Human follicle dermal papilla cells (PromoCell, Heidelberg, Germany) and human dermal microvascular endothelial cells (PromoCell) were cultured in a follicle dermal papilla cell growth medium and endothelial cell growth medium (both from PromoCell), respectively. The cells were incubated at 37 °C in a humidified atmosphere containing 5% CO₂.

Recombinant human Ang1, Ang2, and VEGF were purchased from R&D Systems (Minneapolis, MA, USA). MTT (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide), Wortmannin, PD98059, SB202190, anti-Tie-1, anti-Tie-2, Gly-Arg-Gly-Asp-Ser (GRGDS) peptide, and functional blocking antibodies against integrins α1–6, αv, β1, and α5β1 were purchased from Millipore-Sigma (St. Louis, MO, USA). Other reagents and antibodies used were: anti-cleaved caspase-3, anti-Bax, anti-Bcl-2, anti-Bcl-xL, anti-phospho-AKT, anti-AKT, anti-phospho-ERK1/2, anti-ERK1/2, anti-phospho-p38, anti-p38, anti-integrin α4, anti-integrin α5, anti-integrin αv, anti-integrin β1, anti-integrin β3, anti-integrin β4, and anti-integrin β5 (Cell signaling Technology, Danvers, MA, USA), anti-β-tubulin and peroxidase-conjugated secondary antibodies (Santa Cruz Biotechnology, Dallas, TX, USA), Muse® Annexin V & Dead Cell Assay Kit (FITC) (Luminex Corporation, Austin, TX, USA), and 5′-bromodeoxy-2′-uridine (BrdU) cell proliferation ELISAs (Roche, Indianapolis, IN, USA).

Cell viability was measured using the MTT assay kit (Millipore-Sigma). About 5 × 10³ cells were plated in 96-well plates for 24 h and treated with indicated reagents for 48 h. Thereafter, the cells were treated with 100 μL of MTT (5 mg/mL) for 3 h. The formazan levels were measured using the absorbance at 570 nm.

The apoptotic effect was assessed using the Annexin-V-FITC/PI double-staining assay, following the instructions provided by the manufacturer (Muse® Annexin V & Dead Cell Assay Kit). Cells (3 × 10⁵) were treated with the indicated agents for 48 h. Post incubation, cells were collected in 1 mL of medium containing 1% fetal bovine serum (FBS). Subsequently, the cell suspension (100 μL) was mixed with Muse® Annexin V & Dead Cell reagent and vortexed for 5 s. The mixture was then incubated for 20 min at room temperature and analyzed using the Muse™ Cell Analyzer. Data were processed using the Muse Analysis Software, and cells positive for annexin-V only or annexin-V/PI double staining were considered apoptotic. Each experiment was conducted in triplicate.

Cells were harvested and lysed in radioimmunoprecipitation assay (RIPA) buffer (Thermo Fisher Scientific, Waltham, MA, USA) supplemented with a protease and phosphatase inhibitor cocktail (Thermo Fisher Scientific). Protein concentration was determined using the bicinchoninic acid (BCA) assay (Thermo Fisher Scientific), and equal amounts of protein (30 μg per lane) were resolved on 10–12% SDS–polyacrylamide gels. Following electrophoresis, proteins were transferred onto nitrocellulose membranes (0.45 μm pore size; GE Healthcare, Chicago, USA) using a semi-dry blotting system (Bio-Rad, Hercules, CA, USA). Membranes were blocked in 5% non-fat dry milk diluted in TBST buffer (Tris-buffered saline with 0.1% Tween-20) for 1 h at room temperature. Primary antibodies were diluted (1: 1000 in 5% BSA in TBST) and incubated overnight at 4 °C. Following three washes with TBST (10 min each), membranes were incubated with HRP-conjugated anti-rabbit or anti-mouse secondary antibodies (Santa Cruz Biotechnology, diluted 1: 5,000 in 5% non-fat dry milk/TBST) for 1 h at room temperature. Bands were visualized using an enhanced chemiluminescence (ECL) detection reagent (Thermo Fisher Scientific) and imaged using the ImageQuant LAS 500 system (GE Healthcare).

To measure cell proliferation, a Cell Proliferation BrdU ELISA kit (Roche) was used according to the manufacturer’s protocol. Cells treated with the indicated agents for 48 h were labeled with 10 μM BrdU for 1 h. The anti-BrdU peroxidase conjugated antibody was then incubated for 90 min. After washing, the bound peroxidase was detected based on the substrate reaction, which was measured at 450 nm.

All RNA was extracted from cells and tissues using the RNeasy Plus Mini kit (Qiagen). cDNAs were generated from RNAs (1 μg) using 2.5 μM oligo-dT primers, 1 mM dNTPs, and MuLV reverse transcriptase. qRT-PCR assays were performed in the qPCR Master Mix for SYBR Green PCR Master Mix (Applied Biosystems). qRT-PCR was performed using the following primers: TIE2 (forward: 5’-GCTTGCTCCTTTCTGGAACTGT-3′ and reverse: 5′- CGCCACCCAGAGGCAAT-3′); TIE1 (forward: 5’-AGAACCTAGCCTCCAAGATT-3′ and reverse: 5’-ACTGTAGTTCAGGGACTCAA-3′); ITGA4 (forward: 5’-GCTTCTCAGATCTGCTCGTG-3′ and reverse: 5’-GTCACTTCCAACGAGGTTTG-3′); ITGA5 (forward: 5’-TGCAGTGTGAGGCTGTGTACA-3′ and reverse: 5’-GTGGCCACCTGACGCTCT-3′); ITGAV (forward: 5’-AATCTTCCAATTGAGGATATCAC-3′ and reverse: 5’-AAAACAGCCAGTAGCAACAAT-3′); ITGB1 (forward: 5’-GAAGGGTTGCCCTCCAGA-3′ and reverse: 5’-GCTTGAGCTTCTCTGCTGTT-3′); ITGB3 (forward: 5’-CCGTGACGAGATTGAGTCA-3′ and reverse: 5’-AGGATGGACTTTCCACTAGAA-3′); ITGB4 (forward: 5’-AGACGAGATGTTCAGGGACC-3′ and reverse: 5’-GGTCTCCTCTGTGATTTGGAA-3′); ITGB5 (forward: 5’-GGAGCCAGAGTGTGGAAACA-3′ and reverse: 5’-GAAACTTTGCAAACTCCCTC-3′); and ACTB (forward: 5’-GGGAAATCGTGCGTGACATT-3′ and reverse: 5’-AGTTTCGTGGATGCCACAGG-3′). A mean quantity was estimated from triplicate qRT-PCR reactions following normalization to the control gene.

Statistical analyses were performed using the GraphPad Prism software (GraphPad. Inc., La Jolla, CA, USA). Depending on the experimental design, unpaired two-tailed Student’s t-test (assuming unequal variances), one-way analysis of variance, or two-way analysis of variance followed by Tukey’s post-hoc test was used. A p-value of less than 0.05 was considered statistically significant. All quantitative data are presented as the mean ± standard deviation.

Initially, an MTT assay was conducted to assess whether angiogenic factors, including Ang1 and Ang2, influence the viability of HFDPCs. When HFDPCs were treated with Ang1 for 48 h, similar to VEGF, it enhanced cell viability, whereas Ang2 had no effect (Figure 1A).

Next, to accurately assess whether the enhancement in cell viability induced by Ang1 was associated with cell survival or proliferation, Muse® Annexin V & Dead Cell Assay Kit-based cytometric analysis, western blot analysis, and the BrdU cell proliferation ELISA assay were conducted. Ang1 inhibited DHT-induced apoptosis in HFDPCs (Figure 1B). Similarly, Ang1 prevented DHT-induced increases in cleaved caspase-3 and proapoptotic Bax levels, as well as DHT-induced decreases in antiapoptotic Bcl-2 levels in HFDPCs (Figures 1C,D). However, neither ang1 nor DHT affected antiapoptotic Bcl-xL levels in HFDPCs (Figures 1C,D). In addition, Ang1 increased the proliferation of HFDPCs even when treated alone and prevented the decrease in HFDPCs proliferation caused by DHT (Figure 1E). These results demonstrate that Ang1 prevents DHT-induced increase in apoptosis and decrease in proliferation of HFDPCs.

Next, the mechanisms through which Ang1 promotes survival and proliferation in HFDPCs were explored. Ang1 is widely recognized for activating AKT, ERK1/2, and p38 in endothelial cells (37, 38), with these signaling pathways playing a key role in survival or proliferation (37, 39). Based on this, it was hypothesized that Ang1 may similarly regulate survival or proliferation in HFDPCs through these pathways.

Ang1 enhanced the phosphorylation of AKT, ERK1/2, and p38 in HFDPCs following treatment for 15, 30, and 60 min (Figure 2A). When treated with DHT, HFDPCs exhibited reduced phosphorylation of AKT and ERK1/2, while p38 phosphorylation remained unchanged (Figures 2B,C). Furthermore, Ang1 inhibited the DHT-induced reduction in AKT and ERK1/2 phosphorylation in HFDPCs (Figures 2B,C). To investigate whether Ang1-induced activation of AKT, ERK1/2, and p38 plays a role in apoptosis or proliferation, the AKT inhibitor Wortmannin, the ERK1/2 inhibitor PD98059, and the p38 inhibitor SB202190 were utilized. Wortmannin blocked Ang1-induced AKT phosphorylation in HFDPCs with no impact on the phosphorylation of ERK1/2 or p38 (Supplementary Figure 1A). PD98059 blocked Ang1-induced ERK1/2 phosphorylation with no impact on the phosphorylation of AKT or p38 (Supplementary Figure 1A), while SB202190 blocked Ang1-induced p38 phosphorylation with no impact on the phosphorylation of AKT or ERK1/2 (Supplementary Figure 1B). Interestingly, Wortmannin fully blocked Ang1-mediated survival under DHT treatment in HFDPCs, whereas PD98059 and SB202190 had no effect (Figure 2D). Furthermore, Wortmannin fully prevented the Ang1-induced reduction in cleaved caspase-3 and pro-apoptotic Bax levels, as well as the increase in anti-apoptotic Bcl-2 levels under DHT treatment in HFDPCs (Figures 2E,F). In contrast, PD98059 fully inhibited Ang1-driven proliferation in HFDPCs, while neither Wortmannin nor SB202190 contributed to proliferation. (Figure 2G). These findings indicate that Ang1 promotes survival via the AKT pathway and drives proliferation through the ERK1/2 pathway in HFDPCs.

Western blot and qRT-PCR were subsequently performed to examine the presence of Tie-2 and integrins, which serve as receptors for Ang1, in HFDPCs. Notably, since Tie-2 is specifically and highly expressed in endothelial cells (40), HDMECs, a type of endothelial cell, were utilized. Interestingly, neither Tie-1 nor Tie-2, both members of the Tie receptor family, were expressed in HFDPCs (Figures 3A,B). Furthermore, the mRNA expression of Tie-2 and Tie-1 was nearly undetectable in HFDPCs compared to HDMECs (Supplementary Figure 2A). In contrast, integrin α4, α5, αv, β1, β3, β4, and β5 were prominently expressed in HFDPCs, with their mRNA levels also being significantly detectable (Figures 3C,D and Supplementary Figure 2B). These findings indicate that the Tie-2 receptor is absent in HFDPCs, while integrins are present.

Since Tie-2 is absent in HFDPCs and multiple integrins are expressed, it was hypothesized that Ang1 contributes to survival and proliferation via these integrins, with an aim to identify the specific integrin involved. To determine whether Ang1 promotes the survival and proliferation of HFDPCs through integrins, the Gly-Arg-Gly-Asp-Ser (GRGDS) peptide was utilized to inhibit integrins that recognize the Arg-Gly-Asp (RGD) sequence. GRGDS inhibited the activation of Akt and ERK1/2, which contribute to Ang1-induced survival and proliferation in HFDPCs, while also reducing p38 activation. (Figure 4A). Next, various α integrin neutralizing antibodies were used to identify the specific integrin involved in Ang1-induced survival and proliferation in HFDPCs. Among the tested neutralizing antibodies, only integrin α5 inhibited Ang1-induced phosphorylation of AKT, ERK1/2, and p38 (Figure 4B). Since integrin α5 can form a heterodimer with integrin β1, a neutralizing antibody for integrin β1 and an integrin α5β1 neutralizing antibody were used to investigate whether Ang1 promotes survival and proliferation through integrin α5β1 in HFDPCs. The neutralizing antibodies for integrin β1 and integrin α5β1 inhibited Ang1-induced phosphorylation of AKT, ERK1/2, and p38 in HFDPCs (Figure 4C). Furthermore, the integrin α5β1 neutralizing antibody completely blocked Ang1-mediated survival under DHT treatment in HFDPCs (Figure 4D). Additionally, it fully prevented the Ang1-induced decrease in cleaved caspase-3 and pro-apoptotic Bax levels, while also inhibiting the increase in anti-apoptotic Bcl-2 levels under DHT treatment in HFPDCs (Figure 4E and Supplementary Figure 3A). The integrin α5β1 neutralizing antibody also fully inhibited Ang1-mediated proliferation under DHT treatment in HFPDCs (Figure 4F). These results indicate that Ang1 mediates survival and proliferation in HFDPCs through integrin α5β1.