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Article

Development and Physicochemical Characterisation of Probiotic Emulsions Containing Lactobacillus rhamnosus for Potential Dermal Applications

by
Monika Gasztych
1,
Ruth Dudek-Wicher
2,
Dawid Brzozowski
1,
Arleta Dołowacka-Jóźwiak
3 and
Witold Musiał
1,*
1
Department of Physical Chemistry, Pharmaceutical Faculty, Wroclaw Medical University, Borowska 211, 50-556 Wroclaw, Poland
2
Department of Pharmaceutical Microbiology and Parasitology, Pharmaceutical Faculty, Wroclaw Medical University, Borowska 211, 50-556 Wroclaw, Poland
3
Department of Drug Technology, Pharmaceutical Faculty, Wroclaw Medical University, Borowska 211, 50-556 Wroclaw, Poland
*
Author to whom correspondence should be addressed.
Pharmaceutics 2026, 18(2), 199; https://doi.org/10.3390/pharmaceutics18020199
Submission received: 15 December 2025 / Revised: 21 January 2026 / Accepted: 29 January 2026 / Published: 3 February 2026
(This article belongs to the Section Physical Pharmacy and Formulation)
Browse Figures
Versions Notes

Abstract

Background/Objectives:This study evaluated how variations in emulsion composition influence the viability of a probiotic strain Lactobacillus rhamnosus GG within biphasic systems, as well as the overall stability of the resulting formulations. Methods:Eight biphasic emulsions were prepared, each in two versions—with and without a preservative—and subsequently analysed for pH, FTIR spectroscopy, and emulsion type. The viability of L. rhamnosus GG in each formulation was determined using the plate count method. This method is regarded as the reference technique for the quantitative determination of viable bacteria, expressed as colony-forming units (CFUs). Results: The pH indicated that an emulsion with a pH of 4.65 provides the most favorable conditions for L. rhamnosus GG survival, as values below pH 6 promote its proliferation. This acidity aligns with the natural pH of human skin, although it falls slightly below the recommended 4–5 range for topical formulations. FTIR analysis confirmed the structural stability of the emulsions and revealed spectral shifts attributable to the presence of the bacteria. The spectra remained largely consistent throughout the study period, demonstrating good temporal stability. Conclusions: Microbiological evaluation showed that all produced formulations supported bacterial growth, the presence of the preservative did not inhibit viability of L. rhamnosus GG, corroborating findings from an independent assessment. All emulsions were classified as O/W systems, due to the high water content which is advantageous for microbial viability. Furthermore, O/W emulsions are user-friendly, easy to remove, limit the penetration of the active component into deeper skin layers, supporting their suitability for probiotic-based topical applications.

1. Introduction

The human skin microbiome is recognised as a vital part of cutaneous physiology. Commensal microorganisms coexist with host tissues and actively contribute to skin health by occupying ecological niches, competing with pathogens, and modulating immune responses [1,2]. For example, although they are often considered opportunistic pathogens, Staphylococcus epidermidis and Cutibacterium acnes play a vital role in maintaining microbial equilibrium by producing antimicrobial peptides and modulating the local pH [3].
Disruption to microbial balance, known as dysbiosis, can lead to or exacerbate various dermatological conditions, including atopic dermatitis (AD), psoriasis, acne vulgaris, seborrheic dermatitis, and eczema [4]. Dysbiosis can weaken the skin’s defense barrier, promote the colonisation of pathogenic bacteria such as Staphylococcus aureus and trigger chronic inflammatory responses. Recent evidence also suggests that certain cosmetic ingredients, preservatives, and environmental pollutants can alter the skin microbiota, thereby contributing to barrier dysfunction and inflammation [5,6,7].
Understanding the skin as a symbiotic ecosystem has therefore shifted therapeutic approaches towards restoring microbial balance rather than simply eliminating bacteria. Strategies such as prebiotic, probiotic, and postbiotic treatments have gained significant attention for their ability to modulate the skin microbiome and restore homeostasis [8,9,10,11,12,13].
According to the World Health Organization, probiotics are ‘live microorganisms which, when administered in adequate amounts, confer a health benefit on the host’. Probiotics have traditionally been used to prevent and treat gastrointestinal disorders, but recent research has demonstrated their beneficial effects in extra-intestinal sites, including the skin [14,15].
Probiotics can act directly on the skin’s surface or indirectly via systemic immunomodulatory effects following oral intake. The mechanisms underlying their action include: competitive exclusion of pathogens by occupying adhesion sites and competing for nutrients; production of antimicrobial substances, such as bacteriocins and organic acids; reinforcement of the skin barrier through modulation of lipid metabolism and ceramide synthesis; Regulation of the immune system, including stimulation of anti-inflammatory cytokines (e.g., IL-10) and suppression of pro-inflammatory mediators such as TNF-α and IL-6; enhancement of wound healing by promoting fibroblast proliferation and collagen synthesis [12].
Probiotics have demonstrated clinical potential in managing dermatological disorders. In atopic dermatitis, for example, several studies have reported a reduction in symptom severity and relapse rate following the oral or topical application of Lactobacillus spp. strains [13,16,17]. Similarly, probiotics can inhibit the growth of Cutibacterium acnes in acne vulgaris and restore microbial diversity altered by antibiotic use. Probiotic strains, particularly those belonging to the genus Lactobacillus, have been shown to suppress the growth of Cutibacterium acnes. This is achieved primarily through the production of organic acids and bacteriocins, as well as through competitive interactions on the skin surface. These effects reduce inflammation and restore microbial balance, suggesting that probiotics could be used to manage acne vulgaris [18]. In psoriasis, probiotic supplementation may modulate systemic inflammation by influencing the gut–skin axis [15,19,20].
Of the many probiotic genera, Lactobacillus is one of the most extensively studied. Members of this genus are Gram-positive, non-sporulating, facultatively anaerobic rods that produce lactic acid primarily through carbohydrate fermentation. Common species include L. acidophilus, L. casei, L. brevis, L. bulgaricus and L. rhamnosus [19].
Lactobacillus rhamnosus, in particular, has attracted attention due to its dual functionality, which benefits both intestinal and cutaneous health. This species can survive in mildly acidic environments (pH 4–6), which are typical of the skin surface, and tolerate limited oxygen conditions. It produces antimicrobial metabolites that inhibit pathogens, promote antioxidant defence and support the renewal of the epidermal barrier [15]. Furthermore, L. rhamnosus may interfere with melanogenesis pathways, which could contribute to an even skin tone [20].
Topical applications of Lactobacillus spp. have been reported to reduce skin inflammation and erythema, enhance hydration and strengthen the lipid matrix of the stratum corneum. These effects are not only mediated by live bacteria, but also by bacterial components such as cell wall elements, including peptidoglycans and metabolites, which can act as postbiotics [21].
As well as maintaining bacterial viability, emulsions must meet pharmaceutical standards of homogeneity, physicochemical stability and microbiological safety. These parameters are usually evaluated using methods such as Fourier-transform infrared spectroscopy (FTIR) for structural characterisation, pH measurement, microbial assays and in vitro release studies to evaluate the performance of the formulation [22].
The beneficial effects of probiotics on the skin extend beyond direct antimicrobial action. Recent research highlights their ability to modulate the skin’s innate and adaptive immune responses. Probiotic-derived molecules can interact with Toll-like receptors (TLRs) on keratinocytes and Langerhans cells, activating signaling pathways that influence cytokine production and barrier regeneration [23].
For example, stimulation of TLR2 by Lactobacillus components enhances tight-junction protein expression, improving the cohesion of the stratum corneum. Furthermore, probiotics can modulate the balance between pro-inflammatory Th1/Th17 and anti-inflammatory Th2/Treg responses, thereby reducing chronic inflammation associated with conditions like atopic dermatitis or rosacea [24].
By influencing both local and systemic immunity, probiotics may also play a role in the so-called gut–skin axis, a bidirectional communication system linking intestinal microbiota and skin health. Oral administration of Lactobacillus rhamnosus has been shown to improve skin barrier function, reduce transepidermal water loss, and decrease the incidence of eczema in children [25]. These findings suggest that maintaining microbial balance across multiple body sites may have synergistic effects on skin homeostasis.
Despite the promising results, developing probiotic-based topical products presents several technological constraints. Probiotic bacteria are sensitive to adverse conditions, including temperature fluctuations, exposure to oxygen, UV radiation and the presence of preservatives or surfactants commonly used in pharmaceutical formulations [26]. Ensuring their viability and biological activity throughout production, storage and application remains a significant challenge.
An essential requirement for effective topical delivery systems is development of a stable microenvironment that supports bacterial survival while preserving the desired physicochemical properties of the formulation. Among the systems available, emulsions—mixtures of immiscible oil and water phases stabilised by surfactants—have gained prominence as promising carriers for probiotics [27]. Their biphasic structure enables the bacteria to be evenly distributed within the aqueous phase while being physically protected from the surrounding lipid layer. Furthermore, emulsions can be engineered to optimise texture, spreadability and the controlled release of active ingredients.
Selecting appropriate excipients is essential to ensure compatibility between probiotic cells and formulation components. Non-ionic emulsifiers, humectants and mild preservatives are often preferred to minimise bacterial stress. In addition the pH of the emulsion should match that of the skin (typically 4.5–5.5) to maintain product stability and skin tolerance [28].
In light of the increasing scientific and clinical interest in probiotic-based dermatological products, there is an urgent need for systematic research into the formulation and characterisation of stable probiotic emulsions that are suitable for topical application. Although the beneficial properties of Lactobacillus rhamnosus are well documented, concerns about stability and compatibility limit its use in cosmetic or pharmaceutical emulsions.
It should be noted that the present study constitutes a preliminary evaluation of probiotic-containing topical emulsions, focusing on physicochemical stability, microbiological safety, and bacterial viability. Despite its exploratory nature, this work provides scientifically relevant insights into the formulation-related factors influencing the survival of L. rhamnosus GG in semi-solid topical systems, thereby establishing a foundation for further formulation optimisation and advanced biological investigations.
The present study therefore aimed to develop and evaluate emulsions containing L. rhamnosus GG as a model probiotic strain. The formulations were characterised in terms of their physicochemical parameters (pH, emulsion type), microbiological properties (contamination control, bacterial viability) and structural features, which were analysed using FTIR. Additionally, the impact of preservatives and other excipients on probiotic survival was examined to optimise formulation safety and performance.
The ultimate goal of this research was to create a stable, microbiologically safe emulsion that can serve as a carrier for live probiotic bacteria. This would contribute to the development of innovative topical preparations that support skin health and homeostasis.

2. Materials and Methods

2.1. Reagents

Cetyl alcohol (Sigma Aldrich, Poznan, Poland), beta glucan (Zrob sobie krem, Prochowice, Poland), biobased (Ecospa s.c., Warszawa, Poland), behentrimonium methosulfate (Ecospa s.c., Warszawa, Poland), olivem 1000–cetearyl olivate sorbitan olivate (Ecospa s.c., Warszawa, Poland), lanolin (Fagron, Warsaw, Poland), benzyl alcohol (Ecospa s.c., Warszawa, Poland), dimethicone 350 (Ecospa s.c., Warszawa, Poland), glycerine (Pol-Aura Sp. z o.o., Morąg, Poland), propylene glycol (Pol-Aura Sp. z o.o., Morąg, Poland), glyceryl stearate citrate (Pol-Aura Sp. z o.o., Morąg, Poland), inulin (Info-Farm, Katowice, Poland), stearic acid (Sigma Aldrich, Poznan, Poland), Lactobacillus rhamnosus GG (ATCC 53103, Manassas, VA, USA) (Pharmabest, Warszawa, Poland), glyceryl monostearate (Ecospa s.c., Warszawa, Poland), cetearyl olivate (Ecospa s.c., Warszawa, Poland), sorbitan olivate (Ecospa s.c., Warszawa, Poland), linseed oil (Chmarablend, Suwałki, Poland), sweet almond oil (Chmarablend, Suwałki, Poland), MRS broth (de Man, Rogosa and Sharp, Biomaxima, Lublin, Poland), MRS agar (de Man, Rogosa and Sharpe agar, Biomaxima, Lublin, Poland), carnauba wax (Ecospa s.c., Warszawa, Poland), white beeswax (Ecospa s.c., Warszawa, Poland), DHA BA preservative–benzyl alcohol, dehydroacetic acid (Ecospa s.c., Warszawa, Poland), deionised water.

2.2. Preparation of Emulsions Containing Lactobacillus rhamnosus GG

The emulsions were prepared using the hot emulsification method, which is widely used in the development of thermally processed oil-in-water systems. The components of the oil phase were weighed accurately and transferred to a dedicated laboratory reaction vessel. Meanwhile, the constituents of the water phase were measured and placed into a separate reaction vessel. The oil and aqueous phases were heat-treated separately in a thermostatically controlled water bath at 70 °C for 10 min prior to emulsification, to ensure the lipid phase had completely melted and both phases were thermally equilibrated. During heating, vigorous agitation of the oil phase was employed to facilitate the uniform melting and dispersion of its constituents. The oil and water phases were mixed continuously at 300 rpm for 10 min using a magnetic stirrer (Multi-HS Digital, Velp, Monza and Brianza, Italy) while being heated to ensure complete melting and homogeneity. Once the target temperature range for emulsification was reached, the aqueous phase was gradually introduced into the oil phase while stirring continuously. The resulting mixture was then removed from the water bath and homogenised at a specific speed using a homogeniser until it cooled to ambient temperature to ensure the formation of a stable primary emulsion. Homogenization was performed using a CAT Unidrive X 1000D homogenizer (CAT Scientific, Ballrechten-Dottingen, Germany) at 10,000 rpm for 3 min to obtain a homogeneous emulsion. After cooling, L. rhamnosus GG was added to the formulation as a probiotic additive. The prepared emulsion was then divided into two portions, one of which was supplemented with a natural preservative DHA BA to enhance microbiological stability. Both formulations were transferred to sterilised, tightly sealed containers and stored under refrigerated conditions appropriate for systems containing probiotics. Samples enriched with the preservative were marked for identification purposes. The qualitative compositions of the developed emulsions are presented in Table 1.

2.3. Measurement of pH Value

The pH of the formulated preparations was determined using a CPC-505 pH meter (accuracy ±0.002 pH; Elmetron Sp.j., Zabrze, Poland) coupled with an ERH-11S electrode (Elmetron Sp.j., Zabrze, Poland) suitable for high-viscosity semisolid systems, including ointments, creams, and gels. Each sample was subjected to five independent measurements to ensure analytical reliability.

2.4. Visual Assessment of the Formulation

A preliminary qualitative visual inspection was used to monitor macroscopic changes indicative of physical instability, such as phase separation or visible contamination. The appearance of each preparation was assessed at seven-day intervals under consistent lighting and environmental conditions. This included examining colour, homogeneity, phase separation, sedimentation, changes in opacity and the presence of any visible particulate matter or microbial growth. Any deviations from the initial appearance were documented. This qualitative analysis provided an early indication of potential instability, complementing the physicochemical and microbiological tests performed on the emulsions. Observations were discontinued on day 42.

2.5. Determination of Emulsion Type

The type of the obtained emulsions was assessed using three complementary analytical approaches: the electrical conductivity method, the dye test, and the drop dilution test. Each emulsion was examined using all three methods.

2.5.1. Electrical Conductivity Method

The electrical conductivity of the formulations was measured in analogue multimeter VC-2020 (Voltcraft, Kraków, Poland) with metallic electrodes to differentiate between oil-in-water (O/W) and water-in-oil (W/O) systems. Emulsions exhibiting measurable electrical conductivity were classified as O/W, indicating the presence of a continuous aqueous phase. Conversely, formulations that did not conduct electric current were identified as W/O emulsions, reflecting a continuous oil phase.

2.5.2. The Dye-Solubility Test

The dye test was performed to determine the type of emulsion, based on the physicochemical properties of two dyes: Sudan III and methyl orange. Sudan III is an intensely red, lipophilic dye that is readily soluble in non-polar media such as oils, whereas methyl orange is an orange-coloured dye that is soluble in polar compounds such as water. This approach enables the continuous phase in the investigated emulsions to be identified. For the analysis, two aliquots of each emulsion sample were collected. Sudan III was added to one and methyl orange to the other, followed by gentle mixing. The stained samples were then examined under a stereoscopic microscope (FPN, Nysa, Poland). Selective staining of the continuous phase was used to classify the emulsion type: staining of the continuous oily phase indicated water-in-oil (W/O) emulsions, while staining confined to the dispersed droplets indicated oil-in-water (O/W) emulsions.

2.5.3. Drop Dilution Test

The drop dilution method involved depositing a droplet of 50 µL of deionized water onto the surface of the emulsion. The miscibility behavior of the droplet served as the basis for classification: lack of spreading or coalescence of the water droplet indicated a W/O system, whereas immediate dispersion and merging of the droplet with the sample were indicative of an O/W emulsion.

2.6. Assessment of the Antibacterial Activity of the Preservative

To determine the effect of the preservative concentration on the viability of the microorganisms present in the formulation, the preservative’s antibacterial activity was evaluated using the standard serial microdilution method. A suspension of the reference strain L. rhamnosus GG was prepared from a 24 h culture and adjusted to a density of 1 McFarland unit. This was then diluted 1000-fold to obtain a final concentration of approximately 105 CFU/mL. 100 µL of Mann–Rogosa–Sharpe (MRS) medium (Biomaxima, Poland) was dispensed into the wells of a 96-well microtiter plate. Geometric serial dilutions of the tested preservative were then prepared, with the highest evaluated concentration being 0.2%. Next, 100 µL of the microbial suspension (105 CFU/mL) was added to each well containing the preservative dilutions. Parallel controls were included: a positive growth control (K+), consisting of the microbial suspension in MRS medium without the preservative; and a negative sterility control (K−), consisting of MRS medium alone. The microtiter plate was then incubated for 24 h at 37 °C under 5% CO2, with continuous shaking at 400 rpm (IKA Schüttler MTS 4). After incubation, 20 µL of a 0.1% 2,3,5-triphenyltetrazolium chloride (TTC) solution in MRS medium was added to each well and the plate was incubated again at 37 °C under 5% CO2 for 4 h. The formation of a red colour indicated the reduction of TTC to formazan, which signifies the presence of metabolically active bacterial cells. The test was performed in triplicate.

2.7. Microbiological Evaluation of the Emulsions

A microbiological analysis was performed to confirm the presence of viable L. rhamnosus GG cells in the prepared emulsions. Microbiological analysis of probiotic viability was performed on day 30 after preparation of the emulsions. Each sample underwent sterility testing in accordance with the procedures outlined in the 9th and 10th editions of the European Pharmacopoeia (Chapters 2.6.1 and 2.6.27, respectively) [29]. Approximately 500 mg of each emulsion was aseptically transferred into a sterile glass vessel. 10 mL of sterile MRS (de Man, Rogosa and Sharpe) medium (Biomaxima, Lublin, Poland) was added and the sample was mixed for 30 min using a magnetic stirrer until the formulation was completely dispersed. Subsequently, 10 mL of the resulting mixture was transferred to 90 mL of liquid MRS medium. The suspension was then incubated at 37 °C for 48 h under anaerobic conditions to allow for the recovery and growth of microbes. All procedures were carried out under aseptic conditions. Plate Count method was then performed. A dilution series was prepared in sterile physiological saline ranging from 100 to 10−10. From each dilution, 100 µL was aseptically plated onto MRS agar using the surface spread technique with a sterile spreader. All samples were plated in triplicate. In parallel, the sterility of the physiological saline used for dilutions was tested by inoculating 100 µL of the saline onto MRS agar. A total of four control plates were prepared. All plates were incubated for 48 h at 37 °C in an atmosphere containing 5% CO2. All results of performed count method is presented in the Table S1 of Supplementary Materials. To calculate CFU/mL, agar plates with two successive dilutions were selected based on quantitative range between 15 and 300 CFU per plate. Following incubation, colony-forming units (CFU) were counted. The microbial count per milliliter of the prepared suspensions was calculated using the following formula:
L = d ⋅ a ⋅ C N 1 + 0.1 N 2
where: L—number of CFU in 1 g or 1 mL, d—dilution factor, a—inoculation factor, C—total number of colonies on all plates selected for counting, N1—number of replicates from the first selected dilution, N2—number of replicates from the second selected dilution. This procedure was repeated for each emulsion tested.

2.8. Fourier Transform Infrared Spectroscopy (FTIR)

FTIR analyses were performed using a Nicolet iS50 spectrophotometer equipped with an ATR module (Thermo Scientific, Waltham, MA, USA). Spectra were collected and processed with OMNIC software (version 9, Thermo Fisher Scientific). Each measurement consisted of 32 scans per sample, recorded at a spectral resolution of 4 cm−1 across the range of 4000–400 cm−1. All ATR–FTIR measurements were conducted at ambient temperature. The primary aim of the analysis was to determine whether the infrared spectra of the emulsions changed over time, which would indicate instability, or remained constant, confirming their physicochemical stability. FTIR was also used to detect characteristic signals of L. rhamnosus GG, enabling verification of the presence and persistence of bacterial components within the formulations. Measurements were performed under identical conditions on days 1, 7, 14, 21, 28, 35, and 42 after preparing the emulsions. Between measurement intervals, the samples were stored at 8 °C in opaque containers to minimise light exposure and slow potential degradation processes.

3. Results

3.1. pH Values of Formulation

The pH values of the formulations under investigation (Em 1–Em 8) ranged from 3.65 to 6.31, indicating noticeable variability between compositions (Table 2). The highest pH value was observed for Em 2 (6.31 ± 0.03), which is close to the upper limit of what is acceptable for dermal applications. By contrast, Em 6 (3.65 ± 0.02) and Em 7 (3.87 ± 0.03) had the lowest pH values, indicating a significantly more acidic nature than the other formulations. Most samples (Em 1, Em 3, Em 4, Em 5 and Em 8) exhibited pH values between 4.4 and 5.9, which are generally compatible with the physiological pH of the skin’s acid mantle. These results suggest that most formulations are suitable for topical application without increasing the risk of disrupting the skin barrier [30,31]. The standard deviations (SD) for all formulations were low (0.01–0.03), confirming the reproducibility of the measurements and the stability of the tested systems. The narrow SD values suggest that the pH of each formulation is well controlled and not prone to fluctuations caused by measurement error or component instability. pH measurements were taken one day after preparation and on the 30th day after formulation. The pH value of the individual formulations was not affected by the addition of the DHA BA preservative.

3.2. Visual Examination of Physical Stability

Visual assessment showed that most of the formulations remained stable throughout the observation period. No changes in colour, homogeneity or phase behaviour were observed in any of the emulsions, except for two preparations. On day 42, visible contamination was observed in emulsions Em 2 and Em 8, whereas all remaining formulations remained visually stable, as shown in Figure 1. The contaminant was identified as mould, which had not been present at any earlier time point. All other emulsions remained visually unchanged for the entire monitoring period.

3.3. Emulsion Type Determination

3.3.1. Conductometric Analysis

Measurements revealed a measurable current flow in all formulations tested, indicating the presence of a continuous aqueous phase. This behaviour is characteristic of oil-in-water (O/W) emulsions, in which the external phase capable of conducting electric current is water.

3.3.2. Dye-Solubility Test

When methyl orange was added, all samples exhibited uniform colouration, confirming that water was the continuous phase. In contrast, Sudan III did not stain the continuous phase, indicating that the oil was present as the dispersed phase. Collectively, these findings support the classification of the formulations as oil-in-water (O/W) emulsions.

3.3.3. Drop Dilution Method

The droplets of the examined emulsions mixed readily with water, but did not disperse in oil. This behaviour confirms that water is the continuous phase, thus verifying that the formulations correspond to the O/W emulsion type. Table 3 shows the type of obtained emulsions.

3.4. The Preservative’s Antibacterial Activity Evaluation

The preservative DHA BA used in the developed formulation did not exhibit antimicrobial activity and against the tested strain. The red coloration of the plate wells presented on Figure 2 indicates the presence of metabolically active microorganisms.

3.5. Microbiological Analysis Outcomes

The emulsions Em 1 and Em 1k showed the lowest counts of viable cells. In contrast, the Em 5k and Em 6 formulations exhibited the highest number of living bacteria. Emulsions Em 1k, Em 2k and Em 4k had a significantly lower number of viable bacteria than their preservative-free counterparts. Em 3k also showed a slight reduction in bacterial counts relative to Em 3. By contrast, Em 5k demonstrated a significant increase in probiotic viability compared with Em 5, while Em 7k and Em 8k showed only minor increases in bacterial numbers. After incubation of Em 6k samples no bacterial growth was observed in all dilution of sample. All results of cell count is presented in the Table S1 in Supplementary Materials. The resulting data obtained after calculation described in Section 2 are presented in Table 4.

3.6. Fourier Transform Infrared Spectroscopy (FTIR) Analysis of L. rhamnosus GG

To analyse the emulsion using FTIR, it was necessary to characterise the spectrum of the probiotic bacterium used in the formulation of all tested preparations. The microbial FTIR spectrum was therefore divided into five specific segments (Figure 3). Segment 1 contains peaks that reflect the presence of fatty acids in the bacterial cell membrane. Segment 2 includes peaks characteristic of amide bonds found in peptides and proteins. Segment 3 is a non-selective region in which peaks corresponding to both proteins and fatty acids can be identified. Segment 4 shows peaks confirming the presence of polysaccharides in the bacterial cell wall. The final region, segment 5, is referred to in this study as the “true” fingerprint region, as the bonds expressed in this area cannot be assigned to specific functional groups.
Within segment 1, characteristic peaks indicate the presence of fatty acids in the test sample, most likely originating from the bacterial cell membrane. In segment 2, three peaks can be distinguished at 1741.04 cm−1, 1637.02 cm−1, and 1577.38 cm−1. These correspond to stretching vibrations of C=O groups (typical of aldehydes, ketones, acids, or esters), C=N stretching (imines, oximes), C=C stretching (alkenes), and N–H deformation vibrations. Taken together, these signals indicate the presence of amide bonds, confirming protein components in the analysed sample. The subsequent peaks are essential for confirming the presence of L. rhamnosus GG. Segment 3 includes a peak at 1412.57 cm−1, corresponding to O–H deformation vibrations, which suggests the presence of proteins or fatty acids in the preparation. The next three peaks, at 1039.18 cm−1, 988.88 cm−1, and 923.96 cm−1, correspond to weak C–C stretching vibrations and indicate the presence of polysaccharides, most likely derived from the bacterial cell wall. In the final segment, a peak with a wavenumber of 854.51 cm−1 can be identified [32].

3.7. FTIR Analysis of Emulsions Containing Probiotic Strain

Infrared spectra of all eight formulations were compared with corresponding emulsions lacking L. rhamnosus GG, enabling identification of spectral differences associated with the presence of bacterial components. The analysis was conducted within the previously defined five diagnostic spectral segments, allowing systematic interpretation of functional group–specific vibration changes over the 42-day study period. Emulsions Em 7 and Em 8 were not included in the description because the only difference in their composition compared to Emulsions Em 2 and Em 6 was the presence of inulin, which did not affect the FTIR spectrum image.
  • Em 1
Comparison of the spectra revealed that in segment 1, the emulsion containing L. rhamnosus GG exhibited an additional peak at 2955.28 cm−1, as well as more intense peaks at 2916.50 cm−1 and 2849.44 cm−1. These signals indicate an increased number of C–H and O–H vibrations associated with fatty acids originating from the bacterial membrane. In segment 2, the appearance of a peak at 1735.81 cm−1 suggests a higher contribution of amide-related bonds compared with the control emulsion. Segment 3 displayed multiple additional peaks (1377.41, 1265.22, and 1219.45 cm−1), alongside a notably intensified peak at 1465.69 cm−1, indicating the presence of protein- and lipid-associated functional groups. In segment 4, additional peaks at 1195.89 cm−1 and 1098.96 cm−1 were observed, along with an intensified polysaccharide-related signal at 1098.96 cm−1. These features correspond to carbohydrate structures characteristic of the bacterial cell wall. Segment 5 showed a distinct peak at 719.96 cm−1, which falls within the fingerprint region and cannot be assigned to a specific functional group.
  • Em 2
As shown in Figure 4 Em 2, segment 1 exhibited intensified fatty-acid-related peaks at 2916.50 cm−1 and 2849.44 cm−1 in the probiotic emulsion. In segment 2, a stronger amide-related peak at 1740.36 cm−1 was detected. Within segment 3, the peak at 1465.69 cm−1 was more intense compared with the control, again reflecting protein and lipid contributions. In segment 4, additional peaks at 1196.63 cm−1 and 1101.35 cm−1, along with an increased signal at 1077.51 cm−1, correspond to polysaccharide vibrations characteristic of the bacterial cell wall. A fingerprint-region peak at 720.33 cm−1 appeared in segment 5.
  • Em 3
In this formulation (Figure 4 Em 3), differences between the probiotic and control emulsions were less pronounced; therefore, interpretation focused on peaks coinciding with typical bacterial functional groups. In segment 1, peaks at 2915.47 cm−1 and 2848.61 cm−1 correspond to fatty-acid functional groups. Segment 2 included peaks at 1734.78 cm−1 and 1637.44 cm−1, confirming amide-associated vibrations originating from bacterial proteins. Segment 3 contained a range of protein- and lipid-related peaks, with the most prominent at 1466.14 cm−1 and 1378 cm−1. In segment 4, characteristic polysaccharide peaks appeared at 1197.84 cm−1, 1173.89 cm−1, and 1051.24 cm−1, confirming cell-wall carbohydrate components. No significant features were noted in segment 5.
  • Em 4
Based on spectra in Figure 4 Em 4, only subtle differences were detected between the probiotic and control emulsions. In segment 1, peaks at 2915.64 cm−1 and 2848.61 cm−1 again indicated the presence of fatty-acid functional groups. Segment 2 showed peaks at 1735.17 cm−1 and 1640.51 cm−1, consistent with amide vibrations. Segment 3 included prominent protein- and lipid-associated peaks at 1464.58 cm−1, 1377.27 cm−1, and 1219.40 cm−1. In segment 4, polysaccharide-related peaks appeared at 1196.70 cm−1, 1174.79 cm−1, 1112.84 cm−1, and 1051.24 cm−1. No significant peaks were observed in segment 5.
  • Em 5
In Figure 4 Em 5, spectral differences again followed a pattern similar to the bacterial reference spectrum. Segment 1 exhibited peaks at 2916.34 cm−1 and 2848.69 cm−1, associated with fatty acids. In segment 2, peaks at 1736.49 cm−1 and 1640.59 cm−1 reflected amide-related vibrations. Segment 3 showed notable peaks at 1462.75 cm−1, 1376.79 cm−1, and 1242.56 cm−1, indicating proteins and fatty acids. In segment 4, the dominant polysaccharide peaks occurred at 1162.75 cm−1 and 1039.37 cm−1. Segment 5 did not include diagnostically relevant peaks.
  • Em 6
In Figure 4 Em 6, segment 1 displayed increased intensity at 2916.74 cm−1 and 2849.10 cm−1, confirming the presence of bacterial fatty acids. In segment 2, the peak at 1734.86 cm−1 was enhanced, suggesting elevated amide content. Segment 3 contained additional protein- and lipid-related peaks at 1370.50 cm−1, 1316.58 cm−1, and 1246.58 cm−1. No significant peaks were found in segment 4 for this formulation. Segment 5 featured a characteristic fingerprint-region peak at 718.92 cm−1.
All formulations were monitored over a 42-day period, with FTIR measurements taken every seven days. The spectra remained highly consistent throughout the study across all analysed samples. No new absorption bands appeared and none of the characteristic peaks previously identified for L. rhamnosus GG diminished or disappeared. The only changes observed were minor variations in peak intensity, primarily within segments 2 and 4, which reflected subtle fluctuations rather than structural alterations. Overall, the spectral profiles of all emulsions remained comparable at each time point, indicating the formulations’ molecular stability over the entire duration of the analysis.

4. Discussion

This study aimed to determine the influence of the composition of various emulsion systems on the survival of L. rhamnosus GG when incorporated into topical formulations, and to evaluate the stability of these systems during prolonged storage. The formulations remained stable throughout the 42-day observation period, which is an extended assessment of short-term stability relative to standard pharmacy compounding practice. Careful optimisation of physicochemical parameters and microbiological properties is required when developing preparations containing viable probiotics, as these factors directly affect the therapeutic potential of such formulations. The growing interest in topical probiotics for managing skin disorders has highlighted the need for stable, biologically active delivery systems.
The emulsion compositions were selected to produce a range of formulations with different physicochemical characteristics. The prepared emulsions had visibly different consistencies and textures, which reflected differences in the compositions of the formulations and their microstructures. We designed systems that differed in oil-to-water ratio and type of emulsifying agent to create distinct microenvironments within the emulsions. This diversity was essential for evaluating the survival of L. rhamnosus GG under different formulation conditions. This variation enabled us to evaluate the direct impact of the aqueous phase, which is crucial for probiotic viability, as well as the influence of factors such as pH, emulsifier structure, and the presence or absence of preservatives. Differences in texture and consistency also reflect potential real-world applications, as topical formulations must balance stability, sensory qualities, and microbiological safety. Ultimately, the diversity of the tested emulsions enabled a more comprehensive analysis of the formulation parameters that support or hinder probiotic survival. This provided valuable insights for developing stable, probiotic-enhanced topical products.
To address this challenge, eight emulsions were developed, each with different component ratios (Table 1). Two variants were additionally supplemented with inulin to evaluate their potential to promote bacterial growth. Each formulation underwent complete analytical and microbial characterisation, using standard physicochemical and microbiological methods commonly applied in pharmaceutical formulation studies, including pH assessment, determination of emulsion type, FTIR stability analysis, and microbial viability testing.
The pH of the tested emulsions, spanned a broad range, and this parameter exerted a pronounced influence on the behaviour of L. rhamnosus GG within the formulations (Table 2). Previous studies have shown that this species demonstrates enhanced metabolic activity in acidic environments, particularly when the pH drops below 6 [33,34,35]. Consistent with these observations, the present results indicate that formulations with low pH values—such as in case of Em 6 (pH 3.65)—provided a more favourable microenvironment for probiotic survival than those with pH values approaching neutrality (e.g., Em 2 and Em 8). It is interesting to note that the acidic pH of certain formulations closely corresponds to the natural acidity of human skin, which is typically around pH 5 or lower [30]. However, the obtained values were slightly below the range generally recommended for topical products (pH 4–5) [31]. Collectively, these observations indicate that maintaining an appropriately acidic environment is crucial for ensuring both probiotic viability and the microbiological stability of the final product.
Notably, fungal contamination occurred exclusively in Em 2 and Em 8 after 42 days of storage (Figure 1), further suggesting that the physicochemical conditions in these two formulations were suboptimal for maintaining microbial stability. Both emulsions contained biobase in the oil phase as well as glycerine in the aqueous phase, which may have influenced water activity or nutrient availability in a manner conducive to fungal growth [36]. Both emulsions have virtually the same composition, differing only in the presence of inulin, which is found in Em 8. Many studies have shown that inulin, a multifunctional agent in skin care, may enhance product’s stability and increase shelf-life [37,38]. However, the presence of mold may indicate the growth of fungi of the genera Aspergillus spp., Penicillium spp., and Rhizopus spp., which are the most predominant genera responsible for cosmetic contamination [39,40]. Listed fungi display the ability to produce inulinases-enzymes that hydrolyze inulin into sugars and bioethanol [41,42,43,44]. The loss of microbial stability of Em 8 may indicate that the concentration of inulin used should be increased. In addition, the following formulation- and process-level interventions could be considered to prevent fungal proliferation: selection or optimisation of a broad-spectrum preservative system with proven antifungal activity, such as sorbic acid/potassium sorbate or validated combinations like phenoxyethanol with complementary agents. Adjustment of the product pH to the effective range of the chosen preservative. These measures can markedly improve resistance to mould growth. Further practical measures to limit contamination and ingress of spores include reducing water activity through appropriate humectant balance, improving manufacturing hygiene (GMP) and employing protective packaging (e.g., airless dispensers or single-dose formats). Finally, any proposed changes should be validated by standard microbiological challenge tests to confirm the efficacy of the preservatives under realistic conditions [40]. All tested preparations were confirmed to be oil-in-water (O/W) emulsions (Table 3). This type of system is particularly advantageous for the incorporation of viable probiotics, as the aqueous continuous phase ensures an environment suitable for microbial activity. Moreover, O/W emulsions are widely regarded as user-friendly due to their light sensory profile, ease of spreading, and the minimal risk of promoting deep skin penetration of active components [45]. These characteristics enhance the safety and practicality of probiotic-containing topical products. Alternative carrier systems, such as W1/O/W2 multiple emulsions stabilized by internal-phase gelation, have demonstrated potential for improving probiotic survival and may serve as an innovative direction for further development [46]. Microbiological analyses confirmed the sustained survival of L. rhamnosus in all formulations except of Em 6k (Table 4). The highest probiotic viability was observed in formulation Em 6. This suggests that the specific combination of excipients in this formulation created a more protective microenvironment within the aqueous phase. Em 6 had low pH, which probably improved bacterial tolerance by aligning with the natural acidic preference of L. rhamnosus GG. The ratio of emulsifiers and the nature of the oil phase may also have enhanced droplet stability and reduced oxidative or osmotic stress. These are conditions that are known to influence probiotic persistence in semi-solid matrices [46,47,48]. In contrast, formulations Em 2 and Em 8—despite belonging to the same emulsion class as Em 6—supported bacterial survival to a lesser extent, and were the only systems to develop fungal contamination. Both emulsions contained biobase in the oil phase and glycerine in the aqueous phase; these components may have altered water activity or nutrient availability in ways that were less favourable for the probiotic, but more conducive to fungal growth. The near-neutral pH of these formulations may have further reduced the competitiveness of L. rhamnosus GG, as its metabolic activity decreases above pH 6 [49,50,51]. These observations highlight that favourable bacterial survival in O/W systems does not depend solely on the emulsion type, but rather on the interplay of pH, emulsifier composition, oil-phase characteristics and water-activity-modifying excipients. The exceptional performance of Em 6 emphasises the importance of optimising not only the continuous phase, but also the microenvironment surrounding the dispersed droplets, to ensure the long-term stability of probiotics in topical emulsions. The test showed that there are no bacteria in the Em 6k emulsion. Based on the fact that emulsions Em 6 and Em 6k differ only in the presence of the preservative, it might be concluded that the lack of bacterial growth was caused by interactions between glycerin and DHA-BA. It has been shown that humectants such as glycerin can reduce available water and boost preservative action which in case of Em 6k could lead to eradication of probiotics implemented to this formulation.
This unexpected result may reflect either a formulation-dependent effect or a methodological limitation and warrants further investigation in future studies. This conclusion is based on the fact that the bacterial count of a solution made using the Em 6 emulsion, which differs only in the absence of a preservative, is 3.15 × 1010 CFU/mL [39].
Although the implemented preservative, DHA-BA, did not display antimicrobial activity (Figure 2), clear differences were observed between emulsions with and without the preservative (Table 4). These results imply that the impact of DHA-BA on L. rhamnosus GG may be indirect and highly dependent on the formulation matrix. In some emulsions, DHA-BA may transiently increase osmotic or oxidative stress, slightly suppressing bacterial survival. In other systems—particularly those with a favourable emulsifier–oil ratio—the preservative may modify interfacial organisation or reduce droplet coalescence, thereby stabilising the microenvironment surrounding the bacteria. Such structural differences could alter water activity or diffusion processes within the emulsions, indirectly promoting or reducing probiotic viability. Interestingly, in most formulations the presence of the DHA–BA preservative did not increase bacterial counts and, in several cases, was associated with slightly lower viability. The notable exception was that Em 5k exhibited the highest probiotic count of all the tested formulations, despite the presence of the preservative. This suggests that the preservative system, although mild and accepted for use in probiotic-containing formulations, does not actively contribute to maintaining probiotic viability and may impose a low-intensity stress on the cells. The notable exception is Em 5k, in which the probiotic count was significantly higher despite the presence of the preservative, further highlighting that the overall formulation matrix plays a much more decisive role than any single component. Overall, the results suggest that DHA-BA does not have a consistent impact on probiotics, but rather interacts with the unique composition and microstructure of each formulation, leading to variable survival outcomes [52,53,54]. The lowest viable cell counts were observed in the Em 1 and Em 1k emulsions may also be related to the specific composition of these formulations, which appears less supportive of L. rhamnosus GG survival. Notably, both emulsions contain dimethicone—a silicone oil known for its occlusive properties—which may create a physicochemical environment that limits water availability or interferes with the probiotic’s metabolic activity, thereby reducing bacterial viability [55,56,57]. In contrast, Em 5k emulsion incorporates a combination of two emulsifiers and two oils, a system that seemingly provides a more favorable microenvironment for maintaining probiotic stability. The synergistic interactions between these excipients may contribute to improved dispersion, protection against oxidative stress, or enhanced retention of moisture, all of which can support long-term bacterial survival expressed in high bacterial count observed.
Collectively, these findings underscore the importance of excipient selection and their interactions within the emulsion system. The ability of L. rhamnosus GG to remain viable is strongly dependent on the physicochemical characteristics of the formulation, reinforcing the need for careful design of probiotic-containing semi-solid products.
FTIR spectroscopy is a widely used analytical method in pharmaceutical technology thanks to its ability to detect chemical changes and degradation processes over time [58]. Throughout the 42-day study period, the spectral profiles of the emulsions remained largely unchanged (Figure 3 and Figure 4). However, minor fluctuations in peak intensity were observed, particularly in frequency regions associated with protein- and polysaccharide-derived functional groups. Nevertheless, the fundamental spectral features characteristic of L. rhamnosus remained intact. This suggests that neither the excipients nor the storage conditions caused chemical changes that affected the probiotic’s structural integrity. Overall, the FTIR findings confirm the robustness of the emulsions and the persistence of the bacterial molecular signature throughout the study.
This study is a preliminary proof-of-concept investigation into identifying the key formulation parameters influencing the stability and viability of L. rhamnosus GG in topical emulsions. Although the results are limited to in vitro and physicochemical evaluations, they provide scientifically meaningful evidence that supports the feasibility of using emulsion-based systems to carry live probiotics, and justify further, more advanced studies.
This study was performed on a single bacterial strain and under fixed storage conditions, which limits the generalizability of the findings. Future research should explore the behavior of different probiotic species, assess stability under variable environmental conditions, and investigate the interactions between probiotics and skin microbiota in vivo. Additional attention should also be directed toward optimizing rheological and sensory characteristics, which are important for patient compliance. Expanding this work to include advanced encapsulation strategies may further enhance the longevity and bioactivity of topical probiotics.

5. Conclusions

The proposed formulation compositions resulted in biphasic systems classified as oil-in-water (O/W) emulsions. The conducted analyses demonstrated that FTIR spectroscopy is an effective analytical tool for assessing the viability of Lactobacillus rhamnosus GG in semi-solid pharmaceutical preparations of the O/W emulsion type. All the formulations that were tested stayed viable for the probiotic strain during the 42-day storage period, according to the conditions that were applied in the experiment. However, its abundance differed markedly between the emulsions. This indicates that the specific composition and physicochemical characteristics of each system influenced survival. The highest number of viable bacterial cells was observed in the preparation which exhibited a pH below 4. In contrast, the lowest bacterial survival was recorded in the preparation characterized by a pH exceeding 5. These findings highlight the significant influence of pH on the stability and long-term viability of probiotic microorganisms, importance of optimising continuous phase and microenvironment surrounding the dispersed droplets, to ensure the long-term stability of probiotics in topical emulsions. Obtained observations should be taken into account in the further design of formulations containing live probiotic cultures.

Supplementary Materials

The following supporting information can be downloaded at: https://www.mdpi.com/article/10.3390/pharmaceutics18020199/s1, Table S1: Cell count results in microbiological evaluation of tested emulsions (IC-Incalculable, 0-direct sample without dilution).

Author Contributions

Conceptualization, M.G. and W.M.; methodology, M.G., D.B. and R.D.-W.; software, M.G.; validation, M.G., R.D.-W. and A.D.-J.; formal analysis, M.G.; investigation, M.G. and R.D.-W.; resources, M.G.; data curation, M.G.; writing—original draft preparation, M.G.; writing—review and editing, M.G., R.D.-W. and W.M.; visualization, M.G.; supervision, W.M.; project administration, M.G.; funding acquisition, W.M. All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded by National Science Centre, Poland, grant number 2024/08/X/NZ7/00540.

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

The original contributions presented in this study are included in the article and Supplementary Material. Further inquiries can be directed to the corresponding author.

Conflicts of Interest

The authors declare no conflicts of interest.

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Figure 1. Visual appearance of all developed emulsions after 42 days of storage, with visible contamination observed in emulsions Em 2 and Em 8.
Figure 1. Visual appearance of all developed emulsions after 42 days of storage, with visible contamination observed in emulsions Em 2 and Em 8.
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Figure 2. A 96-well plate with a suspension containing L. rhamnosus GG and different concentrations of preservative. DHA BA—used for the formulations’ development. (0.2: the highest tested preservative conentration, K+: growth control, K−: sterility control).
Figure 2. A 96-well plate with a suspension containing L. rhamnosus GG and different concentrations of preservative. DHA BA—used for the formulations’ development. (0.2: the highest tested preservative conentration, K+: growth control, K−: sterility control).
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Figure 3. FTIR spectra of Lactobacillus rhamnosus GG: (a) segmented spectrum; (b) spectrum of Lactobacillus rhamnosus GG with characteristic peaks marked.
Figure 3. FTIR spectra of Lactobacillus rhamnosus GG: (a) segmented spectrum; (b) spectrum of Lactobacillus rhamnosus GG with characteristic peaks marked.
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Pharmaceutics 18 00199 g004aPharmaceutics 18 00199 g004b
Figure 4. FTIR spectra of all emulsions on days 1, 7, 14, 21, 28, 35 and 42, with the regions corresponding to the characteristic segments of the L. rhamnosus GG spectrum highlighted.
Figure 4. FTIR spectra of all emulsions on days 1, 7, 14, 21, 28, 35 and 42, with the regions corresponding to the characteristic segments of the L. rhamnosus GG spectrum highlighted.
Pharmaceutics 18 00199 g004aPharmaceutics 18 00199 g004b
Table 1. The formulation components of the developed emulsions.
Table 1. The formulation components of the developed emulsions.
Substance NamePhaseEm 1Em 1kEm 2Em 2kEm 3Em 3kEm 4Em 4kEm 5Em 5kEm 6Em 6kEm 7Em 7kEm 8Em 8k
% (w/w)
Cetyl alcoholoil2.02.0
Biobased 6.0 6.0
BTMS 6.0 6.0
Stearic acid3.03.0
Olivem 10004.04.08.0 8.0
Almond oil6.06.0 10
Glyceryl stearate citrate5.05.0
White beeswax3.53.5 6.0 5.0
Carnauba wax1.01.0 5.0 5.0 5.0
Biobased
Linseed oil 6.0 6.0 6.0
Lanolin 4.0 10 10 10 10 4.0
Beta glucan 3.0 3.0 3.0 3.0
Dimethicone 3500.50.5 0.5
Dlyceryl monostearate 8.0 8.0
GSC 2.0 2.0
DHA BA 0.1 0.1 0.1 0.1 0.1 0.1 0.1 0.1
Propylene glycolwater5.05.0 9.5 10
Glycerine 4.0 5.0 10 10 4.0
Inulin 4.0 4.0
Deionised water707072 64 69 64 66 62 68
BTMS—behentrimonium methosulfate, GSC—glyceryl stearate citrate, k—formulations supplemented with a preservative.
Table 2. pH values of all formulated emulsions (EM). SD: standard deviation.
Table 2. pH values of all formulated emulsions (EM). SD: standard deviation.
FormulationEm 1Em 2Em 3Em 4Em 5Em 6Em 7Em 8
pH value
1st day		5.406.314.405.295.003.653.875.92
SD0.010.030.030.020.020.020.030.02
pH value
30th day		5.306.354.555.554.953.753.996.01
SD0.020.030.010.020.030.020.010.01
Table 3. The type of obtained emulsion.
Table 3. The type of obtained emulsion.
FormulationConductometric MeasurementsMethyl Orange/Sudan II StainingDrop Dilution TestEmulsion Type
Em 1, Em 2, Em 3, Em 4, Em 5, Em 6, Em 7, Em 8electrically
conductive		homogeneous colouring with methyl orangemixes well with watero/w
Table 4. Bacterial count obtained in viability test of each prepared emulsion [CFU/mL].
Table 4. Bacterial count obtained in viability test of each prepared emulsion [CFU/mL].
FormulationBacterial Count
[CFU/mL]
Em 11.82 × 108
Em 1k1.99 × 106
Em 22.75 × 109
Em 2k3.25 × 108
Em 33.66 × 109
Em 3k2.49 × 109
Em 42.98 × 109
Em 4k3.36 × 107
Em 52.17 × 109
Em 5k3.78 × 1012
Em 63.15 × 1010
Em 6k-
Em 72.98 × 108
Em 7k3.42 × 108
Em 82.55 × 109
Em 8k3.85 × 109
k—represents the addition of the preservative DHA BA to the formulation.
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MDPI and ACS Style

Gasztych, M.; Dudek-Wicher, R.; Brzozowski, D.; Dołowacka-Jóźwiak, A.; Musiał, W. Development and Physicochemical Characterisation of Probiotic Emulsions Containing Lactobacillus rhamnosus for Potential Dermal Applications. Pharmaceutics 2026, 18, 199. https://doi.org/10.3390/pharmaceutics18020199

AMA Style

Gasztych M, Dudek-Wicher R, Brzozowski D, Dołowacka-Jóźwiak A, Musiał W. Development and Physicochemical Characterisation of Probiotic Emulsions Containing Lactobacillus rhamnosus for Potential Dermal Applications. Pharmaceutics. 2026; 18(2):199. https://doi.org/10.3390/pharmaceutics18020199

Chicago/Turabian Style

Gasztych, Monika, Ruth Dudek-Wicher, Dawid Brzozowski, Arleta Dołowacka-Jóźwiak, and Witold Musiał. 2026. "Development and Physicochemical Characterisation of Probiotic Emulsions Containing Lactobacillus rhamnosus for Potential Dermal Applications" Pharmaceutics 18, no. 2: 199. https://doi.org/10.3390/pharmaceutics18020199

APA Style

Gasztych, M., Dudek-Wicher, R., Brzozowski, D., Dołowacka-Jóźwiak, A., & Musiał, W. (2026). Development and Physicochemical Characterisation of Probiotic Emulsions Containing Lactobacillus rhamnosus for Potential Dermal Applications. Pharmaceutics, 18(2), 199. https://doi.org/10.3390/pharmaceutics18020199

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