Taurine‐based Active Pharmaceutical Ingredient Ionic Liquids Modulate the Perifollicular Microenvironment for Androgenic Alopecia Treatment

ABSTRACT

The pathological process of androgenic alopecia (AGA) is closely associated with oxidative stress in the perifollicular microenvironment. Existing antioxidant small‐molecule drugs are often constrained by low permeability, necessitating the urgent development of active therapeutic agents that can effectively penetrate the skin barrier and precisely deliver antioxidants to hair follicles. This study reported a taurine‐based active pharmaceutical ingredient ionic liquid (CTIL) synthesized via a one‐step method, which not only acted as a delivery carrier but also served as a therapeutic material capable of modulating the perifollicular microenvironment by scavenging reactive oxygen species (ROS). An in vitro study demonstrated that the hydrogen‐bond network structure of CTIL facilitated its penetration through the stratum corneum barrier, thus improving the targeted accumulation of taurine within deep hair follicle structures. More importantly, in the AGA animal model, CTIL effectively eliminated excessive ROS in the perifollicular microenvironment, alleviated tissue oxidative damage and inflammatory infiltration, thereby inhibiting the miniaturization process of hair follicles and promoting hair regrowth. This study provided an innovative strategy integrating delivery and treatment for AGA therapy and revealed the potential of bioactive ionic liquids in regulating the microenvironment of skin diseases.

A taurine‐based active pharmaceutical ingredient ionic liquid (CTIL) is developed for treating androgenic alopecia. CTIL functions simultaneously as a transdermal carrier and a therapeutic agent. It penetrates the skin barrier, scavenges perifollicular reactive oxygen species, alleviates oxidative damage and inflammation, and inhibits hair follicle miniaturization, thereby promoting hair regrowth in an animal model.

1. Introduction

Androgenetic alopecia (AGA) is the most common type of pathological alopecia [1, 2]. It is characterized by progressive miniaturization of hair follicles driven by sustained androgen signaling, resulting in a shortened growth phase and the progressive transformation of terminal hairs into vellus‐like hairs [3, 4]. Although hair loss is not life‐threatening, it can adversely affect an individual's appearance and thereby impair self‐esteem and psychological well‐being, leading to both physical and psychological distress and a reduced quality of life [5, 6]. With the accelerating pace of life and increasing environmental exposure, the prevalence of AGA has been rising and showing a trend toward earlier onset [7].

During the onset and progression of AGA, the aberrant activation of androgen receptor signaling has been reported to enhance the metabolic activity of dermal papilla cells (DPCs), thereby accelerating the mitochondrial respiratory chain reactions within cells and resulting in a marked increase in reactive oxygen species (ROS) production [8, 9]. Excessive ROS accumulation not only directly induces apoptosis of hair follicle‐associated cells but also inhibits the transition from telogen to anagen phase by regulating multiple signaling pathways, ultimately contributing to hair follicle miniaturization [10, 11]. Simultaneously, immune cells such as natural killer (NK) cells, T cells, and neutrophils infiltrate the perifollicular microenvironment and secrete pro‐inflammatory mediators, which further exacerbate follicular damage and impair hair regeneration [12, 13]. In this context, oxidative stress within the hair follicle microenvironment is considered a crucial pathological link connecting androgen signaling, inflammatory responses, and follicular degeneration. Therefore, alleviating oxidative stress and restoring perifollicular microenvironmental homeostasis represents one of the key therapeutic strategies for intervening in AGA.

However, antioxidant‐based therapeutic strategies still face substantial limitations in practical applications. The stratum corneum, as a highly dense biological barrier, markedly restricts the penetration of most antioxidant molecules into the hair follicle region, making it difficult to achieve and maintain therapeutically effective concentrations at the target site. These limitations highlight the need for advanced transdermal delivery systems for AGA treatment [14, 15, 16]. In recent years, ionic liquids (ILs) have garnered increasing attention in the drug delivery field due to their tunable physicochemical properties [17, 18]. ILs are generally defined as organic salts composed of cations and anions with relatively low melting points. Through rational selection of ionic components, their physicochemical characteristics, including solubility, stability, and transmembrane transport behavior, can be precisely regulated [19, 20, 21, 22]. Particularly, active pharmaceutical ingredient ionic liquids (API‐ILs), which are directly formed from pharmacologically active molecules, offer a promising strategy for integrating drug delivery and therapeutic functions within a single system [23, 24]. Unlike conventional formulations that rely on inert carrier materials, API‐ILs can simultaneously serve as both functional delivery media and bioactive therapeutic entities. This carrier‐minimized design offers potential advantages for overcoming complex biological barriers and improving local therapeutic efficacy.

Based on the pathological characteristics and therapeutic requirements of AGA, we developed a choline‐taurine active pharmaceutical ingredient ionic liquid (CTIL) for the treatment of AGA via transdermal administration. Choline, a commonly used component in ionic liquids, has been extensively studied and proven to possess biosafety [25, 26, 27]. Taurine (Tau), an endogenous sulfur‐containing amino acid, possesses multiple biological functions, including antioxidant activity, metabolic regulation, and cytoprotective effects, and has been shown to enhance the resistance of hair follicle‐related cells against oxidative stress [28, 29, 30]. ILs composed of choline and taurine have previously been explored for biomaterial modification applications [31]. In this study, this taurine‐based IL system was further utilized as a bioactive therapeutic platform for the treatment of AGA, thereby extending the biomedical applicability of taurine‐derived ILs in skin disease therapy. We systematically characterized the structural and physicochemical properties of CTIL, evaluated its antioxidant capacity in vitro, investigated its transdermal and follicular delivery performance, and further assessed its therapeutic efficacy in an AGA animal model. The results demonstrated that CTIL not only served as a transdermal delivery medium but also exhibited intrinsic biological activity. By reducing excessive ROS accumulation and alleviating inflammatory responses within the perifollicular microenvironment, CTIL effectively mitigated oxidative stress‐associated follicular damage and promoted hair regeneration. Overall, this study provided a bioactive ionic liquid‐based strategy for transdermal AGA therapy and may offer broader insights into the development of functional materials for skin‐related diseases (Figure 1).

FIGURE 1.

Schematic illustration of active pharmaceutical ingredient ionic liquids CTIL for AGA treatment. (A) The preparation process of CTIL. (B) API‐IL CTIL scavenged ROS and modulated the perifollicular microenvironment for AGA.

2. Results and Discussion

2.1. Synthesis and Characterization of Taurine‐based Active Ionic Liquids

To address the dual challenges of oxidative microenvironment imbalance in hair follicles and the limited efficiency of transdermal delivery in AGA, we designed a bioactive pharmaceutical ingredient ionic liquid with therapeutic functionality. Biocompatible choline was selected as the cationic component, while Tau, a bioactive molecule with antioxidative and cytoprotective properties, was employed as the anionic counterpart to construct a choline‐taurine ionic liquid (CTIL). CTIL was synthesized by a one‐step method through an acid‐base neutralization reaction [23]. The synthetic route is illustrated in Figure 2A, and the resulting CTIL exhibited a moisture content of 3.43% and a Tau content of 51.57%, indicating a relatively active ingredient loading capacity and supporting its potential application as a functional therapeutic material.

FIGURE 2.

Synthesis and physicochemical characterization of CTIL. (A) The synthetic route of CTIL. (B) ¹H‐NMR Spectrum of CTIL. (C) FT‐IR spectra of CTIL and precursor components. (D) Shear viscosity of CTIL. (E) Viscosity and conductivity of CTIL at different amounts of water (n = 3).

Nuclear magnetic resonance (NMR) spectroscopy was employed to characterize the composition and structural features of CTIL. As shown in Figure 2B, the ¹H NMR spectrum exhibited the characteristic methylene signals of Tau at δ 3.03 (m, 4H), along with the methyl and methylene proton signals attributable to choline at δ 3.22 (s, 9H), 3.54 (m, 2H), 4.08 (m, 2H), confirming the coexistence of both ionic components in the final product [32]. The ¹³C NMR (Figure S1) spectrum further verified the successful formation of the choline‐taurine system. Notably, compared with the spectra of the individual components, the proton signals of the choline cation in CTIL displayed slight downfield shifts (Figure S2), accompanied by corresponding shifts in the methylene proton signals of Tau (Figure S3). These chemical shift changes suggested the presence of intermolecular interactions and local electronic environment redistribution within the ionic liquid system, indicating that CTIL was not a simple physical mixture of the two components. Fourier‐transform infrared (FT‐IR) spectroscopy provided additional evidence for the formation of the ionic liquid structure (Figure 2C). In the O‐H/N‐H stretching vibration region, choline hydroxide (ChOH) exhibited a broad absorption band centered at approximately 3350 cm^{− 1} corresponding to O–H stretching vibrations, whereas Tau displayed a relatively sharp N–H stretching band at 3035 cm⁻¹. In contrast, CTIL showed a broadened and shifted adsorption band in this region, indicative of strengthened and diversified hydrogen‐bonding interactions. Moreover, the characteristic S═O stretching vibrations of Tau in the range of 1040–1180 cm^{− 1} exhibited changes in both peak intensity and position in CTIL, reflecting interactions between the sulfonate group and the choline cation. The appearance of new vibrational modes in the fingerprint region of CTIL further supported the formation of a distinct ionic liquid structure. The DSC analysis results (Figure S4) further verified the successful formation of CTIL. Tau exhibited distinct crystallization and melting transitions, in sharp contrast to the amorphous characteristics of choline hydroxide. Notably, the CTIL thermogram displayed a characteristic glass transition temperature (Tg) at −72.68°C without any observable melting transition, indicating the absence of crystalline behavior and confirming the formation of a room‐temperature ionic liquid. In addition, the surface tension of CTIL, determined by the pendant drop method, was measured to be 64.03 mN/m, which was consistent with previously reported values for choline‐based ionic liquids [33, 34].

Viscosity is an important physicochemical parameter for ILs, as it strongly influences molecular mobility, transdermal permeation behavior, and formulation stability [35]. Excessively high viscosity may hinder molecular diffusion and reduce skin permeation efficiency, whereas overly low viscosity may compromise drug solubility and formulation stability [36, 37]. In this study, the viscosity of CTIL was measured by a rotational rheometer. The shear stress‐shear rate curve showed that the viscosity of CTIL was measured to be 1225.3 mPa·s, which reflected the presence of a strong hydrogen‐bonded network within the system (Figure 2D). Upon dilution with water, the viscosity of CTIL decreased significantly (Figure 2E), suggesting that water molecules weakened the interactions between ionic components and thereby enhanced system fluidity. Simultaneously, the conductivity of CTIL gradually increased with increasing water content. The conductivity of neat CTIL was 0.16 mS/cm and increased substantially after water addition. This phenomenon is likely associated with reduced ion‐pair interactions and enhanced ionic mobility in the hydrated state.

2.2. Molecular Mechanism Underlying the Formation of Taurine‐based Active Ionic Liquids

To further uncover the formation mechanism of CTIL, multiscale simulation studies were conducted to investigate these interactions. Quantum chemical calculations revealed that, within a single ion pair, the ─O⁻ group of the Tau anion and the ─OH group of the choline cation formed a stable hydrogen bond, with an O···H distance of 1.55 Å (Figure 3A). Electrostatic potential (ESP) mapping and surface potential distribution analysis (Figure 3B,C) revealed a relatively uniform charge distribution over the ion‐pair surface, with most regions concentrated within low‐to‐medium potential intervals. These results suggested that CTIL possessed moderate polarity, which may contribute to its compatibility with compounds of different polarities.

FIGURE 3.

Molecular simulation of CTIL. (A) Hydrogen bond length and Gibbs free energy change between Tau and choline molecules of a single ion pair. (B) ESP mapping of a single ion pair. (C) ESP distribution of a single ion pair. (D) Snapshots of the synthesis process for CTIL. Purple: hydrogen bond. (E) Interaction energy change of the system during the synthesis process. (F) Lennard‐Jones energy, Coulomb energy, and (G) H‐bound number during the synthesis process in the system.

In order to comprehensively understand the dynamic formation process of CTIL, dissipative particle dynamics (DPD) simulations were performed. As shown in Figure 3D, during the simulation period from 0 to 20 000 ps, Tau and choline gradually evolved from an initially dispersed state into stable aggregated structures, accompanied by a continuous decrease in the system potential energy (Figure S5). Quantitative analysis demonstrated that the interaction energy between choline and Tau progressively increased over time, whereas the interactions between each component and water gradually weakened (Figure 3E,F). Correspondingly, the number of hydrogen bonds between choline and Tau steadily increased, while hydrogen bonding between monomers and water gradually decreased (Figure 3G). These findings collectively indicated that hydrogen‐bonding interactions played important roles in the self‐assembly and stabilization of CTIL. Taken together, these results from both experimental characterization and theoretical simulations confirmed the successful construction of CTIL. The moderate viscosity, tunable ionic interactions, and favorable fluidic properties of CTIL provided physicochemical properties suitable for topical and transdermal applications.

2.3. In Vitro Biocompatibility Evaluation of Taurine‐based Active Ionic Liquids

The clinical translation of functional ionic liquids is closely associated with their biosafety and biocompatibility. Since CTIL was designed as a transdermally administered API‐IL for modulation of the perifollicular oxidative microenvironment, it was necessary to evaluate whether the ionic liquid formulation introduced additional cytotoxicity or hemolytic effects. The cytocompatibility of CTIL was first evaluated in fibroblast cells (L929) and HUVEC using the MTT assay. As shown in Figure 4A, free Tau, choline hydroxide, and their physical mixtures exhibited negligible toxicity over a broad concentration range corresponding to equivalent Tau content, which was consistent with their well‐documented biocompatibility. Notably, CTIL‐treated cells maintained cell viability above 80% across all tested concentrations, indicating that the ionic liquid formulation did not introduce additional cytotoxicity. Similar results were observed in HUVEC (Figure 4B), suggesting favorable cytocompatibility of CTIL toward both fibroblastic and endothelial cell types. Live/dead staining assay was performed to visualize cellular viability. As shown in Figure 4C,D, both L929 and HUVEC cells exhibited predominantly green fluorescence with minimal red signal after treatment, indicating that CTIL does not induce significant acute cytotoxicity and maintains good short‐term cell viability. DPCs, as one of the key regulatory cell populations involved in hair follicle development and cycling, play a crucial role in regulating hair growth [38]. Therefore, the biocompatibility of CTIL toward DPCs was further investigated. The MTT assay results (Figure S6) and live/death staining images (Figure S7) collectively demonstrated that CTIL did not induce significant cytotoxicity toward DPCs within the tested concentration range. Hemocompatibility was assessed using a hemolysis assay. As shown in Figure S8, CTIL exhibited a low hemolysis rate relative to the H₂O positive control, indicating good blood compatibility.

FIGURE 4.

Cytocompatibility of CTIL. Cell viability of (A) L929 and (B) HUVEC treated with different formulations for 24 h (n = 4). Representative live/death staining images of (C) L929 and (D) HUVEC treated with different formulations for 24 h.

In accordance with the OECD guideline [39], C57BL/6 mice were used to evaluate the dermal irritation and corrosion potential of CTIL. PBS and 10% SDS aqueous solution were employed as the negative and positive controls, respectively. As shown in Figure S9, the SDS‐treated group exhibited obvious erythema and edema at 1 h after patch removal, which further progressed to pronounced erythema with partial eschar formation at 72 h, indicating a strong irritant response. In contrast, no visible signs of erythema, edema, eschar formation, or other dermal abnormalities were observed in either the PBS or CTIL‐treated groups throughout the entire observation period. Based on the qualitative evaluation according to OECD criteria, SDS induced a marked skin irritation response, whereas CTIL showed no observable irritation under the tested conditions, suggesting its favorable dermal compatibility and potential suitability for topical application.

Collectively, these results demonstrated that CTIL possessed favorable cytocompatibility, hemocompatibility, and dermal safety. This biocompatibility profile supported its potential application as a functional transdermal API‐IL for localized delivery and modulation of the hair follicle microenvironment in AGA treatment.

2.4. In Vitro Antioxidant Capability of Taurine‐based Active Ionic Liquids

Oxidative stress within the hair follicle microenvironment is a key pathological factor in AGA. To evaluate the intrinsic antioxidant capacity of CTIL, we first employed the ABTS assay. In this system, ABTS is oxidized to form stable ABTS⁺ radical cations with a characteristic blue‐green color, which are reduced in the presence of antioxidants, resulting in a decrease in absorbance. As shown in Figure 5A,B, CTIL exhibited concentration‐dependent ABTS⁺ radical scavenging activity. Detectable antioxidant activity was observed at a concentration of 0.625%, and the scavenging effect increased progressively with concentration, accompanied by a marked reduction in the characteristic absorption peak of ABTS⁺. At 5% concentration, CTIL exhibited over 98% radical scavenging efficiency, accompanied by a visible decolorization of the reaction solution (Figure S10), indicating the intrinsic antioxidant property of Tau within the ionic liquid system. To further evaluate the superoxide anion scavenging ability of CTIL, a pyrogallol autoxidation assay was employed. Under weakly alkaline conditions, pyrogallol undergoes autooxidation to generate •O₂ ⁻ and corresponding colored oxidation intermediates. The rate of pyrogallol autoxidation, reflected by the absorbance at 320 nm, is positively correlated with the generation of •O₂ ⁻ and can therefore be used to evaluate the superoxide scavenging activity of CTIL. As shown in Figure S11, CTIL exhibited a superoxide anion radical scavenging activity comparable to that of an equivalent amount of taurine, indicating that the antioxidant functionality of CTIL is largely retained after ionic liquid formation.

FIGURE 5.

In vitro antioxidant capability of CTIL. (A) UV–vis absorption spectra of ABTS⁺ radical solution after incubation with CTIL at different concentrations. (B) ABTS⁺ radical scavenging efficiency of CTIL at different concentrations (n = 3). (C) Representative fluorescence images of intracellular ROS scavenging by CTIL determined by the DCFH‐DA probe. Scale bar = 300 µm. (D) Quantitative analysis of ROS levels detected by DCFH‐DA assay (n = 3).

The intracellular antioxidant activity of CTIL was further assessed using a cellular oxidative stress model. L929 fibroblasts were exposed to H₂O₂ to induce intracellular ROS accumulation, followed by DCFH‐DA staining for ROS quantification. As shown in Figure 5C,D, H₂O₂ treatment increased intracellular ROS levels by approximately 1.44‐fold compared with untreated controls. Notably, CTIL treatment effectively reduced ROS levels compared with the H₂O₂ group, restoring fluorescence intensity toward control levels. Consistently, similar experiments were further applied to DPCs to validate the antioxidant activity of CTIL in a hair follicle‐relevant cellular model. In agreement with the results obtained in L929 fibroblasts, CTIL significantly attenuated H₂O₂‐induced ROS accumulation in DPCs and maintained intracellular ROS at physiological levels (Figure S12). These results collectively demonstrated that CTIL possessed effective in vitro antioxidant activity, suggesting its potential to alleviate oxidative stress in the hair follicle microenvironment.

2.5. Permeation and Retention in the Skin of Taurine‐based Active Ionic Liquids

As the largest organ of the human body, the skin constitutes a robust defensive barrier that safeguards against pathogenic invasion and environmental damage [16, 40]. Nevertheless, this inherent barrier function concurrently compromises the efficiency of transdermal drug delivery, thereby presenting a critical obstacle to targeted intervention in the follicular microenvironment in disorders such as AGA [15, 19]. Given this understanding, we evaluated the transdermal permeation and intradermal retention ability of CTIL. A Franz diffusion cell system was utilized to simulate physiological skin conditions (Figure 6A), enabling quantitative comparison of the transdermal profiles and intradermal retention of Tau across distinct formulations. As depicted in Figure 6B, CTIL significantly elevated the 24 h cumulative Tau permeation to 11.35 mg/cm², representing an approximately 1.85‐fold enhancement relative to the 6.12 mg/cm² observed in the suspension group. After 24 h of permeation, the quantity of Tau retained in skin tissue was determined. Encouragingly, CTIL also improved intradermal retention, achieving 635.34 µg/g compared with 266.06 µg/g in the suspension (Figure 6C), demonstrating its capacity to maintain therapeutically relevant concentrations of Tau within skin tissue.

FIGURE 6.

Permeation and retention of CTIL in the skin. (A) Schematic diagram of the Franz diffusion cell. (B) Quantification results of skin penetration of Tau (n = 3). (C) Quantification results of skin retention of Tau after 24 h permeation in vitro (n = 3). (D) Fluorescence images and corresponding penetration depth analysis of (E) DiD and (F) RhB following incubation for 24 h. Scale bar = 200 µm.

Next, two fluorescent model probes, DiD (a lipophilic carbocyanine dye) and rhodamine B (RhB, a hydrophilic red dye), were adopted to evaluate the transdermal delivery capability of CTIL. As shown in Figure 6D, DiD in PBS (Free DiD) exhibited strong green fluorescence predominantly localized in the superficial skin layers, with limited penetration into deeper regions. Similarly, RhB solution displayed red fluorescence mainly confined to the stratum corneum, further verifying the restrictive barrier function of the stratum corneum. In contrast, CTIL‐mediated delivery enhanced the skin penetration of both hydrophilic and lipophilic probes, as validated by increased fluorescence intensity in deeper skin layers and enriched signal distribution in hair follicle regions (Figure 6E,F). These findings indicated that CTIL not only enhanced the transdermal delivery efficiency of both hydrophilic and hydrophobic molecules and promote their accumulation within skin appendage structures. The enhanced transdermal penetration of CTIL may be attributed to its unique physicochemical properties. First, the moderate viscosity and low surface tension of CTIL enhanced the wetting and spreading on the skin surface, providing a critical prerequisite for efficient drug penetration. In addition, possible intermolecular interactions between CTIL components and stratum corneum lipids, such as hydrogen bonding and electrostatic interactions, could contribute to disrupting the ordered arrangement of lipids and increasing the fluidity of the stratum corneum, thereby weakening the skin barrier function [41, 42, 43, 44]. Furthermore, CTIL may favor drug partitioning into skin appendage pathways, promoting accumulation within hair follicles [45, 46]. The synergistic effect of these aforementioned mechanisms renders CTIL a safe and efficient carrier for transdermal and hair follicle‐targeted delivery.

2.6. Effect of Taurine‐based Active Ionic Liquids on Hair Regrowth in AGA Mouse Model

Encouraged by the in vitro excellent antioxidant efficacy and skin‐penetrating capacity of CTIL, we further evaluated its in vivo potential to promote hair regrowth. It is well established that in the pathological milieu of AGA, testosterone is converted by 5‐α reductase into dihydrotestosterone (DHT), a metabolite with markedly enhanced biological activity. DHT subsequently triggers its downstream biological responses via binding to androgen receptors, ultimately leading to hair follicle miniaturization and even atrophy [47]. Accordingly, an AGA mouse model was established by intraperitoneal administration of DHT. The experimental design and treatment schedule were illustrated in Figure 7A. Macroscopic observation of dorsal skin pigmentation serves as a reliable indicator of hair cycle progression, since melanin deposition accompanies the transition from telogen to anagen [48]. Regular photography of the treatment areas of mice in each group revealed that on day 12, all groups exhibited varying degrees of hair regrowth (Figure 7B). Specifically, the control group exhibited a gradual color transition from pink to gray‐black by day 10, consistent with normal hair cycle progression. In contrast, the dorsal skin of the model group remained pinkish‐white on day 7 and showed pigmentation until day 10, confirming the successful establishment of the AGA model. Both treatment groups displayed earlier skin darkening and visible hair regrowth relative to the model group. Notably, the CTIL‐treated mice showed accelerated pigmentation and hair emergence, with effects superior to those of the minoxidil group. This improved therapeutic efficacy may be attributed to the adhesive properties of CTIL, which likely prolonged its residence time on the skin surface and supported sustained delivery to hair follicles, in contrast to the rapid loss of aqueous minoxidil formulations.

FIGURE 7.

In vivo hair regrowth effects of CTIL. (A) Schematic illustration of the establishment and treatment process of the AGA mouse model. (B) Representative photographs of the dorsal skin of the mouse treated with different formulations. (C) H&E staining images of the treated skin on day 10. (D) Hair follicle length of the treated skin (n = 9). (E) Quantification of skin thickness (n = 3). (F) Representative SEM images of regrowth hair in each group on day 16. Scale bar = 30 µm.

Histological evaluation further corroborated these macroscopic observations. Hematoxylin and eosin (H&E) staining of dorsal skin harvested on day 10 (Figure 7C,D) revealed that hair follicles in both the model and minoxidil‐treated groups predominantly exhibited a relatively quiescent morphology, characterized by smaller follicular size and oval‐shaped hair bulbs, consistent with the telogen or early anagen transition stage. In contrast, follicles in the CTIL‐treated group displayed markedly enlarged and elongated morphologies with clearly defined hair shafts extending from the follicular structure, indicating a more advanced anagen phase. Increased perifollicular pigmentation was also observed, suggesting enhanced melanogenic activity during hair shaft formation. Additionally, dermal thickness in the CTIL group was significantly increased compared with the model group (Figure 7E), which was consistent with anagen‐phase‐associated follicular activation and dermal remodeling [49]. The structural quality of regenerated hair fibers was further examined by scanning electron microscopy (SEM). As shown in Figure 7F, regrowth hair in the CTIL group exhibited increased fiber diameter and well‐organized cuticle scales, whereas hairs from the model group appeared thinner with less defined surface morphology. Notably, the CTIL‐treated group showed improved hair shaft integrity compared with the minoxidil group, suggesting that CTIL not only promotes hair regeneration but may also facilitate maturation of hair fiber microstructure.

2.7. Taurine‐based Active Ionic Liquids Modulated the Perifollicular Microenvironment In Vivo

Building upon the observed promotion of hair regrowth by CTIL, we next investigated whether its therapeutic effects are associated with modulation of the perifollicular microenvironment. Cellular proliferation within hair follicles was first evaluated by immunofluorescent staining of Ki67, a widely used marker of proliferating cells. As shown in Figure 8A,E, the Ki67 fluorescence signals in the CTIL‐treated group were 1.40‐fold higher than those in the model group and closely comparable to the control group, indicating the enhanced proliferative activity of follicular cells during the anagen phase. To further evaluate follicular stem cell activation, SOX9 expression was analyzed as a representative marker of hair follicle stem cells. Consistent with Ki67 staining, SOX9 immunofluorescence intensity was elevated in the CTIL‐treated group relative to the model group (Figure 8B,F), indicating a more active stem cell niche and suggesting that CTIL may facilitate the activation of follicular stem cell populations involved in hair cycle re‐entry.

FIGURE 8.

CTIL modulated perifollicular microenvironment in vivo. Representative immunofluorescence images of (A) Ki67, (B) SOX9, (C) DHE, and (D) IL‐1β staining in treated skin on day 10. Scale bars: 100 µm for A, B, D; 200 µm for C. Semiquantitative analysis of fluorescence intensity of (E) Ki67, (F) SOX9, (G) DHE, and (H) IL‐1β (n = 3).

Given the central role of oxidative stress and inflammation in disrupting follicular homeostasis, we next evaluated these pathological factors in vivo. Dihydroethidium (DHE) staining revealed substantial ROS accumulation in the skin of AGA model mice, whereas CTIL treatment significantly reduced ROS signals within perifollicular regions (Figure 8C,G). This observation aligned with the potent antioxidant activity of CTIL demonstrated in vitro and confirmed its effectiveness in mitigating oxidative stress in vivo. Inflammatory status was further assessed by immunostaining of interleukin‐1β (IL‐1β), a pro‐inflammatory cytokine associated with hair follicle miniaturization and inflammatory alopecia. As shown in Figure 8D,H, strong IL‐1β expression was detected in the model group, whereas CTIL treatment resulted in a noticeable reduction in IL‐1β expression, suggesting suppression of local inflammatory signaling. In addition, no significant changes in body weight were observed among all groups during the treatment period (Figure S13), indicating that CTIL did not induce detectable systemic toxicity under the experimental conditions. Taken together, these findings demonstrated that CTIL may improve the perifollicular microenvironment by reducing oxidative stress, attenuating inflammatory responses, and enhancing proliferative activity within hair follicles, thereby supporting sustained follicular activation.

To further elucidate the molecular mechanism underlying CTIL‐regulated hair follicle regeneration, we examined the expression level of β‐catenin, a key regulatory molecule in hair follicle regeneration. Immunofluorescence staining results (Figure S14) showed that β‐catenin expression in the AGA model group was relatively weak and primarily localized around hair follicles, suggesting suppression of the Wnt/β‐catenin signaling pathway and impaired hair follicle regenerative activity. In contrast, CTIL treatment enhanced β‐catenin expression in skin tissues, indicating activation of the Wnt/β‐catenin signaling pathway. Combined with the expression profiles of Ki67, SOX9, DHE, and IL‐1β, these results suggested that CTIL not only created a favorable microenvironment for hair follicle regeneration by alleviating oxidative stress and inhibiting inflammatory responses but also promoted hair follicle regeneration and hair cycle progression. Moreover, activation of β‐catenin‐related signaling may have contributed to enhanced proliferation of hair follicle‐associated cells and facilitated the transition of hair follicles from the telogen phase to the anagen phase. Collectively, these findings provide further mechanistic evidence supporting the therapeutic potential of CTIL for AGA treatment.

3. Conclusions

In conclusion, an active pharmaceutical ingredient ionic liquid, CTIL, was successfully constructed via a one‐step strategy. CTIL exhibited well‐defined physicochemical properties, favorable biocompatibility, and rheological characteristics suitable for topical administration. Notably, the extensive hydrogen‐bonding interactions in CTIL were found to improve the poor transdermal permeability and limited skin retention of free taurine, thereby enhancing its penetration through the stratum corneum and facilitating accumulation within hair follicle structures. In vitro studies confirmed that CTIL not only acted as a penetration‐enhancing carrier but also directly scavenged ROS and protected hair follicle cells from oxidative damage. In a DHT‐induced AGA mouse model, topical administration of CTIL alleviated oxidative stress and inflammatory responses in the perifollicular microenvironment, promoted hair follicle proliferation and anagen transition, and ultimately facilitated hair regrowth. Collectively, this study provided a functional transdermal delivery system with both efficient delivery and intrinsic therapeutic activity for the topical treatment of AGA and further expanded the application of bioactive ionic liquids in the topical treatment of skin diseases.

4. Experimental Section

4.1. Materials

Tau was purchased from Adamas‐beta (Shanghai, China). Choline hydroxide (44 wt.% in H₂O) and RhB were purchased from Aladdin Biochemical Technology Co., Ltd (Shanghai, China). DiD and were purchased from MeilunBio (Dalian, China). Reactive Oxygen Species Assay Kit and Calcein/PI Cell Viability/Cytotoxicity Assay Kit were purchased from Beyotime Biotechnology Co., Ltd (Shanghai, China). DHT was obtained from Psaitong (Beijing, China). Minoxidil was purchased from Sunshine Mandi Pharmaceutical Co., Ltd (Zhejiang, China). The detailed information was as follows: Minoxidil Tincture (5%, 60 mL per bottle), National Medical Products Administration (NMPA) approval number H20010714.

4.2. Cell Lines and Animals

L929 mouse fibroblast cells (L929) and the human umbilical vein endothelial cell line (HUVEC) were obtained from the Chinese Academy of Sciences Cells Bank (Shanghai, China). DPCs were purchased from Fuheng Biotechnology Co., Ltd (Shanghai, China). The cells were cultured in complete DMEM cell culture medium, comprising 10% fetal bovine serum, 100 U/mL of penicillin G, and 100 U/mL of streptomycin sulfate.

Male C57BL/6 mice (6–7 weeks, 18–20 g) were purchased from Dossy Experimental Animals Co., Ltd. (Chengdu, China). All animal experiments were approved by the Animal Experimentation Ethics Committee of Sichuan University (Approval No. SCU42‐2512‐07).

4.3. Preparation and Characterization of CTIL

Completely dissolved 15 mmol of Tau in 40 mL of deionized water. Then, under an ice‐water bath, the aqueous solution containing equimolar choline hydroxide was dropped into the stirring Tau solution, continuing stirring for 4 h. Subsequently, most of the solvent was removed using a rotary evaporator under vacuum in a 60°C water bath to obtain the crude product. Freeze‐dried the crude product for 24 h to yield the final product CTIL, which was then stored in a vacuum drying oven for later use.

The structure of CTIL was characterized using nuclear magnetic resonance (NMR, 400 MHz, Bruker, Germany) spectroscopy and Fourier‐transform infrared (FT‐IR, INVENIO R, Bruker, Germany) spectroscopy. Tau in CTIL was quantitatively analyzed by high‐performance liquid chromatography (HPLC, Agilent, USA) with pre‐column derivatization. The chromatographic conditions were as follows: a C18 column (250 × 4.6 mm, 5 µm) was used with a column temperature of 25°C. The mobile phase consisted of a mixture of 15 mm sodium acetate buffer (pH = 4.2) and acetonitrile at a volume ratio of 70:30, and the flow rate was maintained at 1 mL/min. The derived samples were detected at a wavelength of 254 nm. The moisture content, viscosity, and conductivity of CTIL were determined using a Karl Fischer moisture titrator (870 KF Titrino plus, Metrohm, Switzerland), a rheometer (MCR302, Anton Paar, Austria), and a conductivity meter (DDS‐11A, Shanghai Yueping, China), respectively. Thermal analysis was performed using differential scanning calorimetry (DSC1, Mettler Toledo, Switzerland). An appropriate amount of each sample was placed in an aluminum crucible and scanned at a heating rate of 10 K/min under a nitrogen atmosphere. For liquid samples (ChOH and CTIL), the temperature was scanned from −80°C to 25°C, whereas for the solid sample (Tau), the scanning range was 25°C to 360°C.

4.4. Simulations for the Dimer Assembly

The GROMACS 4.67 package and the GROMOS96 force fields were used for the MD simulations [50, 51, 52]. The models for tau and choline were parameterized by the Automated Topology Builder [53, 54]. To study the assembly behavior of tau and choline, one tau and one choline were put into a simulation box with dimensions of 4 × 4 × 4 nm³. The 2168 SPC model water molecules were filled into the simulation box. The NPT ensemble was employed with the temperature kept at 296.15 K and the pressure at 1 bar. A 12 Å cutoff was applied to short‐range non‐bonded interactions, and long‐range electrostatic coupling and the Berendsen bath coupling scheme were employed. The simulations were performed in steps of 2 fs over a period of 2 ns. Following completion of the simulation, the assembled tau and choline dimer structure was obtained to check the free energy change by DFT calculation with a 6–311g++ basis and B3LYP functions using the Gaussian 16 code. Subsequent analysis involved electrostatic potential calculations.

4.5. DFT Calculation of Ionic Liquid Assembly

100 Tau and 100 choline were put into the simulation box randomly with 15765 water molecules filled by dimensions of 8 × 8 × 8 nm³. The 15765 water molecules were filled into the simulation box. The simulations were performed in steps of 2 fs over a period of 20 ns. Other simulation setup details were similar as mentioned above. After the simulation run, the snapshots of the simulation results were taken by VMD.

4.6. Cytocompatibility Assay

The MTT assay was used to examine the cytotoxicity of CTILs against L929 and HUVEC cells. When L929 or HUVEC cells were seeded into 96‐well plates and grew to 60% confluency, the fresh medium (as a control) or medium containing different formulations (Tau, ChOH, Tau + ChOH, and CTIL) at various Tau concentrations (1–2000 µg/mL) was introduced into the wells. At 24 h incubation, the supernatant of each well was aspirated and replaced with 100 µL MTT solution and incubated for another 4 h; then the supernatant was removed and replaced with DMSO. Finally, the viability of the cells was assessed at 490 nm using a microplate reader.

Live/dead cell staining was employed to visualize the cytotoxicity of different formulations. L929 or HUVEC were placed in 12‐well plates and allowed to reach approximately 60% confluency. The cells were then treated with different formulations for 24 h. Staining was performed in accordance with the instructions provided in the live/dead cell staining kit, and the staining results were subsequently observed using a fluorescence microscope.

For DPCs, cytocompatibility was evaluated using the MTT assay and the live/dead staining assay, respectively, following the same experimental procedures as described above for the other two cell lines.

4.7. Hemocompatibility Assay

Fresh mouse blood was collected and centrifuged at 2000 rpm and 4°C for 10 min to remove the supernatant. The pellet was then washed with PBS buffer until the supernatant became colorless and transparent to obtain red blood cells. Gently mix 20 µL of the red blood cell suspension with 980 µL of CTIL. Meanwhile, PBS buffer was used as the negative control, and deionized water served as the positive control. All the aforementioned solutions were incubated in a constant‐temperature shaker at 37°C for 2 h, followed by centrifugation at 2000 rpm and 4°C for 10 min. Subsequently, all samples were photographed, and the absorbance of the supernatant at 540 nm was measured using a microplate reader to calculate the hemolysis rate.

4.8. Acute Skin Irritation Test

Healthy C57BL/6 mice were fed under standard conditions. After dorsal depilation, the animals were randomly divided into three groups (n = 3). 24 h after depilation, 0.5 mL of the test formulation was evenly applied to a piece of gauze, which was then placed on the depilated dorsal skin and secured with adhesive tape. After 4 h of exposure, the gauze was removed, and the application site was photographed at 1, 24, 48, 72, and 96 h post‐removal. Skin irritation was evaluated by scoring erythema and edema according to Tables S1 and S2.

4.9. In Vitro Antioxidant Capability Evaluation

Equal volumes of ABTS solution (7 mm) and potassium persulfate solution (2.45 mm) were mixed, followed by reaction at room temperature in the dark for 12 h to prepare the working stock solution. Subsequently, 1.8 mL of the diluted working stock solution was uniformly mixed with 200 µL of the sample solutions at different concentrations, and the mixture was reacted in the dark for 10 min. Finally, the ultraviolet (UV) spectra of each group of solutions were scanned using a UV–vis spectrophotometer, and the absorbance at the maximum absorption wavelength was recorded to calculate the scavenging rate.

4.10. Superoxide Anion Radical Scavenging Test

Equal amounts of Tau, ChOH, and CTIL were separately dissolved in Tris‐HCl buffer (pH 8.2). Subsequently, freshly prepared pyrogallol solution was added to each system to a final concentration of 250 µm. After incubation at room temperature for 5 min, the absorbance at 320 nm was measured using a microplate reader (Victor Nivo HH3500, Perkin Elmer, Germany), and the ·O₂ ⁻ inhibition rate was calculated. Ascorbic acid (Vc) was used as a positive control.

4.11. Intracellular Antioxidant Effects Evaluation

When L929 cells or DPCs were seeded into 12‐well plates and grew to 60% confluency, the medium was replaced with serum‐free medium containing CTIL (500 µg/mL) and H₂O₂ (500 mm). Serum‐free medium served as the negative control, while serum‐free medium supplemented with H₂O₂ acted as the positive control. After co‐incubation for 4 h, the supernatant was removed, and the cells were washed with PBS buffer. Subsequently, the cells were incubated with the DCFH‐DA probe for 30 min. Finally, ROS levels in each group were observed using a fluorescence microscope (DMi8A, Leica, China).

4.12. In Vitro Quantification of Transdermal Permeation and Skin Retention

The Franz diffusion cell method was employed to evaluate the in vitro transdermal delivery of CTIL. Throughout the experiment, the magnetic stirrer speed was set to 300 rpm, and the temperature was maintained at 32 ± 1°C (human skin surface temperature). C57BL/6 mouse skin was sandwiched between the donor cell and the receptor cell, with an effective permeation area of 1.77 cm³. The receptor cell was filled with 14 mL of PBS buffer. Then, 400 µL of CTIL or Tau suspension (Tau sus, containing an equivalent amount of Tau) was added to the donor cells, respectively. At predetermined time points (1, 2, 4, 8, 12, and 24 h), 2 mL receptor cell solutions were taken, and an equal volume of fresh PBS buffer was added to the receptor cell immediately. At the final time point, the skin was collected and scrubbed three times with water, and the Tau retained in the skin was extracted using methanol. The permeability and skin retention of Tau were analyzed using the precolumn derivatization HPLC method as described earlier.

4.13. In Vitro Visualization of Skin Transport

DiD and RhB were used as representative hydrophobic and hydrophilic molecules, respectively, to investigate the enhancement effect of CTIL on the transdermal delivery of drugs with different properties. Hair on the back of C57BL/6 mice was removed with a rosin paraffin mixture. One day after recovery, DiD‐loaded CTIL (DiD@CTIL) was applied topically to the hair removal area, with free DiD for control (at a dosage of 0.8 mg/kg). Similarly, RhB@CGIL was applied topically with free RhB for control (at a dosage of 0.8 mg/kg). After 24 h, all the formulations were wiped off, and the skin in the administration area was collected. Finally, all the samples were observed under a confocal laser scanning microscope (CLSM, Leica, Wetzlar, Germany).

4.14. Hair Regrowth Assessment in the AGA Model

The AGA mouse model was established via intraperitoneal injection, with a single dose of 1 mg DHT administered per mouse [55]. The DHT administration protocol consisted of 4 consecutive days of injection followed by a 1‐day break, with this cycle repeated throughout the experimental period. At the end of the first cycle, the dorsal hair of the mice was removed using a paraffin‐rosin mixture, and the mice were randomly divided into 4 groups (n = 4) as follows: Control group: healthy mice without any treatment; Model group: AGA mice any treatment; Minoxidil group: AGA mice topically treated with 5% commercially available minoxidil (100 µL per mouse) every day; CTIL group: AGA mice topically treated with CTIL (50 mg/kg) every day. The depilated region of each mouse was photographed to track the changes in skin and hair throughout the experiment. On day 10, dorsal skin samples of each group were collected for H&E staining and immunofluorescence staining to evaluate the hair growth and hair follicle microenvironment. Regenerated hair from mice in each treatment group was harvested and then observed morphology using scanning electron microscopy (SEM, S‐4800, Hitachi, Japan).

4.15. Statistical Analysis

Data were presented as mean ± standard deviation (SD). Group differences were assessed by Student's t‐test, one‐way or two‐way ANOVA for multiple comparisons. Statistical significance was defined as ^* p < 0.05, ^** p < 0.01, ^*** p < 0.001, and ^**** p < 0.0001.

Author Contributions

Jun Cao, Yuanwei Chen: conceptualization. Lianyi Yang: methodology. Lianyi Yang, Yazhen Wang: investigation. Lianyi Yang, Yazhen Wang, Lei Lei: visualization. Jun Cao, Yuanwei Chen: supervision. Lianyi Yang: writing – original draft. Bin He, Jun Cao, Yuanwei Chen: writing – review & editing

Funding

The authors have nothing to report.

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Supporting File: adhm71363‐sup‐0001‐SuppMat.docx.

Data Availability Statement

The data that support the findings of this study are available on request from the corresponding author. The data are not publicly available due to privacy or ethical restrictions.