Regulatory Effects of FGF9 on Dermal Papilla Cell Proliferation in Small-Tailed Han Sheep

Fibroblast growth factor 9 (FGF9) is crucial for the growth and development of hair follicles (HFs); however, its role in sheep wool growth is unknown. Here, we clarified the role of FGF9 in HF growth in the small-tailed Han sheep by quantifying FGF9 expression in skin tissue sections collected at different periods. Moreover, we evaluated the effects of FGF9 protein supplementation on hair shaft growth in vitro and FGF9 knockdown on cultured dermal papilla cells (DPCs). The relationship between FGF9 and the Wnt/β-catenin signaling pathway was examined, and the underlying mechanisms of FGF9-mediated DPC proliferation were investigated. The results show that FGF9 expression varies throughout the HF cycle and participates in wool growth. The proliferation rate and cell cycle of FGF9-treated DPCs substantially increase compared to that of the control group, and the mRNA and protein expression of CTNNB1, a Wnt/β-catenin signaling pathway marker gene, is considerably lower than that in the control group. The opposite occurs in FGF9-knockdown DPCs. Moreover, other signaling pathways are enriched in the FGF9-treated group. In conclusion, FGF9 accelerates the proliferation and cell cycle of DPCs and may regulate HF growth and development through the Wnt/β-catenin signaling pathway.


Introduction
Small-tailed Han sheep have bright coloured hair with a fine and soft texture, making it a good raw material for leather. However, compared with that of Merinos, its wool production performance needs to be improved [1]. Wool is an essential production material and is one of the most economically important traits of sheep. HFs form the basis of wool development, and their physiological activities determine the growth and physical properties of their wool product. The density, fineness, and curvature of hair are primarily determined by HF function. HFs are the only mammalian mini-organs with a lifelong periodic cycle that regenerate by spontaneously undergoing repeated anagen, catagen, and telogen phases [2]. Although HFs are highly regulated by various growth factors, cytokines, neuropeptides, and hormones, the periodic HF cycle is a self-sustaining phenomenon that can persist even when HFs are isolated from organ cultures. Despite the identification of multiple molecular regulators in mutant mouse models with defective HF cycles and characteristic gene expression patterns in different phases, the molecular mechanisms that drive the HF cycle remain unclear [3]. DP, which are found at the base of the HF, serve as regulatory centres and are essential for the formation, development, and regulation of HFs and their cycles [4,5]. They are crucial for morphogenesis and controlling the periodic regeneration of HFs. Only DPCs constitute the DP and are mainly distributed at the base, encircled by hair matrix cells [4]. Numerous growth factors and intercellular signaling molecules, such as IGF-1, KGF, and HGF, are expressed and secreted by DPCs and act on neighbouring cells to control cell proliferation and differentiation in HFs, as well as hair shaft elongation [5]. When in vitro DPC cultures were transplanted into nude mice, the mice grew many HFs with visible hair; if DPCs were reduced, HFs failed to form complete structures [6][7][8]. Dermal papilla cells can also regulate the size and length of hair shafts by inducing the growth and differentiation of hair matrix cells [9]. In vitro cultures show that DP can also aid in HF regeneration. Moreover, DPCs do not only induce epithelial cells to form HFs during the embryonic period, but also regulate the periodic regeneration of mature HFs after birth [10,11]. Signaling pathways and cytokines that regulate the developmental and biological characteristics of the DP are expressed in DPCs, and the number and type of DPCs can determine the size and shape of the hair [11,12]. Research on DP is crucial to understand HF formation and hair growth; therefore, we conducted experiments with DPCs at the core.
The FGF gene family, which is widely expressed in skin HF tissues, plays an important role in the cell proliferation, differentiation, and the periodic growth cycle of HFs. It also participates in many other biological processes in mammals by binding to FGFR and cross-synergising with signaling pathways such as neurogenic NOTCH, WNT, and TGF-β [13][14][15]. Kawano et al. conducted a quantitative analysis of the mRNA expression of 22 FGF family members and four FGFRs in the skin of adult mice at different stages of the hair growth cycle [16]. They found that FGF1, FGF2, FGF5, FGF7, FGF10, FGF13, FGF18, and FGF22 were expressed at different stages of the HF proliferation cycle and that their expression peaks varied. The mRNA expression of FGF18 and FGF13 peaked during the telogen phase, that of FGF7 and FGF10 peaked during anagen phase V, and that of FGF5 and FGF22 peaked during anagen phase VI. Housley et al. demonstrate that FGF5 inhibits hair growth by promoting the transition from the anagen to the catagen phase in HFs [17]. FGF9 participates in promoting the formation of dermal agglutinates during the early developmental stage of HFs [18,19]. Studies show that human peripheral blood γ delta (γδ) T cells can express FGF9 in response to stimulation by isopentenyl pyrophosphate and TGF-β1/interleukin (IL)-15, play important roles in skin tissue injury recovery, and indirectly promote the regeneration of HFs [20]. Komi-Kuramochi et al. [21] and Zheng et al. [22] reported similar results regarding the crucial roles of FGF9 in wound healing.
In vitro and in vivo studies suggest that numerous cytokines of the FGF family have regulatory effects on the growth and development of HFs, among which mutations in FGF5 can maintain a prolonged anagen phase in the HFs of multiple species, resulting in a long-hair phenotype [23]. FGF5 gene-edited sheep produced using CRISPR/Cas9 technology by Zhang et al. showed a significant increase in hair length and average growth rate [24]. Moreover, the FGF21 gene regulates the proliferation and differentiation of fibroblasts and the formation of HFs during the first growth cycle by promoting the transition from the anagen to the catagen phase, acting mainly during the anagen phase [25]. Kimura-Ueki et al. [26] found that FGF18 is expressed in HF stem cells throughout the telogen phase by conditionally knocking out the FGF18 gene in keratin-5-positive epithelial cells of genetically engineered mice, revealing that FGF18 regulates the hair cycle mainly through the non-growth phase. Shu et al. [27] showed that during their first growth cycle, FGF7 and FGF10 might induce HFs to enter a new cycle, whereas FGF22 might play an important role in inducing HFs to enter the catagen phase. However, studies also show that FGF7 is necessary for hair growth but not wound healing, whereas FGF10 is necessary for embryonic epidermal morphogenesis but not development [28,29]. Therefore, members of the FGF family act differently during different HF growth stages and maintain normal HF growth and development. However, there is no direct evidence that FGF9 regulates the growth of sheep wool. The FGF9 gene might participate in determining the composition of sheep HF traits; however, its exact regulatory mechanism remains unclear.
Due to the crucial roles of both DPCs and FGF9, we designed this study to gain an overview of the cellular functions of FGF9 in DPCs and further elucidate the roles of FGF9 in HF growth and development. Therefore, we hypothesised that FGF9 might participate in HF growth and development by regulating cell processes in DPCs. We explored the effects of FGF9 on wool growth by culturing HFs of small-tailed Han sheep in vitro and examining the changes in FGF9 gene expression. To speculate the potential functions of FGF9 as a candidate gene for the growth and development of HFs, we further investigated the roles of FGF9 by examining the effects of additional FGF9 protein treatment and FGF9 knockdown on the proliferation and cell cycle of DPCs. Lastly, transcriptome analysis was performed to gain new insights into the molecular mechanisms through which FGF9 affects the function of DPCs. This study provides a scientific basis for subsequent studies on wool growth and development in sheep, thereby laying the foundation for exploring the genetic mechanisms underlying the composition of different wool traits.

Sample Collection
Three two-year-old female small-tailed Han sheep were used in this study. These sheep have no oestrus, pregnancy, or lactation period, and their wool production, stretch length, and curl are all lower than Merino sheep. Therefore, there remains room for improvement in wool production performance [1]. Their outer layer of wool was removed by localised shearing of the scapulae, whereby shears were applied close to their dorsal lateral skin. The skin was cleaned of any remaining fleece. Then, the skin was sterilised by wiping with 75% alcohol, and a 0.5 cm 2 sample of skin tissue containing different HF cycles was collected using sterile surgical shears. Prior to sample collection, the sheep were locally anaesthetised using brucine hydrochloride. The sheep were raised by the Jilin Academy of Agricultural Sciences. Skin samples were collected in January, April, and July of 2022. The research protocol was reviewed and approved by the Animal Welfare and Ethics Committee of Jilin Academy of Agricultural Sciences (AWEC2021A03, 28 May 2021).

Sectioning and Preparation of Skin Tissues
Fresh skin tissue samples were trimmed, fixed in 4% paraformaldehyde (Boster, Wuhan, China) for 48 h, and rinsed in running water for 24 h. After dehydration via immersion in 70%, 75%, 85%, 95%, and 100% alcohol for 1 h each, the samples were soaked in xylene for 20 min and then in wax for approximately 6 h for embedding. The samples were then longitudinally cut in the direction of the HFs, stained with haematoxylin and eosin (HE; Dingguo, Shanghai, China), and sealed with neutral resin. The tissues used for RNA extraction were immersed according to the manufacturer's instructions in RNAlater preservation solution (Thermo Fisher Scientific, Waltham, MA, USA), and the tissues used for DPC extraction were immediately sent to the laboratory (Institute of Animal Biotechnology, Jilin Academy of Agricultural Sciences, Gongzhuling, Jilin, China) for cell isolation and culture.

Isolation and Culture of DPCs
DPCs were isolated using the methods described by Kobayashi et al. [30] and Topouzi et al. [31], which involved blunt dissection combined with enzymatic digestion. Briefly, the scapular skin of small-tailed Han sheep was digested with neutral protease at 4 • C for 4 h. HFs in the anagen phase with intact DP were selected, and the top of the hair bulbs were cut off under a microscope (Nikon eclipse Ti-s, Tokyo, Japan; 100× objective). The DPCs were then removed and transferred to a Dulbecco's modified Eagle medium/nutrient mixture F-12 (DMEM/F12) (Gibco, Grand Island, NY, USA) medium with 10% fetal bovine serum (FBS) (Sigma, St. Louis, MO, USA) and 1× antibiotic and antifungal agents (Sangon Biotech, Shanghai, China) using mouth pipetting. All cells were cultured at a constant temperature of 37 • C with 5% carbon dioxide (CO 2 ) for approximately 7 days.

Exogenous Addition of FGF9 Protein and Inhibition of FGF9 Gene Expression
Recombinant human FGF9 (Takara Bio, Beijing, China) was dissolved in sterile PBS containing 0.1% BSA according to the manufacturer's instructions to prepare a stock solution at a concentration of 100 µg/mL. Thereafter, it was diluted to the target concentrations in DMEM/F12 (Gibco) containing 10% FBS (Sigma), according to the different dose requirements.
Three siRNA sequences, siRNA-FGF9-1, siRNA-FGF9-2, and siRNA-FGF9-3 were designed according to the mRNA sequence of the sheep FGF21 gene published in GenBank (accession number: XM_027977590.1) and transfected into the DPCs using Lipofectamine 3000 (Thermo Fisher Scientific). The total RNA of the cells was extracted 48 h after transfection, and the transfection efficiency was tested using real-time fluorescence-based qPCR to screen for the best siRNA-FGF9 combination for subsequent experiments. All siRNA sequences were constructed by Jintuosi Biotechnology Co., Ltd., Wuhan, China, and their sequences are listed in Table 1. Table 1. Small interfering RNA (siRNA)-fibroblast growth factor 9 (FGF9) sequences used in this study.

Effects of FGF9 on HF Growth In Vitro
HFs were isolated using the method described by Higgins et al. [22] and cultured in vitro under the following conditions: William's E serum-free culture medium (Gibco) with 2 mmol/L L-glutamine (Sangon Biotech), 2 mmol/L HEPES (Sigma), 10 µg/mL insulin (Sigma), 10 ng/mL sodium selenite (Aikeshiji, Chengdu, China), 10 ng/mL hydrocortisone (Sigma), 10 µg/mL transferrin (Aladdin, Shanghai, China), 100 U/mL penicillin (Sangon Biotech), and 100 µg/mL streptomycin (Sangon Biotech) at 31 • C with 5% CO 2 and saturated humidity. The treated HF group was additionally administered with 100 ng/mL FGF9. Eight intact HFs in the anagen phase were extracted from each group and placed in 24-well plates for in vitro culture. The HF length (the total height of the HFs containing new fibres minus the initial height) was measured daily (in millimetres), and the growth rate was plotted using GraphPad Prism software (version 7.0, GraphPad Prism software, Inc., La Jolla, CA, USA).

Determination of Cell Proliferation Rate and Cell Cycle
The isolated and cultured DPCs of small-tailed Han sheep were uniformly inoculated into 96-well plates at 100 µL/well (containing approximately 2000 cells). The original culture medium was then replaced with complete culture medium containing 0, 10, 100, and 1000 ng/mL FGF9 protein after 24 h of inoculation. After 24, 48, 72, and 96 h, 10 µL CCK-8 solution was added to each well. The absorbance of each well at 450 nm was measured using a microplate reader after 2 h of incubation at 37 • C, and cell proliferation curves were plotted through GraphPad Prism 7.0 software.
The subsequent experimental conditions were set as 100 ng/mL FGF9 treatment for 48 h. To further examine the facilitatory effect of FGF9 on DPC proliferation, we evaluated the changes in the number of DPCs in the proliferative phase using a 5-ethynyl-2 deoxyuridine (EdU) assay. After exogenous addition of FGF9 protein and transfection, DPC proliferation was evaluated using a Cell-Light EdU Apollo567 In Vitro Kit (RiboBio, Guangzhou, China), according to the manufacturer's instructions. At 48 h after transfection, the cell cycle of the DPCs was examined using flow cytometry with the Cell Cycle and Apoptosis Analysis Kit (Beyotime, Shanghai, China) according to the manufacturer's recommendations.

RNA Extraction, Library Construction, and High-Throughput Sequencing
A 1 mL volume of TRIzol (Invitrogen) was added to each treatment group. The bottom cells were repeatedly aspirated to form a cell suspension, which was transferred to a centrifuge tube. Then, 200 µL chloroform was added, and the tube was shaken to obtain a pink turbid solution. The solution was allowed to rest for 10 min at 4 • C before centrifugation for 15 min at 12,000× g. The upper colourless, aqueous phase was transferred to another centrifuge tube. Isopropanol (500 µL) was added, and the solution was mixed thoroughly, left to rest for 10 min at 4 • C, and then centrifuged at 12,000× g for 10 min. The supernatant was removed and discarded. Next, 1 mL 75% ethanol was added, and the tube was then shaken to resuspend the RNA precipitate at 4 • C. The suspension was centrifuged at 7500× g for 5 min, after which the supernatant was discarded and the precipitate was dried at room temperature in an ultra-clean table. Next, 20 µL diethyl pyrocarbonate water was added to fully dissolve the precipitate to obtain the RNA samples. The first strand cDNA was prepared using a PrimeScript™ RT Reagent Kit with gDNA Eraser (Takara Bio) according to the manufacturer's instructions. The RNAs and cDNAs were stored at −80 • C and −20 • C, respectively, for later use. The quality of the RNA was measured using a Quawell Q5000 spectrophotometer (Quawell, San Jose, CA, USA) and RNA integrity was confirmed through agarose gel electrophoresis. The mRNAs were enriched using magnetic beads containing oligo-deoxythymidine (dT) and broken into short fragments by adding a fragmentation buffer. Subsequently, double-stranded cDNA was synthesised via reverse transcription using random hexamer primers. AMPure XP (Beckman Coulter, Beverly, MA, USA) beads were used to purify the double-stranded cDNAs and select fragments based on size, which were then amplified using PCR to construct a cDNA library. RNA was quantified using Agilent 2100 Bioanalyzer (Agilent Technologies, Santa Clara, CA, USA). After the library passed the quality control analysis, transcriptional sequencing was performed using an Illumina HiSeqTM2500/4000 sequencing platform (Allwegene, Beijing, China).

Bioinformatics Analysis
Raw sequencing data were filtered using Trimmomatic software (version 0.33). Cleaned data were mapped onto the reference genome of Ovis aries using STRA software (Reference Genome Version ARS-UI_Ramb_version 2.0 [NCBI]). Gene expression in each sample was analysed using the HTSeq software (version 0.5.4) in combination with a union model. The fragments per kilobase of exon model per million mapped reads (FPKM) value was used as a measurement of the gene expression level. A |log 2 (fold change)| > 1 and p < 0.05 were used as the criteria to identify differentially expressed genes (DEGs).

qPCR
Differences in the expression of FGF9 in skin tissues containing HFs in the anagen, catagen, and telogen phases were also tested using qPCR. ACTB was used as the internal control. After the exogenous addition of FGF9 protein and inhibition of FGF9 gene expression in DPCs, several key genes of the Wnt/β-catenin signalling pathway, including CTNNB1, GSK3β, LEF1, DVL2, LRP5, and LRP6, were selected for qPCR quantification. The first strand cDNAs were created after the total RNA was extracted. Using the Primer Premier (version 5.0) program, primers were created following the assembly sequences of the investigated genes in GenBank (Table 2).

Gene Ontology (GO) and Kyoto Encyclopedia of Genes and Genomes (KEGG) Pathway Enrichment Analyses
To gain insight into the biological relationship between the DEGs of DPCs in the FGF9 protein-supplemented and control groups, we set the significance threshold (p < 0.05) and used cluster analysis software to perform GO (GOSeq, topGO, v1.22) and KEGG (KOBAS, v2.0) pathway enrichment analyses on the DEGs. GO was divided into three categories: molecular function, biological processes, and cellular composition.

Statistical Analysis
The results of the real-time PCR analysis were processed using the 2 −∆∆CT method. Statistical analyses were performed using SPSS 22.0 software. The independent sample T-test and one-way analysis of variance (ANOVA), as well as LSD, Tukey's-b, and Waller-Duncan tests, were used for variance analysis and significance tests. Data analysis was performed using GraphPad Prism 7.0 software. All experimental data are expressed as the mean ± standard error of the mean (SEM), and statistical significance was set at p < 0.05.

HF Cycle Phases and FGF9 Expression in Relation to Sheep Wool Growth
The HF samples extracted in July 2022 were in the anagen phase, in which the DP consisted of fibroblasts embedded in the mesenchyme of connective tissues by the epithelial cells of the bulb. In January 2022, the HFs were in the catagen phase, in which the DPs were located at the proximal ends of the HFs and the DPCs were arranged closely together with minimal ECM. In April 2022, the HFs were in the telogen phase, in which the DP had reduced mesenchyme, the epithelial cells of the bulb shrank, and the DP was released (Figure 1a). These results confirm that the HFs in the skin of small-tailed Han sheep follow the HF cycle.

FGF9 Promotes DPC Proliferation in Small-Tailed Han Sheep
The CCK-8 assay results reveal that the addition of exogenous FGF9 protein facilitates DPC proliferation (Figure 2a). We observed an increase in the number of EdUpositive cells in the proliferative phase following the addition of exogenous FGF9 protein, and a statistical analysis of the number of EdU-positive cells as a proportion of the total number of cells does show an upward trend (Figure 2b,c). Cell cycle analysis  Figure S1). The qPCR results show that FGF9 expression changes throughout the HF cycle. Compared with that in the telogen phase where DP is released, the expression level of FGF9 is higher in the skin tissues, where HFs are in the growth stage. It is speculated that the expression level of FGF9 in the skin tissues may be related to the periodic growth cycle of hair follicles (Figure 1a,b). To evaluate the effects of FGF9 on wool growth, HFs of small-tailed Han sheep were cultured in vitro using the organ culture method with the addition of the FGF9 protein. Hair length markedly increases with the addition of FGF9 protein compared to that in the control group (Figure 1c,d). These results demonstrate that FGF9 participates in hair growth in small-tailed Han sheep.

FGF9 Promotes DPC Proliferation in Small-Tailed Han Sheep
The CCK-8 assay results reveal that the addition of exogenous FGF9 protein facilitates DPC proliferation (Figure 2a). We observed an increase in the number of EdU-positive cells in the proliferative phase following the addition of exogenous FGF9 protein, and a statistical analysis of the number of EdU-positive cells as a proportion of the total number of cells does show an upward trend (Figure 2b,c). Cell cycle analysis shows that, compared to the control group, the number of cells in the S phase is higher after the addition of exogenous

FGF9 Regulates the Expression of Key Wnt/β-Catenin Signaling Pathway Genes
We examined the effects of FGF9 on the mRNA expression of key genes of the Wnt/βcatenin signaling pathway, including CTNNB1, GSK3β, LEF1, DVL2, LRP5, and LRP6, as well as that of β-catenin, a marker protein of the Wnt/β-catenin signaling pathway. After the exogenous addition of FGF9 protein to DPCs, the mRNA expression levels of CTNNB1, GSK3β, DVL2, and LRP6 decrease, with a significant decrease observed in that of CTNNB1 and GSK3β, whereas the expression levels of LEF1 and LRP5 increase (Figure 4a). Although there is no significant difference between them, compared with the control group, the mRNA expressions of CTNNB1, GSK3β, DVL2, and LRP6 in the FGF9 inhibited group show an upregulated trend, while the mRNA expressions of LEF1 and LRP5 show a downward trend (Figure 4b). Western blotting shows similar results; the protein expression levels of β-catenin decrease after the addition of FGF9 protein but increase after the inhibition of FGF9 expression (Figure 4c,d). Our findings show that the proliferation-promoting effects of FGF9 on DPCs may be regulated via the Wnt/β-catenin signaling pathway.  The qPCR results show that siRNA-FGF9-3 has the highest transfection efficiency a 20 nM (Figure 3a). Dermal cell papilla at the growth stage are transfected with siRNA FGF9-3 (hereinafter called siRNA-FGF9). The CCK-8 assay results show that ce proliferation is reduced after the inhibition of FGF9 expression compared with that i analysis of the number of EdU-positive cells as a proportion of the total number of cell does show a decreasing trend (Figure 3c,d). Cell cycle assay results show a significan decrease in the number of cells in the S phase after FGF9 inhibition and an increase i the number of cells in the G0/G1 phase compared to those in the control (Figure 3e,f).  and LRP6 in the FGF9 inhibited group show an upregulated trend, while the mR expressions of LEF1 and LRP5 show a downward trend (Figure 4b). Western blott shows similar results; the protein expression levels of β-catenin decrease after addition of FGF9 protein but increase after the inhibition of FGF9 expression (Fig  4c,d). Our findings show that the proliferation-promoting effects of FGF9 on DPCs m be regulated via the Wnt/β-catenin signaling pathway.

Functional Enrichment Analysis of DEGs
To gain a comprehensive understanding of how FGF9 regulates DPC proliferation, we performed transcriptome analysis on DPCs in the FGF9 protein-supplemented and control groups. The changes in the differential expression of genes between the two groups were analysed using a volcano plot. A total of 629 DEGs are identified, of which 385 genes are downregulated and 244 genes are upregulated (Figure 5a). and control groups, with the most enriched category being biosynthesis-chondroitin sulphate/dermatan sulphate, followed by m advanced glycation end-product (AGE-RAGE) signaling path complications, and ECM receptor interactions. The top 20 most pathways were selected in the order of their p-values and plotted (Fig   Figure 5. (a) Volcano plots of differential gene expression between DPCs tre h and the blank group (green: downregulated genes; red: upregulated gen Gene Ontology (GO) enrichment in the biological processes (blue) and (yellow) categories. The significance level of the enrichment is set to an adj 0.05). (c) Scatterplot for the Kyoto Encyclopedia of Genes and Genomes (K differential genes. The bigger the circle, the more genes there are. The re smaller the Q−value, and the closer the Q−value number is to zero, the m enriched. (c) Scatterplot for the Kyoto Encyclopedia of Genes and Genomes (KEGG) enrichment of differential genes. The bigger the circle, the more genes there are. The redder the colour, the smaller the Q−value, and the closer the Q−value number is to zero, the more it is significantly enriched.
From the KEGG analysis, most single genes are associated with biological regulation and developmental processes. In the cellular components category, most single genes are associated with the ECM. The top 30 most significant GO terms are not enriched in the molecular function category (Figure 5b). The KEGG pathway enrichment analysis was then performed on the significant DEGs screened from the FGF9-treated and control groups, with the most enriched category being glycosaminoglycan biosynthesis-chondroitin sulphate/dermatan sulphate, followed by malaria, the roles of advanced glycation endproduct (AGE-RAGE) signaling pathways in diabetic complications, and ECM receptor interactions. The top 20 most significant KEGG pathways were selected in the order of their p-values and plotted (Figure 5c).

Discussion
The results indicate that FGF9 can accelerate the proliferation and cell cycle of DPCs and regulate HF growth and development through the Wnt/β-catenin signaling pathway. FGF9 expression also varies throughout the HF cycle and participates in wool growth in small-tailed Han sheep. Similarly, previous studies indicate that FGF5, which also belongs to the FGF family, can affect the density of sheep wool by regulating the Wnt/β-catenin signaling pathway [32].
Sectioned skin tissues of small-tailed Han sheep are found to follow the HF cycle, showing regular anagen, catagen, and telogen phases. Additionally, the expression levels of FGF9 varies according to the HF cycle. Similarly, the expression of FGF7, FGF5, FGF10, and FGF22 is also found to peak during the anagen phase in mice [16]. The cell proliferation rate and cell cycle are accelerated in DPCs treated with exogenous FGF9 protein. Simultaneously, the proliferation rate and cell cycle are inhibited in FGF9-downregulated DPCs. This is consistent with the results of Biggs et al. [33] and Gay et al. [19], who show that FGF9 promotes cellular proliferation and HF regeneration. The results of the in vitro HF culture confirm the facilitating effect of FGF9 on hair shaft elongation. Likewise, Cai et al. [34] applied oil produced by FGF9-transgenic safflower seeds to the skin and wounds of mice and found that the hair growth and wound healing rates were higher in the treatment group than those in the control group.
Some previous studies partially explained the association between FGF family members and mechanisms related to the Wnt/β-catenin signaling pathway. Notably, FGFs and the Wnt signaling pathway interact within the DP to regulate the expression of various molecules, including Wnt agonists (R-spondins) and antagonists (DKK2 and Notum) [35]. β-catenin deficiency leads to the degeneration and inflammation of HFs [36]. However, a forced expression of the constitutively active form of β-catenin in the DP does not impair the hair cycle [37]. These results suggest that, while Wnt signaling activity in the DP is essential for maintaining growth, it is not sufficient to counteract hair-promoter-induced signals. Herein, we found that the mRNA and protein expression of a marker gene of the Wnt/β-catenin signaling pathway, CTNNB1, is significantly lower (p < 0.05) in DPCs supplemented with FGF9 protein compared to the control group, while it shows an upregulation trend in DPCs with FGF9 downregulation. Therefore, FGF9 may regulate the growth and development of HFs through the Wnt/β-catenin signaling pathway. Similarly, Enshell-Seijffers et al. [38] show that β-catenin activity in the DP regulates signaling pathways, including that of FGF, thereby mediating the inducing effects of the DP. In conjunction with subsequent experiments, we hypothesised that the positive effect of FGF9 on the proliferation of DPCs may be synergistic through multiple signaling pathways. The signaling cascade in which FGF9 acts in DPCs may act antagonistically with Wnt/β-catenin, and competitive inhibition leads to a decrease in CTNNB1 expression.
The synergistic involvement of multiple signaling pathways in DPC proliferation has been established, and studies show that FGF signaling crosses and acts in concert with the NOTCH, WNT, and TGF-β signaling pathways to control cell maturation [12,13]. Owing to the insignificant changes in the mRNA expression of LEF1, DVL2, LRP5, and LRP6, we speculated that FGF9 might regulate DPC proliferation through other signaling pathways. Therefore, we performed transcriptomic analysis of DPCs in the FGF9 protein-treated and control groups. Through GO and KEGG pathway enrichment analyses of the DEGs, we identified the genes most associated with FGF9 expression that regulate the biological processes and cellular composition during DPC proliferation. In the cellular composition category, most single genes are enriched in the ECM, which is consistent with the fact that FGF9 exerts its biological functions primarily through a paracrine mechanism of action [39].
In the biological processes category, the DEGS are involved in the regulation of biological and developmental processes. Several DEGs are enriched in various signaling pathways, such as TNF, cGMP-PKG, relaxin, FoxO, and PI3K-Akt, suggesting that FGF9 may play a role in promoting cell cycle transition and regulating cell metabolism. By analysing the sequencing results, we speculate that FGF9 synergistically regulates DPC proliferation through multiple signaling pathways. However, the mechanisms of interaction between FGF9 and various signaling pathways still need to be further explored.
In conclusion, our study shows that FGF9 is expressed in varying extents in smalltailed Han sheep and that the addition of exogenous FGF9 protein can encourage DPC proliferation and cell cycle progression. Furthermore, FGF9 may regulate HF growth and development through the Wnt/β-catenin signaling pathway. Lastly, the positive effects of FGF9 on DPC proliferation and the cell cycle may be the result of the synergistic effects of multiple signaling pathways. These findings provide the basis for further production of genetically modified sheep that produce more and better wool. Future research will focus on the molecular mechanism through which FGF9 promotes DPC proliferation to elucidate how FGF9 induces hair follicle growth and development through multiple signaling pathways.

Data Availability Statement:
The data presented in this study are available upon request from the corresponding author.