The specificity of this interaction was further demonstrated by a cold probe competition assay: a wild-type oligonucleotide containing the Gli-binding motif (GACCACCCA) competitively inhibited Gli1-driven sf1 transactivation in a dose-dependent manner, whereas a mutated version (TTAATTAAA) had no effect (Figure 5E).

The functional necessity of Sf1 was confirmed by transplantation assays using sf1^-/- and Sf1-overexpressing (TSL-OnSf1) TSL cells. While TSL-sf1^-/- cells failed to differentiate into Cyp11c1-positive cells even in WT testes, TSL-OnSf1 cells successfully differentiated and expressed Cyp11c1 even in the dhh^-/- testicular environment (Figure 5F–H). These results definitively position Sf1 as the critical downstream effector executing the Dhh-Ptch2-Gli1 signaling cascade during SLC differentiation.

Discussion

This study delineates a critical Dhh-Ptch2-Gli1-Sf1 signaling axis essential for SLC differentiation in Nile tilapia. By combining dhh^-/- XY models with SLC transplantation, we definitively demonstrate that Dhh signaling is dispensable for Leydig cell lineage survival but indispensable for its differentiation. Furthermore, our integrated evidence from targeted gene knockout and functional assays suggests that Ptch2 acts as the primary receptor for Dhh signaling in SLCs and points to Gli1 as the key transcriptional effector mediating the transactivation of sf1. Collectively, these findings implicate Sf1 as the critical downstream target bridging Dhh signaling to steroidogenic maturation, thereby resolving key ambiguities in Leydig cell lineage development and providing evolutionary insights into vertebrate reproductive biology.

Consistent with its established role in mammals, our data in Nile tilapia confirm that Dhh function in male testis development is evolutionarily conserved across vertebrates. While mutations in Dhh lead to testosterone insufficiency and infertility in mice (Pierucci-Alves et al., 2001) and are associated with 46,XY DSD in humans (Wei et al., 2022), the precise cellular mechanism remained unresolved. Our study demonstrates that dhh^-/- tilapia phenocopy these defects, exhibiting testicular dysgenesis, a profound deficiency of Leydig cells, and drastically reduced 11-KT levels (Figure 1). To define the onset of Leydig cell differentiation, we performed a developmental time-course analysis. This revealed that Cyp11c1-positive steroidogenic cells first appear in wild-type testes at 30 dah, while being conspicuously absent in dhh^-/- mutants at this same stage (Figure 2—figure supplement 1). This clear temporal pattern establishes ~30 dah as the developmental window when SLCs initiate their differentiation program in the Nile tilapia. The functional indispensability of Dhh at this specific stage was further confirmed by rescue experiments, in which both SAG treatment and transplantation of Dhh-activated TSL cells restored Cyp11c1-positive cell populations in dhh^-/- testes (Figure 3). Collectively, our findings position Dhh as a crucial niche signal that acts at a defined developmental checkpoint to drive the differentiation of SLCs into steroidogenic Leydig cells.

The functional divergence of Ptch1 and Ptch2 in Hh signaling represents an intriguing aspect of pathway regulation. Structurally, both receptors share conserved 12-transmembrane domains vital for cholesterol transport but diverge in their cytoplasmic C-terminal regions, which are implicated in differential Smo inhibition efficiency (Fleet and Hamel, 2018; Qi et al., 2018). In mammals, Ptch1 serves as the primary, broad-spectrum regulator of Hh signaling during embryogenesis, whereas Ptch2 often assumes context-dependent roles, such as in primordial germ cell migration via its co-receptor Gas1 (Kim et al., 2020). However, their potential redundancy or specificity within the Leydig cell lineage has remained an open question. Our study provides significant insight into this issue. We confirmed that both ptch1 and ptch2 are expressed in Leydig cells (Figure 4A). Functional interrogation using TSL cell lines revealed a marked difference: genetic ablation of ptch2 profoundly attenuated Dhh-induced signaling, as measured by a Gli-responsive luciferase reporter, whereas loss of ptch1 had no detectable effect under the same conditions (Figure 4B–D). This in vitro data, pointing to a primary role for Ptch2, was further supported by in vivo genetic interaction studies. The observation that the dhh^-/- phenotype was fully rescued in dhh^-/-;ptch2^-/- double mutants (Figure 4E–U) is consistent with Ptch2 acting as the inhibitory receptor whose loss alleviates pathway suppression. The specificity for Ptch2 in this context might stem from unique co-receptor interactions or expression patterns within the testicular niche. To preliminarily assess potential compensatory regulation, we examined ptch1 expression in XY testes from WT, ptch2^-/-, and dhh^-/-;ptch2^-/- fish at 90 dah. No significant differences in ptch1 mRNA levels were detected among these genotypes (Figure 4—figure supplement 4), suggesting that loss of ptch2 does not trigger compensatory upregulation of ptch1 at the transcriptional level under the conditions examined. Nonetheless, global ptch2 mutation affects multiple tissues, whereas our mechanistic focus is on SLC differentiation within the testicular niche. Moreover, the early embryonic lethality of global ptch1 mutation in tilapia (Liu et al., 2024) precludes direct assessment of its role in postnatal testis development. Therefore, although our findings strongly support a predominant role for Ptch2 in mediating Dhh signaling in SLCs, definitive resolution of receptor specificity will require future Leydig cell-specific conditional knockout models.

Gli is a nuclear transcription factor, characterized by the presence of a zinc finger domain located in the middle of the protein (Kinzler et al., 1988). Downstream of the zinc finger domain is a phosphorylation cluster containing phosphorylation sites for protein kinase A and glycogen synthase kinase 3, which negatively regulate the transcriptional activity of Gli through phosphorylation (Niewiadomski et al., 2014). In Drosophila, Ci is the sole Gli homolog, possessing both transcriptional activation and repression functions (Huangfu and Anderson, 2006). In vertebrates, there are three Gli homologs: Gli1, Gli2, and Gli3. Gli1 functions solely as a transcriptional activator because it lacks proteolytic processing domains. Gli2 has both activation and repression functions, while Gli3 primarily acts as a transcriptional repressor (Niewiadomski et al., 2019; Pan and Wang, 2007). However, there has been no research on the regulatory mechanism of Gli1/2/3 in SLC differentiation. In this study, we found that although gli1, gli2, and gli3 are all expressed in Leydig cell lineage (Figure 5A–C), the results from TSL cell lines with mutations in gli1, gli2, and gli3 indicated that TSL-gli1^-/-, rather than TSL-gli2^-/- or TSL-gli3^-/-, failed to respond to Dhh signaling (Figure 5D). This suggests that Gli1 is the key transcription factor downstream of the Dhh signaling cascade in SLCs.

Sf1 and Dhh play important roles in Leydig cell commitment and testicular development (Parker et al., 2002; Zhang and Beachy, 2023). Studies suggest a potential interaction between these two factors. For example, in Dhh^-/- XY mice, testicular development is impaired, with a significant reduction in the number of Leydig cells and a marked downregulation of Sf1 expression (Yao et al., 2002). In XX mouse ovaries where Smo is constitutively activated, a large number of ectopic Sf1-positive Leydig cells are generated (Barsoum et al., 2009). Similarly, in sf1^-/- XY tilapia, testicular development is impaired, with a significant reduction in Leydig cell numbers and downregulation of dhh expression (Xie et al., 2016). These findings imply that Dhh and Sf1 may interact through feedback loops or co-regulatory mechanisms. However, direct evidence for the regulation of Sf1 by the Hh signaling pathway remains lacking. This study elucidates the molecular mechanism of Sf1 as a critical downstream effect of the Dhh-Ptch2-Gli1 axis. Transcriptomic profiling revealed an increased expression of sf1 by Dhh regulation signaling in TSL cells (Figure 5—figure supplement 1), identifying it as a key downstream target of Dhh signaling. Promoter analysis identified two conserved Gli-binding motifs (Figure 5F) within the sf1 promoter, and luciferase assays confirmed that Gli1 transactivates sf1 transcription (Figure 5G). Functional validation demonstrated that TSL-sf1^-/- cells failed to differentiate even in WT testes, whereas TSL-OnSf1 cells restored Leydig cell differentiation and steroidogenesis in dhh^-/- testes (Figure 5H–J). These results establish Sf1 as both necessary and sufficient for Leydig cell lineage differentiation.

In conclusion, our study systematically deciphers the signaling cascade governing SLC differentiation, from the niche signal to the core steroidogenic regulator. We definitively show that the Dhh signal from the niche is indispensable for SLC differentiation, but not their survival. Within this pathway, Ptch2 serves as the critical receptor, relaying the signal through the transcription factor Gli1. The pivotal outcome of this cascade is the activation of sf1 by Gli1. Thus, we establish a complete Dhh-Ptch2-Gli1-Sf1 axis, resolving the long-standing question of how a key morphogen controls Leydig cell lineage development in vertebrates.

Materials and methods

Animals

Nile tilapias (O. niloticus) were kept in recirculating freshwater systems at 26°C under a 12 hr light/dark photoperiod. All animal experiments were conducted in accordance with the regulations of the Guide for Care and Use of Laboratory Animals prescribed by the Committee of Laboratory Animal Experimentation at Southwest University, China (IACUC-20181015-12).

Cell culture

TSL cells were cultured in ESM4 medium (Zhao et al., 2023), which is based on DMEM medium with 20 mM HEPES, 100 U/mL penicillin and streptomycin, 2 mM L-glutamine, 1 mM sodium pyruvate, 2 μM Na-selenite, 1 mM nonessential amino acids, 50 μM 2-mercaptoethanol, 10 ng/mL human recombinant basic fibroblast growth factor (R&D Systems, USA), tilapia embryo extract (1 embryo/mL), 15% fetal bovine serum, and 0.2% rainbow trout serum. HEK293 cells were obtained from ATCC and maintained in Dulbecco’s modified Eagle’s medium (Gibco, USA) supplemented with 10% fetal bovine serum (Gibco, USA) in a humidified incubator at 37°C with 5% CO₂. All cell lines were regularly tested for mycoplasma contamination.

Generation of mutant lines by CRISPR/Cas9 in Nile tilapia

The sequences of tilapia dhh (GenBank ID: 100707022) and ptch2 (GenBank ID: 100692939) were retrieved from the NCBI. CRISPR/Cas9-mediated mutagenesis was performed as described previously (Li et al., 2014). Guide RNAs (gRNAs) targeting the first exon of dhh (GCGGGCCCGGTCCGCATCCC) and ptch2 (GTCCCAGGGGCCGGCGTATT) were designed using ZiFit (http://zifit.partners.org/ZiFiT/). gRNA and Cas9 mRNA were synthesized following established protocols (Li et al., 2014). The primers are listed in Supplementary file 1. A mixture of synthetic gRNA (500 ng/μL) and Cas9 mRNA (1000 ng/μL) was microinjected into one-cell-stage fertilized embryos. Mutants were validated by polyacrylamide gel electrophoresis (PAGE) and Sanger sequencing. F0 chimeric XY males were crossed with WT XX females to generate heterozygous F1 offspring. F1 siblings carrying a 13 bp deletion in dhh or a 25 bp deletion in ptch2 were intercrossed to generate homozygous F2 mutants. For double mutants, dhh^+/- XY males were mated with ptch2^+/- XX females to obtain dhh^+/-;ptch2^+/- offspring. These double heterozygotes were then intercrossed to generate dhh^-/-;ptch2^-/- mutants.

Generation of mutant lines by pCas9-NtU6sgRNA system in TSL cells

The tilapia SLC line TSL was maintained in ESM4 medium as described (Huang et al., 2020). Gene sequences for ptch1 (GenBank ID: 100700194), ptch2 (GenBank ID: 100692939), gli1 (GenBank ID: 100710687), gli2 (GenBank ID: 100704788), gli3 (GenBank ID: 100707331), and sf1 (GenBank ID: 100534561) were retrieved from NCBI. Knockout plasmids targeting these genes were designed following established protocols (Zhang et al., 2023). gRNAs were generated using ZiFit, with the following exon-specific targets: ptch1 (GGAGGCGCTCCTGCAGCACC, exon 3), ptch2 (GTCCCAGGGGCCGGCGTATT, exon 1), gli1 (TGACTCATGGATCAGGACCA, exon 2), gli2 (GATTCGCCTGTCACCCCACG, exon 3), gli3 (TGAGGAGCCCTCTACGTCTA, exon 2), and sf1 (GGCTGTGTACTGGTACTGGG, exon 4; designed as previously described) (Zhang et al., 2023). Primers are listed in Supplementary file 1. Plasmids were transfected into TSL cells using the MDMP E-1 microchannel transfection instrument (MENDGENE) with transwell cell compartments (BIOFIT), as per the manufacturer’s guidelines. Post-transfection (48 hr), cells were selected in ESM4 medium containing 500 μg/mL G418 for 7 days. Single-cell clones were isolated via dilution and serial passage, followed by genomic DNA (gDNA) extraction and Sanger sequencing to confirm mutations.

Histological analyses of testes from WT, dhh^-/-, ptch2^-/-, and dhh^-/-;ptch2^-/- XY fish

GSI was calculated for WT, dhh^-/-, ptch2^-/-, and dhh^-/-; ptch2^-/- XY fish at 90 dah as: (gonad weight/body weight)×100%. Testes were fixed in Bouin’s solution for 24 hr at room temperature with agitation, dehydrated through an ethanol series, and embedded in paraffin. Sections (5 μm thickness) were stained with H&E and imaged using an Olympus BX53 microscope. Germ cell classification (spermatogonia, spermatocytes) followed established criteria (Zhao et al., 2023).

Immunofluorescence

Testes from WT, dhh^-/-, WT+11 KT, dhh^-/-+11 KT, ptch2^-/-, and dhh^-/-; ptch2^-/- XY fish at 90 dah were analyzed for germ cell marker Vasa, Leydig cell marker Cyp11c1, and meiosis marker Sycp3. Following tissue fixation and embedding, sections (5 μm) were deparaffinized, rehydrated, and subjected to antigen retrieval. After blocking with 5% bovine serum albumin (BSA; 30 min, room temperature), sections were incubated overnight at 4°C with rabbit polyclonal primary antibodies (Vasa, Cyp11c1, Sycp3; 1:1000 dilution), which had been validated previously (Dai et al., 2021). Alexa Fluor 488-conjugated goat anti-rabbit secondary antibody (1:1,000; Invitrogen) was applied for 1 hr at 37°C, followed by DAPI nuclear counterstaining (1:1000; Sigma-Aldrich). Fluorescence signals were imaged using an Olympus Fv3000 confocal microscope. For each biological replicate (n=5–6 fish per genotype), three non-serial, non-adjacent testis sections were analyzed. From each section, three representative fields of view were captured to ensure non-overlapping sampling. The numbers of cells positive for Vasa, Sycp3, and Cyp11c1 were quantified using ImageJ Pro 1.51 software with default parameters.

FISH and IF

Testes from WT XY tilapia at 90 dah were fixed in 4% paraformaldehyde (Sigma) overnight at 4°C with agitation, paraffin-embedded, and sectioned (5 μm). FISH was performed as described (Zhao et al., 2023). Partial cDNA fragments of ptch1, ptch2, gli1, gli2, and gli3 were cloned into the pGEM-T Easy vector. Sense (control) and antisense RNA probes were synthesized using a DIG RNA Labeling Kit (Roche) and T7 mMESSAGE mMACHINE Kit (Ambion). Signals were amplified using the TSA Plus TMR system (PerkinElmer) according to the manufacturer’s protocols (primers in Supplementary file 1). Concurrently, Leydig cell marker Cyp11c1 expression was assessed by IF.

11-KT and SAG rescue experiments

To rescue dhh^-/- XY fish, offspring from dhh^+/- XY male × dhh^+/- XX female crosses were divided into two groups. The experimental group received 11-KT via immersion from 30 to 90 dah, with treatments refreshed every 2 days. 11-KT concentrations increased incrementally from 57 ng/L at 30 dah to 1254 ng/L by 90 dah (39.9 ng/L increase per treatment cycle; see Figures 2 and 5, Figure 2—figure supplement 1). For SAG rescue, fish received intraperitoneal injections of 10 mg/kg SAG from 30 to 90 dah, with booster doses administered weekly. Testes were collected at 90 dah for morphological and histological evaluation.

TSL cell labeling, transplantation, and IF

TSL, previously isolated from 3-month-old XY tilapia testes (Huang et al., 2020), was used to assess SLC differentiation in WT and dhh^-/- XY fish. TSL cells were treated as follows: (1) untreated TSL, (2) TSL+0.5 μM SAG (Hh pathway agonist), (3) TSL-OnDhh (overexpressing tilapia Dhh), (4) TSL-OnSf1 (overexpressing tilapia Sf1), and (5) TSL-sf1^-/- (sf1 mutation). For the SAG treatment experiment, TSL cells were incubated with 0.5 μM SAG for 48 hr before transplantation. Cells were labeled with PKH26 fluorescent dye (Sigma) and transplanted (5000 cells per testis) into WT or dhh^-/- XY tilapia via the urogenital tract (Huang et al., 2020). For TSL-OnDhh, the Dhh ORF (1377 bp) was amplified from gDNA, cloned into pcDNA3.1 (Invitrogen), and transfected into TSL cells (primers in Supplementary file 1). At 10 days post-transplantation, testes were analyzed by IF to quantify Cyp11c1-positive cells among PKH26-positive populations.

GLI report assay

A firefly luciferase reporter plasmid (8×GLI) was generated by cloning eight tandem copies of the GLI-binding motif (GACCACCCA) into the pGL4.23 vector (minimal promoter). To validate 8×GLI functionality, TSL cells were transiently transfected with: (1) pGL4.23 (negative control), (2) 8×GLI, or (3) 8×GLI plus Dhh overexpression plasmid (OnDhh), for 48 hr. The 8×GLI construct was further transfected into TSL (WT), TSL-ptch1^-/-, TSL-ptch2^-/-, TSL-gli1^-/-, TSL-gli2^-/-, or TSL-gli3^-/-, with or without OnDhh. Transfections included pRL-TK as an internal control. Luciferase activity was measured 48 hr post-transfection using the Dual Luciferase Assay System (Promega) and a Luminoskan Ascent luminometer (Thermo Fisher Scientific). Relative activity was calculated as firefly/Renilla luciferase ratios.

Western blot

The ORFs of Dhh (1377 bp) and Gli1 (4404 bp) were amplified from gDNA by PCR. Two tandem FLAG tag sequences (GATTATAAAGATGATGATAAA) were inserted flanking each ORF (after the start codon and before the stop codon) and cloned into the pcDNA3.1 vector (Invitrogen), generating pcDNA3.1-OnDhh and pcDNA3.1-OnGli1 (primers in Supplementary file 1). TSL cells were transiently transfected with pcDNA3.1-OnDhh or pcDNA3.1-OnGli1 for 48 hr. Protein expression was analyzed by western blot following established methods (Dai et al., 2021). Briefly, total protein lysates were separated by 15% SDS-PAGE, transferred to nitrocellulose membranes, and blocked with 5% BSA in TBST (10 mM Tris pH 7.9, 150 mM NaCl, 0.1% Tween-20) for 1 hr at 37°C. Membranes were incubated overnight at 4°C with rabbit anti-FLAG antibody (1:1000; Cell Signaling Technology), followed by HRP-conjugated goat anti-rabbit secondary antibody (1:1000; Invitrogen) for 1 hr at 37°C. Signals were detected using the BeyoECL Plus Kit (Beyotime) and imaged on a Fusion FX7 system (Vilber Lourmat).

Measurement of 11-KT levels by ELISA

Blood was collected from the caudal vein of 90 dah XY fish (WT, dhh^-/-, ptch2^-/-, dhh;ptch2^-/-, WT+TSL, dhh^-/-+TSL, dhh^-/-+TSL+SAG, dhh^-/-+TSL-OnDhh, and dhh^-/-+TSL-sf1^-/-), incubated at 4°C overnight, and centrifuged to isolate serum. Tissue fluid samples from WT and dhh^-/- XY fish at 5, 10, 20, and 30 dah were similarly prepared. All samples were stored at −80°C until analysis. Serum and tissue fluid 11-KT levels were quantified using a 11-keto Testosterone ELISA Kit (Cayman) validated for teleost samples, as per the manufacturer’s instructions. Individual samples (serum or tissue fluid; n=6 per genotype) were analyzed in triplicate on a single 96-well plate, with concentrations calculated from standard curves.

Transcriptome analyses

Tissue- and stage-specific expression of dhh was assessed using transcriptomic data from tilapia tissues (brain, heart, head kidney, kidney, liver, muscle, ovary, testis) and gonads at developmental stages spanning 5–360 dah (Tao et al., 2018; Tao et al., 2013). For global gene expression profiling, RNA was extracted from TSL-WT, TSL-OnDhh, TSL-OnGli1, and TSL+0.5 μM SAG cells using RNAiso Plus (Takara). For the SAG treatment experiment, TSL cells were incubated with 0.5 μM SAG for 48 hr before collection. For each genotype, cells from three independent culture wells were pooled. Libraries were sequenced on an Illumina platform (GENEBOOK Biotechnology), and clean reads were aligned to the reference genome with HISAT2. Gene expression levels (FPKM: fragments per kilobase per million mapped reads) were quantified using featureCounts. Differentially expressed genes were identified for each condition (TSL-OnDhh, TSL-OnGli1, TSL+SAG) compared to TSL-WT controls using edgeR (threshold: FDR<0.05, |log2(foldchange)|≥1.5).

Real-time PCR

Testes from WT and dhh^-/- XY fish at 90 dah (n=3 per genotype), along with TSL-WT, TSL-OnDhh, TSL-OnGli1, and TSL+0.5 μM SAG cells, were analyzed for gene expression. Total RNA was extracted using RNAiso Plus (Takara), reverse-transcribed into cDNA (PrimeScript II 1st Strand cDNA Synthesis Kit; Takara), and amplified by real-time PCR on an ABI-7500 system (Applied Biosystems). Relative mRNA levels were normalized to β-actin and calculated via the formula R=2^−ΔΔCt (primers in Supplementary file 1).

Dual luciferase report assay

The 5015 bp promoter region upstream of the sf1 start codon was amplified and cloned into the pGL3-basic vector (Promega), generating pGL3-sf1 (primers in Supplementary file 1). To assess Gli1 binding to the sf1 promoter, HEK293 cells were co-transfected with pGL3-sf1, pcDNA3.1-OnGli1, cold probe (GACCACCCA), and mutant cold probe (TTAATTAAA) using the TransIT-X2 transfection reagent (Mirus Bio). Control transfections included empty pcDNA3.1 and pGL3 vectors, with pRL-TK for normalization. Luciferase activity was measured 48 hr post-transfection.

Statistical analyses

Data are presented as mean ± SD from at least three independent replicates. A two-tailed independent Student’s t-test was used to determine the differences between the two groups. One-way ANOVA, followed by Tukey’s multiple comparison, was used to determine the significance of differences in more than two groups. p<0.05 was used as a threshold for statistically significant differences.

Funding Statement

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

Funding Information

This paper was supported by the following grants:

• National Key Research and Development Program of China 2022YFD1201600 to Deshou Wang.

• National Natural Science Foundation of China 32172969 to Jing Wei.

• National Natural Science Foundation of China 31972776 to Jing Wei.

• National Natural Science Foundation of China 32102780 to Wenjing Tao.

• National Natural Science Foundation of China 32473159 to Xiangyan Dai.

Data availability

All data generated or analyzed during this study are included in the manuscript and its supporting files. Materials generated in this study are available from the corresponding author upon reasonable request.