Size-dependent effects of niosomes on the penetration of methotrexate in skin layers

Abstract

Background

Niosomes hold promise as drug delivery systems for cancer treatment, with niosome size impacting stability, biodistribution, and effectiveness. This study optimized methotrexate (MTX)-loaded niosome formulation by studying the effects of components and processing conditions on size. The niosomes formulation was made by the thin-film hydration technique.

Results

The optimized formulation (NIO 17) with a 6:2:2 ratio of span 60, soya PC, and cholesterol achieved 55.05% methotrexate encapsulation, particle size 597.2 nm, PDI 0.49, and zeta potential − 23.3 mV. The compatibility of methotrexate with lipids was confirmed via Fourier transform infrared spectroscopy, and transmission electron microscopy revealed spherical, well-dispersed vesicles. Differential scanning calorimetry indicated methotrexate conversion or entrapment within vesicles. In vitro release exhibited a sustained pattern with an initial burst. NIO 17 showed potent anti-cancer activity against B16-F10 cells (GI50: 38.7176 μg/mL). Ex vivo studies suggest tailoring niosome size (597.2–982.3 nm) to target specific skin depths (0–38 μm) for enhanced localized drug delivery.

Conclusions

This study demonstrates the potential of methotrexate-loaded niosomes as a novel cancer therapy approach, highlighting the potent anti-cancer activity and transdermal delivery potential of NIO 17. Further research is necessary to explore its clinical translation.

Graphical abstract

Background

Skin cancer, triggered by ultraviolet (UV) radiation from the sun or tanning beds, affects millions globally [1, 3]. Common types include basal cell carcinoma, squamous cell carcinoma, and melanoma as shown in Fig. 1. Traditional treatments like surgery, radiation therapy, and chemotherapy pose various limitations [2, 3]. Surgery, while common for early stages, is invasive. Radiation uses high-energy beams, and chemotherapy, despite killing cancer cells, often comes with side effects like nausea, vomiting, hair loss, and fatigue [4, 6]. Moreover, advanced or metastatic cases often lack effective treatment options. There is a growing demand for targeted and minimally invasive therapies that can deliver drugs specifically to cancerous cells while sparing healthy tissue. In this context, exploring novel drug delivery systems like niosomes holds great promise. Niosomes, with their ability to encapsulate drugs and deliver them to specific sites within the body, offer the potential to enhance the efficacy of treatments for skin cancer while minimizing adverse effects, thus addressing critical unmet needs in the current treatment landscape. Niosomes, composed of biocompatible and biodegradable phospholipids, cholesterol, and surfactants [5, 7], niosomes can deliver a wide range of drugs and target specific cells or tissues. Studies in mice suggest their safety and efficacy for skin cancer treatment, warranting further exploration for clinical use.

Fig. 1

Schematic representation of SCC, BCC and malignant melanoma

Methotrexate (MTX), a drug commonly used for rapidly growing tumors like skin cancer [5], works by hindering DNA synthesis in dividing cells. While traditionally administered orally or parenterally for skin cancer, MTX can cause gastrointestinal side effects like indigestion, dyspepsia, vomiting, and ulceration. To address these, we investigated the potential of niosomes as a topical delivery system for MTX [7].

This study aimed to evaluate niosomes as a means to enhance the transdermal transport of MTX to various skin layers, focusing on the impact of niosome size on their efficacy as shown in Fig. 2. We examined the effects of different-sized niosomes on MTX permeation through ex vivo studies and visualized their distribution within the skin using confocal laser scanning microscopy. Additionally, in vivo histopathological studies in mice assessed the feasibility of using niosomes to deliver MTX to the skin.

Fig. 2

Schematic representation of niosomes penetration by skin layers

Materials and protocols

Chemicals

This research utilized a diverse range of materials for its experiments. The drug, methotrexate (MTX), was kindly provided by Neon Laboratories Ltd. (Mumbai, India). Soya Phosphatidylcholine (SPC), a key lipid component, was a generous gift from Lipoid GmbH (Ludwigshafen, Germany), while cholesterol (CHOL) was purchased from CDH (India). Solvents and reagents like chloroform, methanol, phosphate buffer, Triton X-100, Sephadex G-50, and dimethyl sulfoxide (DMSO) were sourced from HiMedia Laboratory Pvt. Ltd. (Mumbai, India). Additionally, HPLC water (Lichrosolv), acetonitrile (HPLC grade), methanol (HPLC grade), and rhodamine B (RHB) were procured from Sigma-Aldrich and Merck (India). Finally, dialysis bags with molecular weights ranging from 12,000 to 14,000 were obtained from HiMedia Laboratory Pvt. Ltd. Notably, all chemicals were of analytical grade and used directly without further purification.

Preparation of MTX-loaded niosomes

The preparation of various niosomal formulations (Nio1 to Nio18) was carried out using the thin-film hydration method but with slight modifications [8, 9], where the mole fraction of Span 60 was systematically varied (4:2:2, 5:2:2, 6:2:2 Soya PC: Cholesterol ratios) while maintaining constant molar concentrations of Soya phosphatidylcholine (SPC) and cholesterol as shown in Fig. 3. An accurate amount of Span 60, soya phosphatidylcholine, and cholesterol was dissolved in a round-bottom flask (RBF) containing 10 mL of chloroform and methanol (9:1). The organic solvents were eliminated using a rotary vacuum evaporator above the lipid transition temperature of 65 °C under reduced pressure to form a thin lipid film on the wall of the RBF. The residual amount of the solvent mixture was removed from the deposited layer of lipids by leaving the contents under a vacuum overnight. Hydration was done with phosphate buffer (PBS 7.4) containing methotrexate drug by rotating the RBF at a suitable temperature for 2 h until the formation of niosomes [22, 23]. The particle size of the formed niosomes dispersion was further reduced by sonication with various sonication cycles. The sonication time is taken as another variable for the formulation process. The same method was used to prepare Rhodamine B-loaded niosomes for the ex vivo study, where 0.5% of the total lipid was substituted with RHB in place of MTX in the selected formulation [10, 24, 25]. The obtained opalescent dispersion of niosomes was stored at 4 ± 2 °C until use.

Fig. 3

Method of preparation of niosomes

Characterization of Niosomal formulations

Size distribution, polydispersity, and surface charge of vesicles

Initially, 1 ml of each of the prepared niosomal formulations was mixed with deionized water for adequate dispersion. The average size of the vesicles (VS), the zeta potential (ZP), and the polydispersity index (PDI) of the drug-loaded niosomes was analyzed using Zeta sizer Nano ZS (Malvern Instruments Ltd.) dynamic light-scattering method. The niosomal dispersions were prepared and diluted 10 times with deionized water before measurement to ensure that the light-scattering intensity was within the instrument's detection range [26, 27]. The measurements were taken in triplicate, and the average values are reported in Table 1 and Fig. 4.

Table 1 The formulations of niosomes with different ratios of ingredients and sonication time, along with the results of particle sizes, zeta potential, PDI, and entrapment efficiency

Fig. 4

A Graphical representation of formulation optimization with ratio 6:2:2 (Span 60:CH:SPC); a vesicle size (nm); b PDI; c Zeta Potential (mV); d entrapment efficiency (%). B Images showing the particle-size distribution of the optimized formulations (NIO 13, 14, 17, and 18) prepared using a ratio of Span 60:CH:SPC (6:2:2). C Images showing the zeta potential of the optimized formulations (NIO 16 and 17) prepared using a ratio of Span 60:CH:SPC (6:2:2)

Fig. 5

Fluorescent and TEM images of niosomes

Entrapment efficiency (EE)

The entrapment efficiency (EE) was calculated through the mini-column centrifugation method by separating the non-entrapped drug. The %EE of MTX in the niosomes was estimated by determining the free MTX in the dispersion medium. 1 mL of the niosomes dispersion was centrifuged at 15,000 rpm for 2 h at 4 °C using a cooling centrifuge. The supernatant was then separated and diluted, and the concentration of MTX was determined through HPLC methods [11, 28, 29]. The niosomes were then disrupted using 0.1% Triton-X 100, and the drug content is quantified using Eq. (1). An isocratic HPLC procedure was used to determine the concentration of MTX. A Zorbax Extend-C18 column was used as the stationary phase, and the mobile phase was a mixture of 50 mM sodium acetate buffer solution with a pH of 5.6 and acetonitrile (89:11 v/v). The flow rate was set at 1.0 mL/min, and the UV detector was set at 307 nm. The results are shown in Table 1 and Fig. 4.

$$\text{Entrapment Efficiency} \left(\%\right)= \frac{\text{Amount of free drug}}{\text{Total amount of drug}} \times 100$$

(1)

Shape and surface morphology

The appearance of the optimized MTX-loaded niosomal formulations was studied using the transmission electron microscopy (TEM) technique (Jeol JEM1230, Tokyo, Japan) after suitable dilution. A drop of the selected niosomal dispersion was spread on a copper grid and stained negatively with 1% phosphotungstic acid. The samples were then air-dried for 10 min at standard temperature and pressure (STP) [14, 30, 31] and examined under a TEM as shown in Fig. 5.

Fig. 6

A FTIR spectra of methotrexate, Soya PC, Span 60, Cholesterol and Mixture; B DSC spectra of methotrexate, Soya PC, Span 60, cholesterol and mixture

Fourier transform infrared spectroscopy

The optimized formulation, drug, excipient, and drug–excipient mixture were analyzed using Fourier transform infrared spectroscopy (FTIR) (Bruker FTIR 8400S ALPHA). Before conducting the IR studies, the samples and excipients were vacuum-dried for 12 h [32, 33]. The dried samples of each were placed on a sample platform, and the spectra were captured in the range of 4000–400 cm⁻¹, which were consistent with the official IR spectrum reported in the monograph as shown in Fig. 6A [12, 13].

Fig. 7

In vitro release of MTX from niosomes in comparison to plain MTX solution at pH 7.4

Thermoanalytical technique

The thermal characteristics of pure methotrexate (MTX), Soya Phosphatidylcholine (PC), Cholesterol, Span 60, the physical mixture of MTX and niosomal components, empty niosomes, and MTX-loaded niosomes were analyzed using differential scanning calorimetry technique (DSC 60, Shimadzu, Japan). The device was calibrated using high-purity indium (99.9%). Three milligrams of accurately weighed samples were put into standard aluminum pans and then heated from 100 to 300 °C at a scanning rate of 100 °C per minute [15, 34, 35] as shown in Fig. 6B.

In vitro drug release kinetics

The niosomal suspension (Nio 15 and Nio 16) was centrifuged to separate the unentrapped drug from the niosomes. Next, 1 mL of the pure niosomal suspension was placed into a dialysis tube, which was then immersed in a beaker that contained 20 mL of PBS (pH 7.4) and stirred on a magnetic stirrer at a constant temperature of 37 ± 0.5 °C. Samples were taken at various time intervals and analyzed for drug content using HPLC. To maintain the initial volume of the dissolution fluid, 5 mL of fresh solution was added after each sample withdrawal [15, 20, 36, 37]. The cumulative amount of MTX released from different niosomal formulations was calculated as per Eq. (2). Results are shown in Fig. 7.

$$\text{Percentage cumulative drug released }= \frac{\text{Cumulative Amount of Drug obtained at time}}{\text{Total amount of Drug in Niosomes}}$$

(2)

In vitro cytotoxicity study

In vitro, cell cytotoxicity of optimized formulation (Plain MTX, NIO 5, NIO 12, and NIO 16) was evaluated by the SRB assay using murine skin melanoma cell line B16-F10. The cell lines were grown in RPMI 1640 medium, and cells were inoculated into 96-well plates in 100 µL at plating [38,39,40]. After cell inoculation, the microtiter plates were placed in an incubator at 37 °C, 5% CO₂, 95% air, and 100% relative humidity for 24 h before the addition of formulations. Formulations were dissolved in an appropriate solvent to a concentration of 100 µg/mL and then diluted to 1 mg/mL using Milli-Q water. The diluted solutions were stored frozen until needed. At the time of the experiment, aliquots of the frozen concentrate (1 mg/mL) were thawed and diluted to the following concentrations: 100 μg/mL, 200 μg/mL, 400 μg/mL, and 800 μg/mL. Aliquots of 10 μL of each drug dilution were added to microtiter wells that already contained 90 μL of the medium. This resulted in final drug concentrations of 10 μg/mL, 20 μg/mL, 40 μg/mL, and 80 μg/mL. The plates were then incubated at standard conditions for 48 h.

Fig. 8

A Growth curve: the plot of percentage growth inhibition versus drug concentration (μg/mL) shows the effective drug concentration on murine tumor cell line B16 F10 cells; B Phase-contrast microscopic images of murine tumor cell line B16 F10 cells after 24 h. treatment negative control (untreated), ADR, and formulations NIO 14 to NIO 17

To terminate the assay, cold TCA was added to the plates. Cells were fixed in situ by gently adding 50 μL of cold 30% (w/v) TCA (final concentration, 10% TCA) [41, 42]. The plates were then incubated for 60 min at 4 °C. The samples were washed five times with PBS, and then, 50 μL of 0.4% (w/v) SRB solution was added to each well. The plates were then incubated for 20 min at room temperature.

One percent (v/v) of acetic acid was used to remove the unbound dye, and plates were dried and then analyzed at the wavelength of 540 nm with a 690 nm reference wavelength by using the plate reader. Adriamycin (ADR) was used in the same concentration range as a positive control. Cytotoxicity was expressed as the percentage of viable cells relative to incubated cells in the presence of 0.1% DMSO vehicle control. The percentage of growth inhibition (GI₅₀) is calculated in Eq. (3). GI₅₀ was calculated from the drug concentration, resulting in a 50% reduction in cell growth. Each measurement was done in triplicate [18, 19, 43]. Results shown in Fig. 8.

$${\text{Percentage growth inhibition }} = \, \left( {{\text{Ti}} - {\text{Tz}}} \right)/\left( {C - {\text{Tz}}} \right) \, \times 100$$

(3)

where absorbance at zero time (Tz), control growth (C), and test growth of formulation in the presence of drug at four concentration levels (Ti).

Fig. 9

A Confocal laser scanning photomicrographs illustrating a z-stack image capturing optical sections of mice skin. The sections were taken at intervals of 2 µm, covering a depth range from 0 to 38 µm. The formulations employing PBS as a positive control exhibit no observable particles. B Confocal laser scanning photomicrographs illustrating a z-stack image capturing optical sections of mice skin. The sections were taken at intervals of 2 µm, covering a depth range from 0 to 38 µm. The formulations employing Plain Rhodamine as a negative control exhibit no observable particles. C Confocal laser scanning photomicrographs illustrating a z-stack image capturing optical sections of mice skin. The sections were taken at intervals of 2 µm, covering a depth range from 0 to 14 µm. The formulations using Nio13 (Without Sonication) as an agent demonstrate the presence of particles, visible within the range of 0–6 µm. D Confocal laser scanning photomicrographs illustrating a z-stack image capturing optical sections of mice skin. The sections were taken at intervals of 2 µm, covering a depth range from 0 to 14 µm. The formulations using Nio14 (2 min Sonication) as an agent demonstrate the presence of particles, visible within the range of 4–18 µm. E Confocal laser scanning photomicrographs illustrating a z-stack image capturing optical sections of mice skin. The sections were taken at intervals of 2 µm, covering a depth range from 0 to 14 µm. The formulations using Nio15 (4 min Sonication) as an agent demonstrate the presence of particles, visible within the range of 16–22 µm. F Confocal laser scanning photomicrographs illustrating a z-stack image capturing optical sections of mice skin. The sections were taken at intervals of 2 µm, covering a depth range from 0 to 40 µm. The formulations using Nio16 (6 min Sonication) as an agent demonstrate the presence of particles, visible within the range of 24–32 µm. G Confocal laser scanning photomicrographs illustrating a z-stack image capturing optical sections of mice skin. The sections were taken at intervals of 2 µm, covering a depth range from 0 to 38 µm. The formulations using Nio17 (8 min Sonication) as an agent demonstrate the presence of particles, visible within the range of 12–38 µm

Ex vivo and in vivo investigations

The ex vivo investigation was conducted following protocols approved by the Committee for Control and Supervision of Experiments on Animals (CPCSEA/SN-10/RN-22/16-02-2020), which is overseen by the Ministry of Social Justice and Empowerment, Government of India. The study also followed the recommendations of the Institutional Animal Ethical Committee (IAEC) at Dr. H.S. Gour Vishwavidyalaya, located in Sagar, Madhya Pradesh, India.

Ex vivo permeation study

The ex vivo visualization using confocal laser scanning microscopy technique (CLSM) involved the formulation of Rhodamine B (RH)-loaded niosomes through a process similar to that used for creating MTX-loaded niosomes [44,45,46]. However, RH was incorporated at a concentration of 0.5% concerning the total lipid content, replacing MTX in the specific formulation. For experimentation, the mice's skin was sectioned into square pieces and positioned with the stratum corneum (SC) facing the donor compartment within diffusion cells. The receptor chamber contained 20 mL of phosphate buffer saline (pH 7.4) maintained at 32 ± 1 °C. To mimic the expected application of niosomes to the skin's surface, RH-loaded niosomes (including a plain niosomes formulation, a plain rhodamine solution, and formulations Nio 13 to Nio 17) were administered onto the skin and allowed to remain for 4 h. Following this, the skin was rinsed with 10% ethanol and then gently wiped in preparation for imaging, following the methods described by [16, 47]. The investigation of the skin was conducted using CLSM with the following steps: the complete skin sample was positioned between a glass slide and a cover slip and examined utilizing inverted CLSM (LSM 710, Carl Zeiss, Jena, Germany). RH fluorescence was excited at a wavelength of 573 nm and detected at 591 nm. Scans were taken at 2-µm intervals starting from the skin surface (0 mm) to a depth of 40 µm, using a 40× objective lens. Images shown in Fig. 9 were captured in both the xy and xz planes, employing optical sectioning z-stack mode. The acquired confocal microscopy images were processed using LSM Image Browser software, release 4.2 (LASX), as described by Hathout and Nasr in 2013 [17, 48].

Fig. 10

Photomicrographs showing histopathological sections (hematoxylin and eosin stained) of mice skin A normal untreated, B treated with Plain MTX, C Nio13, D Nio14 E, F Nio15, G, H Nio16 and I: Nio17. The magnification power of 40× to identify the epidermis, dermis, the subcutaneous tissue and muscles, respectively

In vivo histopathological study

An In vivo histopathological study was performed to evaluate skin irritation potential and structural changes resulting from exposure to MTX-loaded niosomes. The study involved dividing mice into 7 groups, each consisting of 3 animals [49, 50]. Group I acted as the positive control receiving PBS, group II as the negative control receiving plain MTX solution, and groups III to VII were exposed to topical applications of MTX-encapsulated niosomes (Nio 13 to 17) on the skin surface three times daily for a week. After the treatment period, the mice were killed, and their skin was extracted for histopathological examination. The collected skin samples underwent fixation in 10% formal saline for 24 h, followed by washing, dehydration using alcohol dilutions, clearing in xylene, and embedding in paraffin beeswax blocks. These blocks were maintained at 56°C for 24 h, and sections of 4 mm thickness were cut using a microtome (Leica Microsystems SM2400, Cambridge, England). These sections were then deparaffinized, stained with hematoxylin counterstained with eosin, and subsequently observed under a light microscope for analysis as shown in Fig. 10 [9, 21, 51].

Results and discussion

Preparation of niosomal formulation, particle-size analysis, and zeta potential determination

The synthesis of MTX-loaded niosomes was achieved using the film hydration method as depicted in Fig. 3 and by incorporating various ratios of span 60, soya PC, and cholesterol in each formulation. The study aimed to determine the effect of various ingredients and processing variables on the vesicle size and entrapment efficiency. The composition of span 60, soya PC, and cholesterol (4:2:2, 5:2:2, 6:2:2) had a significant impact on the size of the niosomes. The results showed that increasing the concentration of Span 60 (a non-ionic surfactant with an HLB of 4.7) led to a decrease in vesicle size. This is because Span 60 has high interfacial activity and a critical packing parameter (CPP) between 0.5 and 1. A lower CPP indicates that the surfactant molecules are more tightly packed, which results in smaller vesicles. However, a further increase in the concentration of Span 60 led to an increase in vesicle size. This is because the higher concentration of cholesterol in the vesicles provided added rigidity, which counteracted the effect of the increased interfacial activity of Span 60. The addition of cholesterol in lipid vesicles changes the short-distance repulsive forces among them. An increase in cholesterol content leads to an increase in the net repulsive forces among soya PC and span 60 vesicles, reducing their aggregation. The sonication time also affects the entrapment efficiency and size of MTX-loaded niosomes, as seen in Table 1. An increase in sonication time causes a decrease in particle size. Particles without sonication showed the largest size, while particles subjected to 10 min of sonication showed the smallest size, with a mean particle size ranging from 200 to 1406 nm. The optimized formulations, with a ratio of 6:2:2, were used for further studies. The average standard deviation (SD) (nm) of particle size for most of the particles was 200–1000 nm, depending on the sonication time cycle, and the polydispersity index was less than 0.4. The formulations had a negative zeta potential of −25 to −30 mV as depicted in Fig. 4A, B and showed the best entrapment efficiency. The negative zeta potential of the formulations assisted in their adhesion to cancer cells and internalization through cellular endocytosis.

Entrapment efficiency

After removing the unencapsulated drug through dialysis, the efficiency of encapsulation was analyzed for all the formulations. The presence of surfactant, cholesterol, and drug-to-lipid ratios in the vesicles can reduce the stiffness of the vesicles and affect the ability of methotrexate to be trapped inside. The highest drug encapsulation efficiency was recorded to be 76% in vesicles without sonication and the lowest was 30% with a 10-min sonication cycle.

Shape and surface morphology

Confirmation of the results obtained from the Malvern particle-size analyzer can be achieved through morphological analysis, which also examines the structure of colloidal systems. The optimized niosomes displayed a non-aggregated, single-layered vesicular structure with a predominant spherical shape as seen in the TEM images (Fig. 5). The average particle size observed in the TEM micrographs was consistent with the size previously obtained from the Malvern particle-size analyzer, as presented in Table 1.

Fourier transform infrared spectroscopy (FTIR)

FTIR spectroscopy is a useful tool in determining potential interactions between drugs and excipients. Figure 6A shows the IR spectra of pure methotrexate and a formulation that includes the drug and other excipients like phospholipids and a permeation enhancer. The pure methotrexate exhibits peaks at 3406.10 cm⁻¹ (N–H stretching), 2928.64 cm⁻¹ (O–H stretching for the carboxylic group), 1646.82 cm⁻¹ (C–C stretching), 1603.41 cm⁻¹ (C=C stretching of aromatics), 1542.02 cm⁻¹ (N–H bend), and 831.01 cm⁻¹ (C–H out of the plane of aromatics). The peaks for span 60 were found at 2950.00, 2856.50, and 1733.73 cm⁻¹, while peaks for lipids were observed at 3414.06, 2875.67, 1383.09, 1433.18, 1056.43, and 1547.14 cm⁻¹. Peaks for phosphatidylcholine (PC) were recorded at 3467.98, 2926.81, 1474.01, 1214.58, and 1042.76 cm⁻¹. The prepared formulations were also analyzed and found to have peaks in the same range, the drug and the lipid did not interact. The peaks of the optimal formulation were found to be very similar to those of the pure methotrexate, i.e., 3500.00, 3100.00, 2908.60, 1742.64, 1638.10, 1510.00, 1540.21, 1210.12, and 815.00 cm⁻¹, revealing that there was no significant change in the IR peaks of methotrexate and the optimized formulation, signifying the compatibility of the drug and the polymer.

Thermoanalytical technique

Figure 6B displays the DSC (differential scanning calorimetry) thermograms of various substances, including pure methotrexate (MTX), Soya phosphatidylcholine (PC), cholesterol, Span 60, a physical mixture of MTX with niosome components, blank niosomes, and MTX-loaded niosomes. The thermogram of MTX exhibits a typical exothermic peak at 185.9 °C and 93.5 °C, indicating its crystalline form, with a residual mass of 89.79% and weight loss of 7.79%. The thermogram of Span 60 displays a characteristic endothermic peak resulting from melting at 61.6 °C, with a residual mass of 93.00% and weight loss of 3.82%. The thermogram of cholesterol indicates an endothermic peak due to melting at 121.6 °C and 150.3 °C, with a residual mass of 98.25% and weight loss of 2.75%. The DSC of the physical mixture of MTX with niosome components reveals the peaks of both Span 60 and Soya PC, as well as the MTX peak, indicating that it maintains its crystalline form. However, the decrease in MTX peak intensity may be related to the dilution effect caused by the excipients. The thermogram of the unloaded niosomes demonstrates the absence of the MTX characteristic peak. The endothermic peak of Span 60 is present in both the blank niosomes and drug-loaded niosomes but shifted to a lower temperature (61.6 °C). For the drug-loaded niosomes, the MTX characteristic peak is completely gone, which could indicate that MTX was either encapsulated inside the niosomes or transformed into a molecular state within the surfactant mixture. As reported by Nasr et al. (2013) [17], the absence of the drug's crystalline melting peak after encapsulation may also indicate the presence of a strong interaction between the surfactant bilayers of the vesicles and the entrapped drug.

In vitro drug release kinetics

Figure 7 depicts the in vitro release profile of the optimized niosomal suspension loaded with methotrexate at pH 7.4 and 35 ± 0.5 °C. The optimized formulation exhibited a cumulative drug release of 75.09% over 24 h. The release curve indicated an initial burst release where approximately 28% of the drug was released within the first two and a half hours. The subsequent drug release from the optimized niosomes was sustained. The initial burst release was attributed to the presence of free methotrexate in the external phase.

In vitro cell proliferation studies

The anti-cancer efficacy of the formulations Plain MTX, NIO 5, NIO 12, and NIO 16 was tested against the murine skin melanoma cell line B16-F10 using the SRB assay, with ADR (Adriamycin) serving as a positive control. Figure 8A, B shows the activity of the formulations against B16-F10 cells. The results, presented in Table 2, indicate that NIO 16 was significantly more effective than other formulations like NIO 5 and Nio12. The GI₅₀ values were 8.05738 μg/mL (NIO 5), 2.5311 μg/mL (NIO 12), 38.7176 μg/mL (NIO 16), and 40.2238 μg/mL (Plain MTX). The investigation revealed that Nio16 (with a GI50 of 38.7176 μg/mL) exhibited considerable potential in countering cancer activity within B16-F10 cells. It displayed enhanced efficacy in curbing cell growth when contrasted with alternative formulations. This heightened effectiveness might be attributed to potential factors such as the diffusion of niosomes from the stratum corneum, the amplification of permeation through the influence of Span 60 surfactants present in the niosomal formulation, or even modifications to the stratum corneum to augment the permeability of the intercellular lipid barrier. The drugs (MTX) can cross through the niosomes due to their interactions with aggregation, fusion, and adhesion, leading to a large thermodynamic gradient. Our research found that Nio16 had a potent cell inhibition capacity against B16-F10 cells, suggesting that the modified formulation would be beneficial in the treatment of cancer.

Table 2 Cytotoxic effects of formulations NIO 14 to NIO 17 on murine tumor cell line B16F10 cells

Ex vivo and In vivo investigations

Ex vivo skin penetration study

The ex vivo drug permeation study was conducted to gain insights into the potential in vivo performance of RH-loaded niosomes as a transdermal drug delivery system. Employing CLSM, z-stack images were captured to depict optical sections of mice skin at 2 µm intervals, covering a depth span of 0 to 38 µm. Formulations utilizing PBS as a positive control did not exhibit any detectable particles. Likewise, formulations with Plain Rhodamine as a negative control also displayed no observable particles. The findings from this ex vivo skin permeation study, which identified specific depth ranges for particle presence in various niosome formulations, have significant implications for potential applications in skin cancer treatment. Notably, Nio13 (982.3 nm), with particles observed at a depth of 0–6 µm, suggests a potential for superficial skin cancers. Nio14 (783.9 nm), spanning from 4 to 18 µm, may have relevance in treating lesions at slightly greater depths. The formulation Nio15 (707.7 nm), demonstrating distinct particles between 16 snf 22 µm, could be well-suited for targeting tumors residing slightly deeper within the skin. For more deeply seated skin cancers, Nio16 (671.5 nm), with particles at depths ranging from 24 to 32 µm, may offer a solution. Additionally, Nio17 (597.2 nm), with particles distributed within a broad range of 12 to 38 µm, could be instrumental in addressing skin cancers with varying depths or infiltrative characteristics. The particles of Nio 18 (205.0 nm) will go into systemic circulation. This study suggests that tailoring niosome size and properties to specific depth ranges has the potential to enhance localized drug delivery for skin cancer treatment, increasing the precision and effectiveness of therapeutic interventions. These findings offer valuable insights into the distribution patterns and potential penetration capabilities of the RH-loaded niosomes across different skin layers. CLSM has emerged as a valuable tool for studying the skin's permeation and nanocarrier distribution due to its real-time imaging, multi-depth capabilities, and noninvasive nature. In this study, Rhodamine B (RH) acted as a model lipid-soluble fluorophore, mimicking a hydrophilic drug in niosomes. Before imaging, untreated skin displayed no fluorescence within the fluorophore's detection range. The visualizations (Fig. 9) showcased the widespread distribution of fluorescence across skin layers following treatment with fluorolabeled niosomes. The interaction of proposed surfactant-containing niosomes with skin surface lipids might have facilitated intracellular drug penetration. This study's findings highlight the potential of MTX-loaded niosomes as effective transdermal drug carriers and underscore the utility of CLSM in comprehending their skin permeation behavior.

In vivo histopathological study

The in vivo histopathological examination provided valuable insights into the potential skin irritation of different treatment groups. The untreated control group (Group I) displayed a healthy and intact skin structure with well-defined layers, as seen in Fig. 10. This serves as a baseline for comparison. Skin treated with only the MTX solution (Group II) also showed an intact epidermis and dermis, despite minor congestion in deeper blood vessels. This suggests that the MTX itself might not be a significant skin irritant in this context. Skin sections from groups receiving niosomal formulations (Groups III–VII) revealed a key finding: no epidermal erosion. However, crucially, this erosion was not accompanied by any signs of inflammation, such as edema or erythema. This strongly suggests that the irritation was mild and manageable, likely influenced by the presence of penetration enhancers like Span 60 and Soya PC in the niosome formulations. Importantly, other skin layers remained intact, and no major disruptions were observed beyond the topmost keratinized stratum corneum. While the qualitative observations are valuable, adding a quantitative scoring system to assess the degree of epidermal erosion and congestion across groups could further strengthen the conclusions about niosomal safety. This would provide more objective and comparable data for future studies. These findings support the idea that MTX-loaded niosomes exhibit a relatively acceptable safety profile. The absence of significant inflammation despite mild erosion suggests that the potential for irritation in clinical settings is likely low. However, future studies implementing quantitative scoring would further bolster these conclusions and provide even more robust evidence for the safety of these niosomes as drug delivery systems.

Discussion

The discussion of the study's findings reveals important insights into the formulation and properties of MTX-loaded niosomes, as well as their potential for cancer treatment.

The study's investigation into the impact of formulation components, particularly the ratios of span 60, soya PC, and cholesterol, on niosome properties revealed a complex relationship between these components and vesicle size. The initial decrease in vesicle size with a higher span 60 concentration highlights the surfactant's interfacial activity, facilitating smaller vesicle formation. However, beyond a certain point, increased span 60 content coupled with higher cholesterol led to larger vesicle sizes due to increased rigidity. This observation underscores the delicate balance required between these components to achieve optimal vesicle properties. A possible solution lies in fine-tuning the component ratios to strike the right balance between interfacial activity and vesicle stability.

The study's findings on the encapsulation efficiency of MTX shed light on the influence of sonication time on drug loading. The variations in encapsulation efficiency can be attributed to the interplay of sonication energy, vesicle size, and drug solubility. To address this, a controlled and standardized sonication process should be established, accounting for both drug entrapment and vesicle size optimization.

The FTIR spectroscopy results confirm the compatibility of MTX with niosome components, supporting the formulation's potential for effective drug delivery. This is a significant assurance of the stability and efficacy of the loaded drug within the vesicles.

The DSC thermograms' insights into the thermal behavior of components, including the disappearance of the drug's crystalline peak after encapsulation, suggest a change in the drug state or its encapsulation within the vesicles. While this finding holds promise for sustained drug release, further investigations are warranted to understand the exact mechanism of drug encapsulation and release from the vesicles.

The study's anti-cancer efficacy evaluation revealed the superior activity of the Nio17 formulation against B16-F10 cells, attributed to the presence of span 60 as a permeation enhancer. This underscores the potential of niosomes as effective carriers for anti-cancer agents, particularly when incorporating permeation enhancers.

The ex vivo drug permeation study provided visual evidence of the penetration of RH-loaded niosomes into different skin layers, showcasing their potential for transdermal drug delivery. This underscores the practical relevance of the formulation's structure and composition in facilitating drug penetration through the skin. The presence of particles was observed in Nio13 (without sonication), with an average size of 982.3 nm (nm) visible within 0 to 6 µm (μm). This suggests that sonication may be beneficial for enhancing skin penetration. The formulations with sonication durations of 2, 4, 6, and 8 min (Nio14, Nio15, Nio16, and Nio17, respectively) produced progressively smaller particles at increasing depths. The particle sizes ranged from 783.9 nm (4–18 μm), 707.7 nm (16–22 μm), 671.5 nm (24–32 μm), to 597.2 nm (12–38 μm), indicating improved permeation. The positive results of Nio17, revealing the presence of particles within skin layers rather than in the bloodstream, underscore its potential as a promising candidate for effective drug delivery.

The in vivo histopathological examination indicated no epidermal erosion or inflammation, supporting the formulation's potential for transdermal delivery in skin cancer therapy. However, the absence of severe inflammation suggests the formulation's acceptability for clinical use. This observation highlights the importance of considering local skin effects in the development of transdermal drug delivery systems.

This study offers valuable insights into the formulation and properties of MTX-loaded niosomes, demonstrating their potential as effective transdermal drug carriers for cancer treatment. The findings underscore the need for careful optimization of formulation components and process parameters to achieve desired vesicle characteristics, encapsulation efficiency, and drug release profiles.

Conclusion

This study comprehensively investigated the formulation, properties, and potential of MTX-loaded niosomes for transdermal cancer therapy. The complex interplay of formulation components, particularly span 60, soya PC, and cholesterol, on vesicle size and encapsulation efficiency necessitates fine-tuning to achieve optimal properties. Compatibility between MTX and niosomes was confirmed by FTIR, while DSC hinted at a possible sustained release mechanism. Notably, Nio16 demonstrated superior anti-cancer activity against B16-F10 cells, highlighting the benefits of permeation enhancers. Ex vivo studies visually confirmed niosome penetration into skin layers, with smaller particles achieving deeper penetration. While mild epidermal erosion was observed in vivo, the absence of severe inflammation suggests acceptable safety. This study establishes the promising potential of MTX-loaded niosomes for transdermal cancer treatment, emphasizing the importance of optimizing formulation parameters and exploring complementary techniques like CLSM for further validation. This research lays the groundwork for developing innovative and effective transdermal drug delivery systems for cancer therapy.

In the future, our study will be advanced by comparing niosomes with market-available formulations for skin diseases and other nanovesicles under research. Physicochemical properties of our niosomes will be compared with liposomes, polymeric nanoparticles, and other nanocarriers using standardized in vitro assays, focusing on size, stability, and drug loading capacity. In vitro drug release profiles will be analyzed to understand potential differences in release kinetics and therapeutic effects. Additionally, comparative in vivo studies in relevant animal models will evaluate biodistribution, pharmacokinetics, and efficacy of our niosomes compared to other nanocarriers, considering factors like targeted delivery, systemic exposure, and potential side effects.