Skin aging mainly depends on ultraviolet (UV) irradiation, called photoaging, rather than intrinsic aging, unlike other organs (Fisher et al., 2002; Rittié and Fisher, 2015; Zhao et al., 2023). Chronic exposure of the human skin to UV irradiation increases the expression levels of various matrix metalloproteinases (MMPs), which degrade collagen fibrils and other dermal extracellular matrix (ECM). This molecular phenomenon eventually leads to wrinkle formation and the loss of skin elasticity in photodamaged human skin (Lee et al., 2008; Han et al., 2014).

UV irradiation causes the generation of hydrogen peroxides and other reactive oxygen species (ROS) and decreases antioxidant enzymes, which alter gene and protein structure and function, leading to skin damage (de Jager et al., 2017; Papaccio et al., 2022). These features have been observed in chronologically aged human skin. UV irradiation stimulates the signal transduction pathway of mitogen-activated protein kinases (MAPKs), including the extracellular signal-regulated kinase (ERK), Jun N-terminal kinase (JNK) and p38 (Xu et al., 2014; Limtrakul et al., 2016; Kim et al., 2018b; Oh et al., 2020). In addition, UV exposure significantly reduces the transforming growth factor-β (TGF-β), which contributes to increased collagen synthesis (Gambichler et al., 2007; Han et al., 2022).

Pinitol, a 3-methoxy analog of D-chiro-inositol, is a naturally occurring compound found in fine wood, derived from legumes such as soy and carob (Ceratonia siliqua). Pinitol has been reported to possess insulin-like effects and is used as an anti-diabetic agent, driving creatine and other nutrients into muscle cells (Bates et al., 2000; Nascimento et al., 2006). Pinitol is structurally related to the phosphatidylinositol phosphates, which participate in insulin signaling pathways that regulate glucose transport (Yap et al., 2007; Dang et al., 2010). Pinitol significantly downregulates plasma glucose concentration in streptozotocin-induced diabetic rats and enhances glucose utilization in insulin-resistant animals (Sivakumar et al., 2010). Recently, pinitol intake has been reported to increase the lifespan of Drosophila melanogaster (Hada et al., 2013). However, the effect of pinitol on UV-induced skin damage remains unknown.

In this study, we investigated whether pinitol ameliorates ultraviolet A-induced damaged fibroblasts. We found that pinitol significantly increased the number of collagen fibers. In addition, pinitol significantly reversed the UVA-induced phosphorylation levels of ERK and JNK and specifically enhanced the TGF-β signaling pathway for collagen synthesis.

Reagents

Pinitol (3-O-Methyl-D-chiro-inositol) was derived from carob bean (Ceratonia siliqua).

Wound healing analysis

HDFs were seeded in a 60 mm dish at 2×10⁵ cell density. Subsequently, HDFs were incubated at 37°C in a 5% CO₂ incubator for 24 h to reach confluence. Then, a straight line was scratched across the confluent cell in the middle of the dish using a 200 μL pipette tip. After removing the culture medium, DPBS was added and exposed to UVA radiation at 10 J/cm². Immediately after UVA radiation, DPBS was removed and the culture medium was added to the HDFs. HDFs were treated with pinitol 10 μM and incubated for 24 h at 37°C in a CO₂ incubator. To compare 0 h and 24 h, all dishes are photographed using a microscope (Nikon Corporation, Tokyo, Japan).

The closure area of the wound was calculated as follows:

Migration area (%)=(A₀-A_n)/A₀ X 100

(A₀: the area of the initial, A_n: the area of the remaining)

Preparation of human dermal equivalents

Normal human dermal fibroblasts (NHDFs) (Lonza, Basel, Switzerland) were cultured in Dulbecco’s Modified Eagle’s Medium (DMEM) (Lonza) with 10% fetal bovine serum (FBS; Lonza), 100 μg/mL of streptomycin, and 100 U/mL of penicillin in a 5% CO₂ incubator at 37°C. NHDFs were used for experiments in passages 3 through 8. Human dermal equivalents (HDEs) were prepared using type I collagen (BD Biosciences, Bedford, MA, USA). NHDFs were mixed with neutralized collagen in DMEM at a density of 2.5×10⁵ cells/mL. The final concentration of type I collagen was 1 mg/mL. The NHDFs-collagen mixture was dispensed onto 6-well tissue culture plates (Nunc, Rochester, NY, USA) in aliquots of 2 mL per well and allowed to polymerize for 1.5 h at 37°C, after which 2 mL of DMEM containing 10% FBS was added to each well.

Treatment of human dermal equivalents and human skin equivalents

The EpidermFT-400 full-thickness human skin equivalents (HSEs), normal human 3D skin tissue, which were generated by keratinocytes that make up the epidermis and fibroblasts that make up the dermis layer of the skin were purchased from MatTek Corporation (Ashland, MA, USA). The skin tissues were placed in 6-well plates with 2 mL of EFT-400-MM medium and incubated overnight at 37°C/5% CO₂ conditions. After overnight incubation, the medium was replaced with a fresh culture medium.

Before UVA irradiation, tissues were washed two times with DMEM (Phenol red-free; Lonza). HSEs and HDEs were irradiated with 10 J/cm² of UVA using a BIO-SUN (Vilber Lourmat Deutschland GmbH, Eberhardzell, Germany). After irradiation, DMEM was aspirated and exposed with DMEM containing 1 or 10 μM of pinitol.

Total RNA extraction and real-time polymerase chain reaction (RT-PCR)

For analysis of mRNA by RT-PCR, total RNA was extracted using Trizol (Invitrogen, Carlsbad, CA, USA) following the manufacturer’s protocol. Two micrograms of total RNA were reverse transcribed to cDNA using ReverTra Ace reverse transcriptase (Toyobo, Osaka, Japan). All values were normalized to human GAPDH (43333764F) (Applied Biosystems, Foster City, CA, USA). Real-time RT-PCR was performed on a 7500 Fast Real-time PCR system (Applied Biosystems). TaqMan^® Gene Expression Assays were purchased from Applied Biosystems. cDNA samples were analyzed to determine the expression of COL1A1, Hs00164004_m1; MMP1, Hs00899658_m1; TIMP1, Hs00171558_m1 and PEPD, Hs00165445_m1.

Transmission electron microscopy

For conventional transmission electron microscopy, skin equivalents were fixed in Karnovsky fixative, followed by further fixation in 2% osmium tetroxide in a sym-collidine buffer. After staining with uranyl acetate, specimens were dehydrated in ethanol and embedded in Embed-812 (Electron Microscopy Sciences, Hatfield, PA, USA) at 60°C for 48 h. Ultrathin sections (approximately 70-90 nm) were stained with uranyl acetate and lead citrate and were then observed with a transmission electron microscope (TEM; JEM-1200 EX, JEOL, Peabody, MA, USA). TEM images were recorded on negative film and transferred into a computer using a scanner (EPSON Perfection V700 PHOTO™, Long Beach, CA, USA) at 1200 dpi.

Histochemical analyses

Tissue samples from each condition were fixed for 24 h in 10% neutral buffered formalin (Sigma-Aldrich, St. Louis, MO, USA). The tissue blocks were cut in serial sections of 3 μm, which were then stained with hematoxylin and eosin (H&E), Masson`s trichrome, and immunohistochemistry for MMP1 and collagen I by routine techniques then examined by light microscopy (BX41, Olympus, Tokyo, Japan).

Briefly, slides were deparaffinized, rehydrated, and incubated with proteinase K solution for 10 min. After the washing procedure with double-distilled water, slides were covered for 10 min with 3% H₂O₂ to block endogenous peroxidase activity, followed by an additional washing procedure with PBS. Slides were then placed in a humid chamber and incubated for 2 h with the primary rabbit anti-MMP1 (ab38929) (Abcam, Cambridge, UK) or collagen, type I antibody (ab21285) (Abcam). After three rinses in washing buffer, the slides were incubated with the HRP-conjugated donkey anti-rabbit antibody (ab6802) (Abcam) for 1 h. Tissue staining was visualized with a DAB substrate chromogen solution. Slides were counterstained with hematoxylin, dehydrated, and mounted.

Western blot analysis

Cells were isolated from human dermal equivalent (HDEs) by using bacterial collagenase (Sigma-Aldrich) and lysed with RIPA buffer (Millipore, Billerica, MA, USA) supplemented with phosphatase inhibitors (Sigma-Aldrich) and proteinase inhibitors (Roche-applied-sciences, Basel, Switzerland). Fifteen micrograms of protein samples were separated by 4-12 % Bis-Tris gels (Life Technologies, Carlsbad, CA, USA), transferred to PVDF membrane (Roche-applied-sciences), and analyzed by western blotting. Levels of glyceraldehyde 3-phosphate dehydrogenase (GAPDH) were determined in the same samples as controls using rabbit polyclonal anti-GAPDH antibody (Santa Cruz Biotechnology, Dallas, TX, USA). The level of phospho- and total mitogen-activated protein kinases (MAPKs), c-Jun, and Smad3 were detected using a rabbit polyclonal anti-phospho-JNK antibody (Cell signaling technology, Danvers, MA, USA), rabbit polyclonal anti-JNK antibody (Cell signaling technology), rabbit polyclonal anti-phospho-ERK antibody (Cell signaling technology), rabbit polyclonal anti-ERK antibody (Cell signaling technology), rabbit polyclonal anti-phospho-p38 antibody (Cell signaling technology), rabbit polyclonal anti-p38 antibody (Cell signaling technology), rabbit polyclonal anti-phospho-c-Jun antibody (Cell signaling technology), rabbit polyclonal anti-c-Jun antibody (Cell signaling technology), rabbit monoclonal anti-phospho-Smad3 antibody (Cell signaling technology), and rabbit polyclonal anti-Smad3 antibody (Novus biological, Littleton, CO, USA).

Measurement of H₂O₂ released from HDFs

Secreted H₂O₂ levels from cells were measured using an Amplex red hydrogen peroxide/peroxidase assay kit (Life Technologies), according to the manufacturer’s instructions. Briefly, 50 μL of culture media were harvested immediately after UVA irradiation, and incubated with 100 μL of reaction mixtures containing 100 μM Amplex red reagent and 0.2 U/mL horseradish peroxidase for 10 min. The fluorescence of each sample was measured at 590 nm emission following excitation at 560 nm using a Gemini XPS microplate reader (Molecular Device, Sunnyvale, CA, USA).

Statistical analysis

All of the data are presented as the mean ± SD. Significant differences between treatment groups were identified using a t-test. The p-values less than 0.05 were considered statistically significant.

Pinitol enhanced the migration of UVA-damaged HDFs

To investigate the wound healing efficacy of pinitol on the migration of HDFs, a wound healing assay was performed. HDFs were scratched and irradiated with UVA (10 J/cm²), and treated with 10 μM pinitol. After 24 h, pinitol treatment significantly improved the wound healing capabilities in UVA-damaged HDFs (Fig. 1). When converted into a graphical representation, the migration extent indicated that pinitol could restore wound healing in HDFs damaged by UVA (Fig. 1).

Figure 1. Pinitol antagonized UVA-induced collagen-related genes in the normal human equivalents (HDEs). The HDEs were irradiated with UVA (10 J/cm²) and treated with either 1 or 10 μM pinitol for 24 h. Total RNAs were extracted, and quantitative real-time RT-PCR was performed to detect the collagen I (COL1A1), MMP1, TIMP1, and prolidase (PEPD) levels. The graphs are shown as the means ± SD of three independent experiments. *p<0.01.

Pinitol antagonized UVA-induced damaged collagen-related genes in the dermal equivalents (DEs)

To investigate whether pinitol ameliorates UVA-induced damage in HDEs, we treated DEs with pinitol (1 or 10 μM) for 24 h after UVA (10 J/cm²) irradiation and analyzed the mRNA expression levels of collagen type I, tissue inhibitor of metalloproteinases 1 (TIMP1), and prolidase, which is one of collagen homeostasis-associated genes that play an essential role in the recycling of proline (Shim et al., 2012). Interestingly, pinitol significantly reversed the UVA-induced down-regulation of collagen type I, TIMP1, and prolidase (Fig. 2). In addition, pinitol significantly antagonized the UVA-induced upregulation of MMP1 (Fig. 2).

Figure 2. Pinitol recovered wound healing in UVA-damaged NHDFs. The HDEs were irradiated with UVA (10 J/cm²), and treated with 10 μM pinitol for 24 h. The image shows the wound healing efficacy. The depicted graph represents the migration area of each sample. ***p<0.001 indicate a significant difference compared to the UVA-irradiated group. All data are expressed as median ± standard deviation (n=3).

Pinitol improved UVA-induced down-regulation of collagen in HSEs

To confirm whether pinitol improved the UVA-induced damages, we irradiated HSEs with UVA (10 J/cm²) and treated these specimens with pinitol (10 μM) for 24 h.

As shown by Masson’s trichrome staining and staining with anti-collagen antibody, the collagen layer of UVA-irradiated HSEs treated with pinitol was thicker than that of the UVA-irradiated HSEs (Fig. 3A). In addition, TEM analysis revealed that pinitol facilitated the maturation of collagen fibrils, as indicated by an increased number of long banded, well-organized collagen fibrils when treated on HSEs or UVA-irradiated HSEs (Fig. 3B). These findings suggest that pinitol treatment qualitatively improves collagen fibers.

Figure 3. Pinitol improved UVA-induced down-regulation of collagen in human skin equivalents (HSEs). Each HSEs was treated with pinitol, ultraviolet A, or pinitol after ultraviolet A irradiation. (A) Each sample was stained with hematoxylin (red) and eosin (blue) (H&E), Masson’s trichrome, and immunohistochemically stained against MMP1 or collagen I, and then examined by light microscopy. (B) Ultrastructural analysis using reconstructed HSEs has revealed the effect of pinitol on dermal collagen. The bottom four panels show structural details of the boxed areas. Scale bars, 1 μm.

Pinitol antagonized UVA-induced ERK/JNK activation through scavenging H₂O₂

UV generally induces ROS levels in cells (de Jager et al., 2017). Thus, we investigated whether pinitol could prevent ROS generation. As expected, UVA irradiation significantly increased extracellular H₂O₂ secretion, but was decreased by pinitol treatment (Fig. 4). UVA irradiation also induces the activity of MAPKs, including p38, ERK, and JNK (Oh et al., 2020). Once activated, MAPKs phosphorylate c-Jun and lead to an increase in the transcription factor AP-1, which induces both the downregulation of procollagen I and the upregulation of MMPs (Hong et al., 2015; Bang and Choung, 2020). Therefore, we evaluated whether pinitol could prevent UVA-induced MAPK activation. UVA irradiation increased the phosphorylation of ERK and JNK but not p38. However, pinitol treatment prevented the activation of ERK and JNK. Simultaneously, the UVA irradiation-induced increase in c-Jun phosphorylation was inhibited by pinitol (Fig. 5). These results indicate that pinitol interrupts the signaling pathways involved in UVA-induced photodamage.

Figure 4. Pinitol relieved UVA-induced H₂O₂ generation. NHDFs were pre-treated with pinitol (10 μM) and irradiated with UVA (10 J/cm²). The extracellular H₂O₂ release of each sample was measured by Amplex red hydrogen peroxide/peroxidase assay kit. Data shown are the means ± SD of results from 3 independent experiments. **p<0.01.

Figure 5. Pinitol antagonized UVA-induced ERK/JNK activation through scavenging H₂O₂. The HDEs were irradiated with UVA (10 J/cm²) and then treated with pinitol (10 μM) for 2 h or 4 h. The cell lysates were analyzed with immunoblotting using anti-ERK, anti-phosphorylated ERK, anti-p38, anti-phosphorylated p38, anti-JNK, anti-phophorylated JNK, anti-c-jun, anti-phophorylated c-jun, or GAPDH. The results are the representative images of three independent experiments.

Pinitol enhanced the phosphorylation of Smad 3

TGF-β plays a critical role in regulating multiple cellular responses in wound healing processes (Werner et al., 2007; Hameedaldeen et al., 2014). TGF-β/Smad pathway impairment has been reported to reduce type I procollagen expression in dermal fibroblasts (Kim et al., 2008; Gao et al, 2018). TGF-β downregulates the expression of specific enzymes related to the breakdown of collagen, such as MMP-1 and MMP-3 (Liu et al., 2018; Heo et al., 2021). UV exposure decreases type I procollagen expression via the inhibition of the TGF-β signaling pathway such as the upregulation of Smad7, an inhibitory Smad (Kim et al., 2018a; Jin et al., 2021). Thus, we further investigated whether pinitol affects the TGF-β/Smad signaling pathway. Interestingly, pinitol specifically increased the phosphorylation level of Smad3 (Fig. 6). In addition, the phosphorylation level of Smad2 and the expression levels of Smad4 and Smad7 were unaffected by pinitol (Fig. 6). We found that both UVA and pinitol did not alter significantly the mRNA expression of TGF-β1, TGF-β2, TGF-β receptor 1 or TGF-β receptor 2 (Supplementary Fig. 1).

Figure 6. Pinitol enhanced the phosphorylation of Smad 3. The HDEs were irradiated with UVA (10 J/cm²) and subsequently treated with pinitol for 24 h. The phosphorylation or total of each Smad isoform was detected with corresponding antibodies. The results show representative images of three independent experiments.

In this study, we demonstrated that pinitol exerted anti-aging effects on UVA-induced damage of HDEs and HSEs. Interestingly, pinitol antagonized several types of damage caused by UVA irradiation, including collagen fiber degradation and increased MMP-1 expression (Fig. 3). As its underlying mechanism, pinitol reversed the UVA-induced phosphorylation levels of ERK and JNK and enhanced Smad3 phosphorylation (Fig. 5, 6).

A previous report demonstrated that pinitol treatment on Drosophila melanogaster significantly enhances the nuclear localization of dFOXO, an anti-aging target, by specifically activating dJNK and dS6K, which extends the life span of Drosophila melanogaster (Hada et al., 2013). However, no changes in S6K caused by pinitol treatment were detected in this study (data not shown), implying that the molecular mechanism underlying pinitol may differ among species.

We observed that pinitol significantly reduced the UVA-induced generation of H₂O₂ (Fig. 4). However, the precise mechanism underlying this effect require further investigation. Pinitol reverses defective endothelial function by decreasing ROS, key molecules in diabetes related to endothelial dysfunction (Nascimento et al., 2006). Other studies have demonstrated that pinitol can prevent diabetes mellitus by effectively decreasing hyperglycemia and hypertriglyceridemia in streptozotocin-induced type 2 diabetic animals (Bates et al., 2000; Nascimento et al., 2006). Pinitol reduces the postprandial blood glucose level and stimulates GLUT4 translocation in skeletal muscle (Dang et al., 2010).

A previous study reported that Tsumura-Suzuki obese diabetic mice showed thinner collagen bundles, decreased the collagen fiber density, and skin tensile strength compared to non-obese mice (Ibuki et al., 2012). This skin fragility is associated with oxidative stress and MMP overexpression in the subcutaneous adipose tissue, which may partially influence dermal fragility via a paracrine pathway. Several diabetic patients suffer from cutaneous pathologies such as necrobiosis lipoidica diabeticorum, which is characteristic of collagen degeneration and cutaneous atrophy (Oikarinen et al., 1987; Nern, 2002; Ahmed and Goldstein, 2006). In particular, chronic diabetic patients suffer from diabetic foot ulcers, and in certain cases, the symptoms worsen because of bacterial infection and eventually lead to amputation of the leg (Attinger and Brown, 2012; Yip et al., 2023). Accordingly, we recently observed that pinitol exhibited a wound healing effect in damaged fibroblasts treated with lipopolysaccharide derived from E. coli (data not shown). Therefore, we suggest that pinitol can be topically administered to the skin of diabetic patients because of its ability to increase collagen synthesis.

In conclusion, this study demonstrated that an anti-diabetic agent, pinitol can be a potential therapeutic agent for preventing skin damage or improving UV-induced skin damage.

The authors have declared no conflicting interests.

This work was supported by the National Research Foundation of Korea (NRF) grant funded by the Korea government (MSIT) (2022R1A2C1093305).