A two-stage transdermal drug delivery system comprising sono-phase-change transfersomes for non-invasive deep dermal delivery

Abstract

Non-invasive deep transdermal drug delivery is required for conditions such as keloids. However, the skin’s stratum corneum barrier is the primary obstacle for transdermal drug delivery methods. Most energy sources and operational controls are located externally, making it difficult to regulate the deep delivery of a drug once it has penetrated the stratum corneum barrier. Additionally, some transdermal administration methods that provide an internal energy source are frequently invasive and can cause damage. Therefore, achieving non-invasive drug delivery to the deep dermis for such conditions is challenging. Here, we propose sono-phase-change transfersomes (SPCTs) that stably encapsulate perfluoro-n-pentane within transfersomes. These SPCTs, combined with low-frequency and low-intensity ultrasound (LFLIU), are designed to create a two-stage transdermal drug delivery system that integrates passive and active penetration. In the first stage, the deformability of SPCTs enables them to stably and passively penetrate the epidermis. In the second stage, SPCTs undergo a phase change when ultrasound irradiation is applied, transforming into microbubbles. As the microbubbles penetrate deeper, they gradually expand, rupture, and release their encapsulated substances, providing supplementary energy from within to achieve active penetration. Both in vitro and in vivo experiments revealed a significant increase in the efficiency, depth, quantity, and distribution range of the contents entering the dermis. In addition, the SPCTs combined with the ultrasound group in the keloid nude mouse model exhibited the fastest keloid volume reduction, with various indicators demonstrating that its therapeutic effect was significantly better than that of the other control groups. These findings indicate that this two-stage transdermal drug delivery system can non-invasively and safely achieve deeper and higher-dose dermal administration, offering a new strategy for treating diseases such as keloid.

Graphical Abstract

The working principle of the two-stage transdermal drug delivery system.

Supplementary Information

The online version contains supplementary material available at 10.1186/s12951-025-03710-6.

Introduction

Keloid is a skin disease that is difficult to heal spontaneously, characterized by excessive fibroblast proliferation in the dermis, massive extracellular matrix deposition, and abnormal epidermal structure [1, 2]. Its rigid nature hinders effective drug penetration and deep dermal distribution. Despite various treatment methods, including drug injection [3] and surgical resection [4, 5], keloid has high recurrence rates [6] and complications including pain, subcutaneous atrophy, and capillary dilation caused by injection [7]. Currently, there is a lack of effective treatment methods for keloids, which presents a challenge for clinical treatments. While transdermal drug delivery can reduce trauma, pain and minimize side effects [8–10], how to effectively utilize its advantages while enhancing the efficiency of drug penetration has become a focus of current transdermal drug delivery methods for keloid treatment.

Transfersome is a specific type of liposome that consists of an aqueous core and lipid bilayer, which is composed of phospholipids and a surfactant [11–13]. Surfactants are distributed in the lipid bilayer to increase the deformability of the vesicle membrane, allowing it to deform through channels many times smaller than its own particle size [14–16]. When applied to the skin surface in an unconfined state, transfersomes can penetrate into the water-rich epidermis as water evaporates, spontaneously deforming across the stratum corneum to enter the deeper epidermal layers while maintaining the vesicular structure [17, 18]. Compared with conventional liposomes with rigid structure that can only deposit in the stratum corneum, transfersomes increase the transdermal flux by several-fold [19–23]. Compared with ethosomes, transfersomes better maintain structural integrity during penetration [24–26]. In contrast to phase-change nanodroplets, transfersomes accommodate a broader spectrum of drugs and achieve higher transdermal efficiency [27–29]. Consequently, due to their advantages in transdermal efficiency, transfersomes have been widely used for transdermal drug delivery of a wide range of drugs [30–34], and have now also been used for transdermal drug delivery in keloid [35, 36].

However, while transfersomes can effectively penetrate the stratum corneum, they lack the driving force of the hydration gradient after reaching the dermis, restricting their subsequent delivery [37, 38]. Therefore, it is necessary to seek better treatments for diseases such as keloid to achieve higher efficiency, increased doses, and deeper drug penetration into the dermis [39]. Evidently, relying solely on transfersomes to achieve the target is difficult, and several physical transdermal methods (iontophoresis, microneedles, and ultrasound) have been used in keloid treatment [40–43]. It has also been proposed that transfersomes combined with physical approaches facilitate drug penetration [44–47]. However, the dermis is richly vascularized and tends to be significantly thickened in keloid tissues [48]. Among existing approaches, it is difficult to reach a depth of 500 μm using iontophoresis and laser [49–52]. Additionally, microneedle penetration greater than 960 μm causes significant pain sensations, while bleeding and the risk of infection occur at depths beyond 1450 μm [53–56]. Moreover, keloid is prone to recurrence due to trauma, so it is not applicable. In these studies, most energy sources and operational controls are located externally, making it difficult to regulate the deep delivery of a drug once it has penetrated the stratum corneum barrier. Providing an internal energy source is frequently invasive. As a result, further research is needed to achieve efficient and safe drug delivery into the deep dermis through transdermal administration.

Herein, based on transfersomes, we propose sono-phase-change transfersomes (SPCTs) that encapsulate delivery substances (e.g., drugs, fluorescent markers) and an ultrasound-responsive substance (perfluoro-n-pentane [PFP]) (Fig. 1A). As the material foundation enabling synergistic transdermal drug delivery, SPCTs retain the deformability and hydrophilicity characteristic of transfersomes while incorporating ultrasound responsiveness. When combined with ultrasound, SPCTs collaboratively constitute a two-stage transdermal drug delivery system. Fig. 1B indicates that this system’s delivery process involves the following two stages: (1) Driven by the hydration gradient, SPCT deforms to pass through the stratum corneum barrier and diffuse deeper. (2) Ultrasound irradiation facilitates further penetration of SPCT and triggers an internal PFP phase transition, leading to its gradual expansion and enlargement. Under continuous ultrasound irradiation, SPCT ruptures, achieving deeper drug penetration primarily through the microjet effect generated by cavitation.

Fig. 1.

The working principle of the two-stage transdermal drug delivery system. (A) Preparation and structural schematic diagram of SPCT. (B) When the SPCT solution is loaded on the skin surface, SPCTs deform through intercellular channels in the stratum corneum and steadily penetrate the stratum corneum. When ultrasound is applied, SPCTs further penetrate and trigger the phase transition of internal PFP to form microbubbles, which ultimately rupture and release the contents into the deep dermis

During this process, SPCT deforms to pass through the stratum corneum barrier while maintaining its structural integrity, driven by the permeable hydration gradient (Fig. 1B). Within the skin structure, the water content increases from the stratum corneum to the dermis, allowing SPCT to continuously diffuse downward [57, 58]. Once SPCT reaches the dermis, the difference in water content suddenly decreases, failing to provide sufficient driving force for further diffusion. After that, ultrasonic irradiation can drive SPCT to penetrate deeper into the dermis, and trigger the phase transition of PFP, resulting in the gradual expansion of SPCT volume [59–64]. As the SPCT microbubbles expand, their cavitation rupture generates microjets accompanied by the mechanical friction effect, facilitating drug delivery to the deeper layers of the dermis [65–67]. This ensures penetration efficiency and dosage, allowing a large amount of content to penetrate deeper into the dermis. The response intensity can be adjusted by controlling the amount of PFP and ultrasonic parameters. We chose keloid as the target and prepared SPCTs containing common keloid treatment drug (5-fluorouracil [5-FU]) and fluorescent markers to evaluate the effect of SPCTs combined with ultrasound for transdermal drug delivery. The results revealed that SPCTs could spontaneously pass through the keloid epidermis and enter the dermis more deeply with appropriate ultrasound irradiation. As demonstrated by in vitro and in vivo evaluations, this approach illustrates the relative safety and efficacy of transdermal delivery, higher efficiency, higher drug doses, and deeper dermal penetration when SPCTs are combined with ultrasound. Combining SPCT with ultrasound could be a new transdermal drug delivery system with improved therapeutic outcomes.

Materials and methods

Materials

Soybean phosphatidylcholine (SPC) was procured from Avanti Polar Lipids (Alabaster, AL, USA). Nonionic surfactant Span^® 80 was obtained from Aladdin (Shanghai, China). Phosphate-buffered saline (PBS, pH = 7.4), 5-FU, and PFP were acquired from Macklin Biochemical Technology (Shanghai, China). Water-soluble CdTe quantum dots (CdTe QDs) solution was obtained from Xingzi New Material Technology Development (Shanghai, China). 3, 3’-Dioctadecyloxacarbocyanine perchlorate (DiO, ex/em: 484/501 nm) was obtained from Sigma-Aldrich (St. Louis, USA), chloroform and methanol were procured from Chongqing Chuandong Chemical (Chongqing, China), and polycarbonate membranes were acquired from Aladdin (Shanghai, China). Polydimethylsiloxane (PDMS) was procured from Dow Corning (Midland, USA), and the Opti-mum cutting temperature (OCT) compound was obtained from Sakura Finetek USA, Inc. (Torrance, CA, USA). Agarose was purchased from Thermo Fisher Scientific Inc. (Waltham, USA). The frozen section antigen repair solution was acquired from Solarbio (Beijing, China). Ki67 was procured from Cell Signaling Technology (Boston, USA). Cy3 labeled goat anti-rabbit lgG (H + L), BCA protein concentration determination kit (Enhanced), and QuickBlockTM Western blocking solution were obtained from Beyotime (Shanghai, China). Collagen type I polyclonal antibody and collagen type III (N-terminal) polyclonal antibody were acquired from Proteintech (Chicago, USA). Goat anti-mouse IgG was obtained from Elk Biotechnology (Wuhan, China).

Preparation of conventional liposomes, transfersomes, and SPCTs

The drug carriers were prepared using the conventional rotary evaporation ultrasonic method [15, 68]. First, SPC, Span^®80 (mass ratio 4:1) with 5 µM DiO was dissolved in chloroform and methanol (2:1, v/v) [69]. The organic solvent was subsequently completely removed, and a film was formed by a rotary evaporator operating under vacuum. The solution containing CdTe QDs and 5-FU was then added to hydrate the film, and the mixed solution was sonicated at 60 W for 2 min in an ice bath using an ultrasonic probe (VCX150, Sonics & Material Inc., Newtown, CT, USA). Similarly, liposomes were prepared as described above, except that Span 80 was excluded. After the synthesis of transfersomes (Ts), a 200 µL of PFP solution was added [70, 71], and the mixed solution was sonicated for 2 min in an ice bath using the same sonicator to create CdTe-DiO-FU SPCTs (CDF@SPCTs). In the preparation of SPCTs containing only the drug 5-FU (5-FU@SPCTs) (without fluorescent dyes), the steps were the same except no fluorescent dye was added. Finally, the solution was filtered through a 220 nm polycarbonate membrane.

Characterization of SPCTs

Measurement of vesicle size and zeta potential

The particle size distribution and zeta potential of the prepared SPCTs were measured using dynamic light scattering (DLS, ZSU3200, Malvern Panalytical, Malvern, UK). Each sample was measured in triplicate.

Entrapment efficiency (EE) and drug loading (DL)

An ultraviolet spectrophotometer (UV-3600iPLUS, Shimadzu, Kyoto, Japan) was used to measure the absorbance at different concentrations (20, 40, 60, 80, and 100 µg/mL) at 266 nm. Subsequently, the standard curve of 5-FU was drawn based on the above data. The SPCT solution was centrifuged at 10,000 r/min for 10 min at 4℃ to determine the EE and DL of SPCTs. The absorbance of the supernatant was measured, and the encapsulation efficiency of 5-FU was calculated using the formula (1) EE = (m₁–m₂)/m. The 5-FU DL rate was calculated using the formula (2) DL = (m₁–m₂)/m₃, where m₁ represents the total amount of 5-FU input, m₂ represents the free supernatant of 5-FU, and m₃ represents the total amount of SPCTs used.

Degree of deformability

The 5 mL of liposome or SPCT suspension was diluted ten times with PBS. The mixture was squeezed through a 220 nm polycarbonate membrane at the same pressure and at the extrusion temperature of 24℃, in which the membrane size was determined based on the particle size of the carrier obtained from the experiment. The particle sizes of the liposomes and SPCTs were then measured using DLS. The deformation index was calculated as follows:

where j represents the volume of the suspension extruded, r_v represents the particle size of the carriers after extrusion, and r_p represents the pore size of the polycarbonate membrane.

Transmission electron microscopy (TEM)

TEM (h-7500, Hitachi, Tokyo, Japan) was used to observe the morphology of SPCTs. The SPCT solution was dripped onto a copper grating coated with a carbon film, and excess liquid was removed with filter paper. The samples were observed after drying naturally for 24 h.

Stability test

The SPCT solution was stored at 4℃ in a refrigerator. In the following week, the average size and potential were measured using the same method as that used to measure the vesicle size and potential.

Measurement of cavitation signals in SPCTs combined with ultrasound

The hydrophone (RHS(A)−20, Hangzhou Applied Acoustic Research Institute, Hangzhou, China) and LFLIU device (SY23R7151K002, Chongqing Ronghai Ultrasonic Medical Engineering Research Center Co., Ltd., Chongqing, China) were placed 45° into a container filled with degassed water. The frequency of the LFLIU device was 360 kHz and its power was set to 1 W. The ultrasound instrument probe was submerged in a cylindrical chamber containing the SPCT solution. A hydrophone was used to detect the ultrasound signals, and an oscilloscope (MSO64, Tektronix, OR, USA) connected to it displayed the waveforms and recorded the data in real time. The collected signals were subsequently subjected to fast Fourier transform (FFT) using Matlab software (Mathworks, Natick, MA, USA). The resulting spectrograms were analyzed to observe cavitation.

The controlled-release experiment of SPCTs

First, we added 1 ml of SPCT solution to the hole of an agarose skin model (3% w/v). The model was subsequently placed in a container that enabled temperature control and ultrasound diagnostic equipment (My Lab ClassC Advanced, Esaote, EsaoteS.p.A. Genova, Italy) was used to irradiate the agarose model from the side to observe the ultrasound imaging of SPCTs in the hole. The ultrasound transdermal drug delivery device (360 kHz, 1 W) was used to apply ultrasound from the top of the hole. The brightness change of SPCTs in the hole was monitored to quantify the ultrasound response of SPCTs.

In vitro cytotoxicity assay

Human immortalized keratinocytes (HaCaT) were used as model cells to assess the cytotoxicity of Ts and SPCTs without 5-FU. In addition, the cytotoxicity of the two carriers combined with LFLIU (360 kHz, 1 W) irradiation for 40 min was also measured. The cells were cultured in α-MEM medium supplemented with 10% fetal bovine serum and antibiotics at 37℃ and 5% CO₂. The cells were suspended in a cell culture medium and inoculated in 96-well plates at a density of 1 × 10⁴ cells per well. The cells were exposed to fresh medium containing the corresponding samples and treated accordingly. 100 µL of fresh medium and 10 µL of cell counting kit-8 (CCK-8) were added to each well and incubated for 2 h. An enzyme marker was used to measure the absorbance at 450 nm. The cell viability was calculated as follows:

Cell viability (%) = (absorbance of the treated cells/absorbance of the control group) × 100.

Penetration effect of SPCTs combined with ultrasound in vitro

The isolated keloid tissues of patients were selected for experiments (approved by the Ethics Committee of Chongqing Hospital of Traditional Chinese Medicine (No. 2024-KY-HY-40)). Each patient or their surrogates provided informed consent. Franz diffusion cells were utilized in the in vitro experiment [72]. The keloid was fixed between the donor chamber and the receptor chamber, with the epidermis facing upwards. Since the receiving solution was used to mimic the subcutaneous circulation of human blood, the receptor chamber was filled with PBS (pH = 7.4), maintained at a temperature of 37 ± 0.5℃ with a stirring rate of 240 r/min. In this experiment, the donor chamber was replaced with a Liquid storage cell composed of PDMS, which was completely affixed to the skin to hold 1 mL of solution. The PDMS and the remaining solution on the tissue surface were removed after passive administration. A coupling agent was applied to the ultrasound probe, and ultrasound was used just above the tissue epidermis.

The keloid samples were embedded in OCT, frozen at − 20℃, and sectioned into longitudinal or horizontal slices to observe the fluorescence penetration depth of each group. The thickness of both longitudinal and horizontal slices was 10 μm. Confocal laser scanning microscopy was used to observe the tissue sections. After 5-FU drug administration, the keloid was ground into a tissue homogenate and dissolved in hydrochloric acid to determine the amount of drug in the tissue. The supernatant was obtained by centrifuging the resulting mixture. High-performance liquid chromatography (HPLC) (Agilent C18, USA) was used to analyze the supernatant. The mobile phase was methanol-water (10:90 v/v at a flow rate of 1.0 mL/min), and the detection was conducted at a wavelength of 266 nm.

Drug release curves of SPCTs

Using a Franz diffusion cell, fill the receptor chamber with 25 mL of PBS, add 1 mL of 5-FU@ SPCTs to the donor chamber, and place the skin tissue with the stratum corneum facing upwards in it. Before applying LFLIU irradiation, 5-FU@SPCTs were allowed to passively permeate for 40 min. Subsequently, ultrasound irradiation was performed, and samples from the donor chamber were collected every 10 min, with PBS supplemented to maintain a constant volume. The absorption peak at 266 nm was measured using a UV spectrophotometer, the release amount of 5-FU was calculated, and the corresponding drug release curve was plotted.

Penetration effect of SPCTs combined with ultrasound in vivo

We selected 6–8-week-old male BALB/c nude mice (Enswell Company, Chongqing, China) to create animal models. All experimental procedures involving animals followed the protocol reviewed and approved by the Laboratory Animal Welfare and Ethics Committee of Chongqing Traditional Chinese Medicine Hospital (No. 2024-DWSY-XW). Keloid samples were obtained after the patients provided written informed consent, and the procedure was approved by the Ethics Committee of the Chongqing Traditional Chinese Medicine Hospital (No. 2024-KY-HY-40). The keloid tissues were immediately cut into cylindrical shapes of 8 × 8 × 3 mm using an 8 mm diameter circular punch once they arrived at the laboratory. Subsequently, the nude mice were anesthetized with gas, and a 6 mm diameter punch was used to remove 6 × 6 mm of skin from the backs of the nude mice. Four keloid tissue samples from the same donor were implanted on the backs of nude mice to observe the modeling effect. After transplantation, keloid tissue samples were taken from the nude mice weekly and H&E staining was used to observe the tissue structure.

Keloid tissues from the same donor were implanted in nude mice to observe the therapeutic effect of each group after administration. During the in vivo experiments, none of the drug carriers added CdTe quantum dots or DiO membrane fluorescence. The transdermal drug delivery experiments began after the wound had healed. First, the SPCT solution was added to the nude mouse-implanted keloid tissues based on the parameters of in vitro experiments to demonstrate the safety of the administration. Then, the infrared thermometer camera (Fotric, Shanghai, China) was used to measure the real-time temperature of the nude mice within 60 min of ultrasound exposure. Additionally, the two drug carriers were combined with ultrasound separately, and the tissues were subsequently sectioned and stained with H&E.

Keloid samples from the same nude mice were divided into four groups: Control (without treatment), 5-FU@SPCTs passive penetration only, transfersomes loaded with 5-FU (5-FU@Ts) combined with ultrasound, and 5-FU@SPCTs combined with ultrasound. Relevant methods were used for treatment. Except for the Control group, we added 1 mL (drug concentration = 1 mg/mL) of carrier solution to the other three groups. Treatment was performed twice weekly for four weeks. Before each administration, the short (a) and long (b) diameters of the keloids were measured using calipers and recorded. When measuring the volume, only the underlying surviving keloid tissue was measured. The formula for keloid volume was a²b/2.

Immunofluorescence

After four weeks, we removed keloid tissues from the animal models for immunofluorescence detection. The keloid samples were embedded in OCT (4583, Sakura, CA, USA), frozen at − 20℃, and microtomized into longitudinal sections. The sections were incubated with antigen repair solution (C1035, Solarbio, Beijing, China) at room temperature for 10 min and subsequently incubated with primary antibody Ki67 (9129, Cell Signaling Technology, BO, USA) overnight at 4 °C. After this process, we added the secondary antibody (Cy3-labeled goat anti-rabbit lgG (H + L), Beyotime, Shanghai, China) to the sample and incubated again at room temperature for 1 h. A confocal microscope (Leica SP8 LIGHT, Leica, SO, Germany) was used to observe the results.

Western blotting

Total keloid proteins were extracted using a protein extraction buffer (P0013B, Beyotime, Shanghai, China). BCA protein assay kit (P0010S, Beyotime, Shanghai, China) was used to calculate protein concentration. Proteins were separated on 10% SDS-PAGE gels and transferred onto a polyvinylidene fluoride (PVDF) membrane. The PVDF membrane was subsequently blocked with QuickBlockTM Western blocking solution (P0252–500 mL, Beyotime, Shanghai, China) for 1 h at room temperature. The membrane was subsequently incubated overnight at 4 °C with primary antibodies against type I collagen (14695-1-AP, Proteintech, Chicago, USA) and type III collagen (22734-1-AP, Proteintech, Chicago, USA). After washing thrice with TBS-T, the membrane was incubated with HRP-conjugated secondary antibodies (SA002, Elk Biotechnology, Wuhan, China) at room temperature for 1 h. An enhanced chemiluminescence approach was used to visualize the protein bands, and images were acquired through a gel imaging system.

Statistical analysis

Data are represented as mean ± standard deviation (SD) (n ≥ 3). ImageJ software (National Institutes of Health, USA) was used to quantify fluorescence intensity. GraphPad Prism software (version 9.5.0) was used for data analyses. The Student’s t-test was used to determine if there was a significant difference between the two groups. Differences between multiple groups were analyzed using one-way analysis of variance. The data follow a normal distribution and conform to variance homogeneity. The data were deemed to be statistically significant when p < 0.05 (*), p < 0.01 (**), and p < 0.001 (***) were observed, while “ns” indicated no significant difference.

Results and discussion

Characterization of SPCTs

TEM images revealed that the prepared 5-FU@SPCTs were spherical with a particle size of approximately 200 nm (Fig. 2A). The size distribution of 5-FU@SPCTs, measured using the laser particle size analyzer, was 240.36 ± 1.38 nm (Figure S1), consistent with the TEM results. The UV curves of Ts, 5-FU@SPCTs and free 5-FU solution are shown in Fig. 2B. The absorption peaks of 5-FU and CDF@SPCTs were observed at a wavelength of 266 nm, indicating that SPCTs had successfully encapsulated the drug. The 5-FU EE was 70%, and the 5-FU loading was 2.02 wt% calculated using the UV standard curve of 5-FU (Figure S2). The deformability of 5-FU@SPCTs was 19.41 (Table S1). Consequently, the particle size of the prepared 5-FU@SPCTs, EE, and deformability were appropriate for transdermal drug delivery [73, 74]. Fig. 2C depicts the changes in particle size and zeta potential after one week; no significant difference was observed, indicating that the prepared 5-FU@SPCTs were stable for one week. Before ultrasound irradiation (0 min), no microbubbles were generated. After 20 min of ultrasound irradiation, 5-FU@SPCTs gradually expanded and formed obvious microbubbles (Fig. 2D), indicating that PFP was successfully encapsulated in 5-FU@SPCTs.

Fig. 2.

Preparation and characterization of 5-FU@SPCTs. (A) TEM images of 5-FU@SPCTs. (B) The UV absorption of Ts, 5-FU@SPCTs, and free 5-FU solution. (C) Particle size and zeta potential stability of 5-FU@SPCTs over 7 days. (D) In optical images, the change of 5-FU@SPCTs after ultrasonic irradiation (360 kHz, 1 W)

Penetration effect of SPCTs in vitro (without ultrasound)

In comprehensive characterizations, the penetration effect of passive transdermal drug delivery penetration in keloids was first tested without the application of ultrasound. Fluorescent markers were used to demonstrate the permeation effect of each group. CdTe QDs (red fluorescence) were encapsulated in the aqueous phase of CDF@SPCTs, whereas DiO (green fluorescence) was loaded on CDF@SPCTs’ membrane. Fig. 3A depicts the selected passive penetration duration of 60 min, with longitudinal sampling and observation occurring every 10 min. We found that CdTe QDs loaded directly onto the surface of keloids underwent a static process lasting for 60 min. The red fluorescence was visible only outside the stratum corneum (Fig. 3B). For transdermal drug delivery, low molecular weight and fat-soluble substances exhibit feasible and easier transdermal penetration. Alternatively, achieving penetration can be difficult. As a consequence, the stratum corneum completely blocks water-soluble CdTe QDs [75]. Similarly, green fluorescence (DiO) and red fluorescence (CdTe QDs) were predominantly outside the stratum corneum when CdTe-DiO-FU liposomes (CDF@Liposomes) were loaded onto the keloid surface for 60 min (Fig. 3B). This phenomenon might be attributed to the rigid structure of liposomes and the dense structure of keloid tissue, which prevent liposomes from penetrating through the stratum corneum barrier. The experimental results of CDF@SPCTs standing for 40 min revealed significant green and red fluorescence near the superficial layer of the demis. The entry depth did not differ significantly when CDF@SPCTs were left to stand for 60 min (Fig. 3B). The above results are due to the hydrophilic and deformability of transfersome. The water content of the epidermis ranges from 10% to 30%, whereas that of the active epidermis increases to 75%, thus SPCTs are driven by the hydration gradient and deform through intercellular channels within the stratum corneum and subsequently stably pass through the epidermis [76, 77]. Accordingly, the penetration effect of SPCTs is significantly better than that of no carrier or conventional Liposomes. However, the driving force for SPCTs to continue penetrating deeper into the skin diminished as the water content stabilized after entering the dermis. Consequently, 40 min was an appropriate choice as the application time for passive drug delivery in subsequent experiments.

Fig. 3.

In vitro passive penetration and ultrasound controlled-release effects. (A) Schematic diagram of the passive permeation in vitro experiment. (B) Fluorescence distribution images of passive penetration: CdTe QDs, CDF@Liposomes, and CDF@SPCTs. Scale bar = 100 μm. (C) Schematic diagram of observing the controlled-release effect of SPCTs. (D) B-mode and contrast-mode images of the CDF@SPCT solution stored in an agarose skin model at 32℃ for 40 min and (E) B-mode and contrast-mode images of continuous ultrasonic irradiation for 60 min. (F) The mean gray value of the CDF@SPCT solution in B-mode and contrast-mode within 40 min of standing. (G) The mean gray value of the CDF@SPCT solution in B-mode and contrast-mode within 60 min of ultrasound. (H) Schematic diagram of the measurement of the cavitation signal. (I) Spectrum diagram of ultrasound combined with degassed water. (J) Spectrum diagram of ultrasound combined with CDF@SPCTs.

Controlled-release effects of SPCTs

The epidermis is the outermost layer of human skin. It is located at the skin surface and is nonvascularized; therefore, it is closer to the temperature of the tissue surface (32℃) [78, 79]. The passive delivery of SPCTs reaches only the superficial dermis. The phase change temperature of PFP is 29℃; however, the temperature increases when PFP is encapsulated into the carrier. Additionally, PFP is susceptible to phase transition when low-frequency ultrasound irradiates the carrier [76]. The CDF@SPCT solution was added to the agarose model as shown in Fig. 3C. Temperature control was performed at the bottom, an imaging ultrasound probe was placed on the side for observation, and a therapeutic ultrasound probe was placed on the top. When the CDF@SPCT solution was kept in the agarose model without ultrasound irradiation at 32℃ for 40 min, the mean gray value in B-mode and contrast-mode did not differ significantly, indicating that PFP did not undergo phase transition (Fig. 3D and F). This ensures that the internal PFP does not undergo a phase change while passing through the stratum corneum or even the entire epidermis, thereby preserving the structural integrity of SPCT during penetration through the epidermis. Subsequently, therapeutic ultrasound was used to irradiate the CDF@SPCT solution according to our previous work on transdermal carriers loaded with PFP [80]. We employed LFLIU (360 kHz, 1 W) as a therapeutic ultrasound to irradiate the CDF@SPCT solution in the agarose model under the same environment. The brightness of the CDF@SPCT solution gradually increased in B-mode and contrast-mode imaging when ultrasound irradiation was applied, indicating a phase transition of the PFP inside (Fig. 3E). The brightness gradually increased within 20 min (B-mode: from 19.97 ± 1.2 pixels to 30.06 ± 1 pixels; contrast-mode: from 4.81 ± 0.47 pixels to 24.62 ± 0.96 pixels) and was maintained until 40 min (Fig. 3G). Due to the constant amount of CDF@SPCT solution, the brightness began to weaken as PFP gradually vaporized. After 60 min, the brightness inside the hole became very weak (B-mode: 18.93 ± 1.68 pixels; contrast-mode: 14.83 ± 1.49 pixels). The analysis results of the gray values in both modes are consistent with the images.

To observe the cavitation of SPCT, the 5-FU@SPCT solution was diluted 100 times, and ultrasound was applied and observed under an optical microscope. Microbubbles formed within the first 40 min of ultrasound irradiation. After 40 min, all microbubbles produced by 5-FU@SPCTs had ruptured, and the observation field was devoid of obvious microbubbles (Figure S3), consistent with the findings from the previous gray value analysis. Fig. 3H illustrates that the signal was monitored during the process of 5-FU@SPCTs combined with ultrasound. When degassed water was used as the control group, the combination of degassed water and ultrasound did not produce significant cavitation signals (Fig. 3I). However, 5-FU@SPCTs combined with ultrasound produced significantly higher harmonic signals (Fig. 3J), indicating that inertial cavitation had occurred. The above results suggest that SPCTs can spontaneously pass through the stratum corneum barrier and reach the dermis stably. In the two-stage transdermal drug delivery system composed of SPCTs and LFLIU, SPCTs would expand and enlarge to form microbubbles under continuous ultrasound irradiation, followed by gradual rupture. The system achieves active drug delivery through the cavitation effect generated by SPCTs combined with LFLIU.

Penetration effect of SPCTs combined with ultrasound in vitro

As mentioned previously, transfersomes are superior to conventional liposomes or no carriers in transdermal drug delivery applications. However, after SPCTs reach the dermis, the absence of a hydration gradient and small differences in water content within the dermis, further penetration lacks sufficient driving force and delivery efficiency decreases. Here, we conducted transdermal administration using Ts or SPCTs combined with LFLIU to investigate the relative ultrasound effect. CdTe-DiO-FU transfersomes (CDF@Ts) or CDF@SPCTs were loaded on the surface of keloids for 40 min. Subsequently, after 10 min of irradiation using LFLIU, the keloid tissues were longitudinally cut, and the fluorescence distribution was observed (Fig. 4A). The fluorescence entry depth and intensity distribution for the CDF@Ts combined with LFLIU irradiation for 10 min (CDF@Ts + LFLIU-10 min) group and the CDF@SPCTs combined with LFLIU irradiation for 10 min (CDF@SPCTs + LFLIU-10 min) group were analyzed statistically. Both groups observed red and green fluorescence in the dermis, and the depth of fluorescence penetration increased on the basis of passive penetration, indicating that ultrasound facilitates further penetration of Ts and SPCTs. Compared with the CDF@Ts + LFLIU-10 min group, the red fluorescence in the CDF@SPCTs + LFLIU-10 min group penetrated significantly deeper into the dermis (Fig. 4B). It exhibited more vigorous fluorescence intensity and a broader distribution area. The effect of CDF@SPCTs combined with ultrasound to penetrate the dermis was significantly better than that of CDF@Ts. Moreover, no significant difference was observed in the green fluorescence distribution depth compared to the red fluorescence distribution depth of the CDF@Ts + LFLIU-10 min group (dashed line marked area, Fig. 4B). However, the depth of green fluorescence distribution in the CDF@SPCTs + LFLIU-10 min group was significantly shallower than that of the red fluorescence distribution. The results confirmed that the PFP vaporization resulted in the rupture of SPCT, and the microjets generated during the process pushed the contents deeper into the dermis. Fig. 4C manifests that the fluorescence penetration depth of the CDF@SPCTs + LFLIU-10 min group was about 485 μm, while the depth of the CDF@Ts + LFLIU-10 min group was about 310 μm. Fig. 4D displays that the CDF@SPCTs + LFLIU-10 min group exhibited a fluorescence intensity of 26.12 ± 2.82 pixels within the dermis range. However, the fluorescence intensity of the CDF@Ts + LFLIU-10 min group under the same conditions was 8.12 ± 0.6 pixels. These statistical results indicated that the penetration depth and amount of SPCTs combined with ultrasound in keloids are significantly better than that of Ts.

Fig. 4.

In vitro penetration depth combined with ultrasound. (A) Schematic diagram of passive permeation combined with ultrasound administration. (B) Fluorescence distribution images of CDF@Ts and CDF@SPCTs after 40 min of passive penetration and 10 min of ultrasonic irradiation. (C) Fluorescence entry depth and (D) fluorescence intensity into keloids for the above two groups (n = 3, mean ± SD). (E) The depth of penetration into the keloid of a series of LFLIU irradiation times combined with CDF@Ts or CDF@SPCTs. (F) Fluorescence distribution images of CDF@SPCTs in a keloid after 40 min of passive penetration followed by 40 min of ultrasonic irradiation. Scale bar = 100 μm

We aimed to determine the optimal duration of ultrasound irradiation for combination with SPCTs, considering the time-dependent controlled release effect of SPCTs. We tested different ultrasound irradiation times (10, 20, 30, 40, 50, and 60 min) for the Ts or SPCTs in keloid tissues, made longitudinal slices of the tissues, and conducted statistical analyses to evaluate the effect on dermal penetration. We found that the penetration depth into the keloid of SPCTs combined with LFLIU was higher than that of Ts combined with LFLIU at each selected ultrasound irradiation duration (Fig. 4E). The penetration depth in the SPCTs combined with LFLIU group rapidly increased as the irradiation time was extended from 10 to 40 min. However, the growth rate of penetration depth decreased as the ultrasonic irradiation time was increased from 40 to 60 min. This is consistent with the findings of previous controlled-release effect experimental results. In the group of Ts combined with LFLIU, the increased amplitude of penetration depth was relatively slow.

This finding indicates a positive correlation between the depth of content entry and the duration of ultrasound irradiation and confirms that combining SPCTs with ultrasound is beneficial for the penetration depth into keloids. Furthermore, the research findings indicated that within 40 min of ultrasound combined with SPCTs, ultrasound enhanced the penetration effect of SPCTs in the dermis layer. This was mainly due to the cavitation effect of ultrasound, which caused a phase transition to form microbubbles in the PFP of SPCT. The ultrasound pushed SPCTs deeper into the dermis before the microbubbles ruptured, followed by the gradual rupture of the SPCTs, which facilitated the sustained diffusion and release of the drug deeper into the dermis under the action of microjets. After 40 min, as many SPCTs had ruptured, the primary effect was due to ultrasound alone, which continued to move the contents into the deep dermis. However, since the effect of ultrasound alone is weaker than the synergistic effect of ultrasound and SPCTs, the increase in penetration depth slowed down. From the results, it can be seen that most SPCTs experienced an ultrasound response within 40 min. The longitudinal sections indicated that the red fluorescence penetrated to a maximum depth of 2,000 μm in the keloid (Fig. 4F). H&E staining of the isolated keloids revealed that the combined action did not cause structural damage to the tissue and that the safety characterization was confirmed (Figure S4). Consequently, 40 min was an appropriate choice as the application time of ultrasonic irradiation in subsequent experiments.

Ts or SPCTs were combined with LFLIU to conduct transdermal drug delivery to further clarify the distribution of delivery substances at different depths of the keloid dermis. The study was divided into two groups: one group had CDF@Ts loaded onto the surface of the keloid, left for 40 min and irradiated under LFLIU for 40 min (CDF@Ts + LFLIU-40 min), and the other group had CDF@SPCTs loaded onto the surface of the keloids for 40 min and irradiated under the same LFLIU for 40 min (CDF@SPCTs + LFLIU-40 min). Subsequently, the keloid tissues were sectioned transversely. The findings indicated that the fluorescence distribution in the CDF@SPCTs + LFLIU-40 min group was significantly broader and more abundant at the same depth of horizontal slices. The fluorescence distribution was more apparent in horizontal slices from 300 to 2,000 μm depth (Fig. 5A-B). Further quantitative analysis was conducted on the fluorescence intensity of each horizontal slice at the same depth. At each depth, the fluorescence intensity of the CDF@Ts + LFLIU-40 min group (Fig. 5C) was significantly lower than that of the CDF@SPCTs + LFLIU-40 min group (Fig. 5D). For the CDF@Ts + LFLIU-40 min group, the peak fluorescence intensity was observed at a depth of approximately 300 μm. However, the mean gray value of the CDF@SPCTs + LFLIU-40 min group revealed a peak fluorescence intensity at approximately 800 μm, which was significantly relatively deeper. The pharmaceutical content of 5-FU that entered the two groups of keloid tissues was detected using HPLC. According to the HPLC standard curve of 5-FU (Figure S5), the drug content in the tissues of the CDF@Ts + LFLIU-40 min group was 273.34 µg. However, the drug content in the tissues of the CDF@SPCTs + LFLIU-40 min group was 533.34 µg, which was twice that of the CDF@Ts + LFLIU-40 min group (Fig. 5E). The drug release curves showed that the drug release rates of both Ts and SPCTs increased with the extension of ultrasound irradiation time, showing a significant upward trend (Fig. 5F). Under the same ultrasound irradiation, the cumulative drug release rate of the 5-FU@SPCTs + LFLIU-40 min group was 39.31% ± 2.54%, which was 2.08 times higher than that of the 5-FU@Ts + LFLIU-40 min group. This also suggests that PFP in SPCTs plays a key role in further improving the drug delivery efficiency through synergistic application with ultrasound. In summary, SPCTs combined with ultrasound can promote more drugs to penetrate deeper into the skin.

Fig. 5.

In vitro distribution at different depths and pharmaceutical contents. Fluorescence distribution at different depths of keloid tissues after (A) CDF@Ts or (B) CDF@SPCTs were left to stand for 40 min and then irradiated by ultrasound for 40 min (horizontal sections). Scale bar = 200 μm. The fluorescence intensity corresponds to different depths within the keloid tissues after administration of (C) CDF@Ts or (D) CDF@SPCTs. (E) The amount of 5-FU drug accumulated in keloid tissues by both drug carriers after 40 min of passive penetration and then 40 min of ultrasound irradiation (n = 3, mean ± SD). (F) The drug release curves of 5-FU@Ts and 5-FU@SPCTs after 40 min of passive penetration followed by ultrasound irradiation

Penetration effect of SPCTs combined with ultrasound in vivo

We constructed an animal model by transplanting human keloid tissue onto the backs of nude mice (Figure S6A-B). The transplanted human keloid tissue was 2.5 mm in height and retained all the tissue layers of the human keloid: the epidermis, papillary dermis, and reticular dermis. Seven days after the keloid nude mouse model was established, a four-week sampling period was implemented, with samples taken once a week on weekdays for H&E staining sections. The distribution of keloid tissue can be identified by the clear and transparent collagen clusters in the H&E staining results (indicated by red dashed lines; Figure S6C). One week later, no gap was observed between the keloid tissue and the nude mouse skin, indicating the successful construction of a model for transdermal drug delivery to keloid tissue. Weekly sampling and H&E staining observations were conducted for the subsequent 21 days. The findings revealed that the keloid tissue and the skin of the nude mice remained closely articulated, and the volume of the keloid tissue did not shrink, confirming the successful maintenance of the model for more than 28 days. We followed the regimen outlined in Fig. 6A to evaluate the in vivo effect of SPCTs combined with ultrasound. 5-FU@Ts and 5-FU@SPCTs containing only 5-FU were prepared for subsequent experiments. The in vivo experiments were divided into four groups: untreated (Control), 5-FU@SPCTs passively permeated for 40 min without ultrasound (5-FU@SPCTs), 5-FU@Ts combined with 40 min of LFLIU (5-FU@Ts + LFLIU-40 min), and 5-FU@SPCTs combined with 40 min of LFLIU (5-FU@SPCTs + LFLIU-40 min). The mice received the transdermal drug delivery treatment twice weekly for four weeks (28 days). After 28 days of transdermal drug treatment, significant large keloid tissue persisted in the Control, 5-FU@SPCTs and 5-FU@Ts + LFLIU-40 min groups, while only sporadic keloid tissue remained in the 5-FU@SPCTs + LFLIU-40 min group. The 5-FU@SPCTs + LFLIU-40 min group exhibited the most significant reduction in keloid tissue and the best treatment effect (Fig. 6B). As shown in Fig. 6B-C, the volume of the 5-FU@SPCTs + LFLIU-40 min group decreased fastest compared with the other three groups. When measuring volume, only the surviving keloid tissue below was measured. After 28 days of treatment, the remaining volume of keloids in the Control, 5-FU@SPCTs and 5-FU@Ts + LFLIU-40 min groups was 59.75% ± 6.98%, 34.3% ± 6.25% and 20.23% ± 3.5%, respectively. However, the keloid volume in the 5-FU@SPCTs + LFLIU-40 min group was only 8.46% ± 5.46%, significantly reducing the keloid volume (p < 0.01).

Fig. 6.

In vivo penetration effect and safety evaluation. (A) Schedule for transdermal drug delivery therapy in nude mouse keloid model. (B) Images of keloid changes in nude mice over 28 days after treatment with the following four groups: Control, 5-FU@SPCTs, 5-FU@Ts + LFLIU-40 min, and 5-FU@SPCTs + LFLIU-40 min. (C) Volume variation curves of keloids in the four groups over 28 days. (D) H&E staining of keloids from the above four groups. Scale bar = 500 μm

In addition, we evaluated the safety of the transdermal drug delivery in various aspects in vivo. The safety of ultrasound irradiation was assessed, and real-time temperature measurement revealed that the body temperature of the nude mice remained within the safe temperature (42℃) for 60 min of applying ultrasound without thermal damage (Figure S7) [81]. After 28 days of treatment, H&E staining of the above four groups revealed a clear tissue structure of the skin of keloid nude mice; no tissue defects on the surface were observed, the cells were orderly arranged, the cell nuclei were homogenously stained, and no other abnormalities were observed, indicating that the treatment did not cause significant damage to the skin of the nude mice (Fig. 6D).

We used HaCaT cells to assess the cytotoxicity of unloaded Ts and unloaded SPCTs in conditions with and without ultrasound by CCK-8 assay. The cell survival of both samples was above 80% in both conditions, indicating their biocompatibility (Fig. 7A-B). Furthermore, 5-FU decreases the volume of keloid tissue by inhibiting cell proliferation and collagen synthesis (primarily collagen I and III) [82–84]. We further identified the expression levels of relevant biomarkers, collagen proteins. The proliferative activity of fibroblasts in keloid tissues was observed by detecting the red fluorescence of Ki-67 (a marker of cell proliferation) in keloid fibroblasts across the four groups after four weeks of administration. The 5-FU@SPCTs + LFLIU-40 min group exhibited the weakest fluorescence intensity compared with the other three groups (Fig. 7C). The quantitative fluorescence intensity analysis indicated that Ki-67 expression was significantly down-regulated in the 5-FU@SPCTs + LFLIU-40 min group (Fig. 7F). These findings indicate that the 5-FU@SPCTs + LFLIU-40 min group exhibited the lowest fibroblast proliferative activity, confirming its highest drug penetration and the best therapeutic effect on keloid tissue. The protein concentration of each experimental group after administration was calculated from the BCA protein standard curve (Figure S8). The relative expression levels of type I and III collagens after drug administration showed a reduction in the expression of both collagens in the keloid tissues of the other three treated groups compared with the Control group, with the 5-FU@SPCTs + LFLIU-40 min group exhibiting the most significant reduction (Fig. 7D-E and G-H). This confirmed that the 5-FU@SPCTs + LFLIU-40 min group for transdermal delivery of 5-FU was the most effective, leading to the fastest reduction in keloid volume. In addition, our weekly analysis of the relative expression levels of type I and III collagens in keloid tissues revealed that the protein expression of both types was down-regulated in the 5-FU@SPCTs + LFLIU-40 min group compared with the Control group (without treatment) (Figure S9). In summary, the 5-FU@SPCTs + LFLIU-40 min group demonstrated the best therapeutic effect in terms of inhibiting cell proliferation activity and collagen synthesis.

Fig. 7.

Cell viability, biomarker and protein expression. (A) HaCaT cell viability after treatment with Ts or SPCTs, relative to the control group. (B) HaCaT cell viability in the Control group, and in groups treated with Ts or SPCTs, after exposure to LFLIU for 40 min. (C) Immunofluorescence of keloid tissue in four groups after four weeks of treatment: Control, 5-FU@SPCTs, 5-FU@Ts + LFLIU-40 min, and 5-FU@SPCTs + LFLIU-40 min. Scale bar = 100 μm. (D) The protein expression levels of collagen I after 4 weeks of treatment. (E) The protein expression levels of collagen III after 4 weeks of treatment. (F) Analysis of the fluorescence intensity (n = 3, mean ± SD). (G, H) Gray value analysis of protein expression differences corresponding to collagen I and collagen III (n = 3, mean ± SD)

Conclusion

This study proposed a novel transfersome, specifically a phase-change transfersome. The sono-phase-change transfersomes (SPCTs) were combined with LFLIU to form a two-stage transdermal drug delivery system. This system can utilize the SPCT deformability to achieve passive administration in the epidermis. However, ultrasonic irradiation is applied once SPCTs have penetrated the epidermis to control their phase-change to form microbubbles. The rupture of SPCT microbubbles caused microjets and mechanical friction effects, ensuring an active administration effect. We successfully prepared the SPCTs loaded with PFP, which exhibited good deformability and a particle size of 240.36 ± 1.38 nm. They can stably and passively penetrate the stratum corneum and enter the epidermis, maintaining stability at epidermal temperature. Under the selected LFLIU parameters (360 kHz, 1 W), controlled release was achieved during ultrasound irradiation, and the contents were released into the deep dermis through active administration (up to 2,000 μm). Notably, the analysis of in vitro and in vivo permeability characteristics revealed that SPCTs combined with ultrasound achieved more effective and deeper penetration and more extensive distribution in the dermis of keloid tissues than using only the Ts or the combination of the Ts with ultrasound. This was achieved with a significantly increased drug dosage, improved treatment efficiency, and no damage to the skin. Our study preliminarily demonstrated that the two-stage transdermal drug delivery system formed by SPCTs and LFLIU can achieve non-invasive and deep dermal drug delivery of 5-FU in keloid, laying the foundation for the subsequent application of this system in transdermal delivery of other drugs and diseases.

Author contributions

Z.Y., H.X. completed in vitro and in vivo experiments, data collection, and finished the original draft. M.X. participated in the original draft of the article and data analysis. Y.Y.K. participated investigation and image visualization. L.F.Q., H.X.F., L.H., Y.Y.J., and W.Y. are involved in research supervision and methodology. T.L. and D.Y.L. participated in in vitro experiments and validation. W.Z.Y. and X.W. contributed to the conceptualization, article revision. W.Z.Y., X.W. and W.Y. participated in funding acquisition of this study. All authors read and approved the final manuscript.

Funding

This work was supported by the National Natural Science Foundation of China (Grant NO. 81501617), the Special Funds for Technology Innovation and Application Development of Chongqing, China (Grant NO. cstc2020jscx-msxmX0064), the Program for Youth Innovation in Future Medicine, Chongqing Medical University (W0155), and Science grants from the Chongqing government (Grants NO. JBGS2024-004, 2023ZDXM033, YXGD202465, CSTB2023JXJL-YFX0028).

Data availability

No datasets were generated or analysed during the current study.

Declarations

Ethics approval and consent to participate

All experimental procedures involving animals followed the protocol reviewed and approved by the Laboratory Animal Welfare and Ethics Committee of Chongqing Traditional Chinese Medicine Hospital (No. 2024-DWSY-XW). The keloid samples were obtained after the patients provided written informed consent, and the procedure was approved by the Ethics Committee of the Chongqing Traditional Chinese Medicine Hospital (No. 2024-KY-HY-40).

Consent for publication

All the authors agree with the publication.

Competing interests

The authors declare no competing interests.