Equine Colostrum-Derived Mesenchymal Stromal Cells: A Potential Resource for Veterinary Regenerative Medicine

1. Introduction

Colostrum is the first, thick, nutrient-rich secretion produced by the female mammary gland during the final 1-2 weeks of pregnancy, as a response to hormonal changes [1,2]. In equine species, due to the epitheliochorial nature of the placenta, colostrum is vital for foal survival, serving as the newborn's sole natural source of immunoglobulins, fulfilling essential nutritional needs and supporting gastrointestinal tract maturation [2,3,4].

The mammary gland is a highly metabolically active organ that undergoes cyclic phases of proliferation and hypertrophy during pregnancy and lactation, followed by post-lactation involution [5,6,7]. These processes are thought to be supported by mammary stromal cells (MaSCs), which possess the potential to differentiate into the various cellular components essential for lactation [8,9].

The functional unit of the mammary gland is the epithelium, composed of luminal and myoepithelial cells [10,11], both of which are thought to originate from a common progenitor derived from MaSCs [8,9,12] These MaSCs are likely of epithelial origin and, probably, they are present in milk, having been shed from the ductal and luminal epithelial layers. This shedding may result from the increased turnover of secretory tissue or mechanical shear forces induced by the continuous cycles of milk filling and emptying during lactation [7,13]. In 2007, Cregan et al. [14] provided the first evidence that breast milk contains cells with stem cell/progenitor properties. They showed that cell colonies established in culture from breast milk contained cells positive for the mammary stem cell marker CK5 and the general stem cell marker Nestin [14]. The ex vivo presence of CK5- and Nestin-positive cells in breast milk was subsequently confirmed by other investigators [7,15] and further expanded with the identification of breastmilk stem cells (BSCs) [16]. BSCs not only exhibit self-renewal capabilities in culture but also express pluripotency-associated genes, including the core transcription factors OCT4, SOX2 and NANOG, along with downstream targets KLF4, REX1, and GDF3 [16]. Remarkably, these BSC subpopulations appear highly plastic, capable of differentiating into cell types derived from all three germ layers, including neurons, glia, hepatocytes, pancreatic cells, cardiomyocytes, osteoblasts, chondrocytes, and adipocytes [10,16,17,18].

In addition to BSCs, various types of stem cells, including MSC-like (Mesenchymal stromal cells)-like cells, have been identified in colostrum and milk across different species [7,10,12,16,19,20,21,22,23,24,25,26]. Patki et al. [10] confirmed the mesenchymal characteristics of human breast milk-derived stem cells, which exhibit dual properties by being positive for both the epithelial marker E-Cadherin and the mesenchymal marker SMA (Smooth Muscle Actin), supporting the hypothesis that these cells may be of myoepithelial origin [10]. Moreover, these human breast milk-derived MSCs have been shown to possess the ability to differentiate into various human cell types, underscoring their potential applications in regenerative medicine [10].

These findings pave the way for further research into the use of milk and colostrum-derived stem cells as a non-invasive, accessible source for therapeutic applications in regenerative medicine, including in veterinary medicine. In vivo, these stem cells contribute not only to the structural remodeling of the mammary gland and the development of neonatal tissues but also play a role in epigenetic regulation in the offspring [26]. Studies in rats and mice demonstrated that maternal stem cells can survive in the gastrointestinal tract, enter the bloodstream, and integrate into various neonatal organs, where they differentiate into functional cells in a process known as microchimerism [27,28]. This process, which begins in utero and continues through breastfeeding, suggests that milk- and colostrum-derived stem cells not only support neonatal development but may provide long-term immune benefits and foster unique biological tolerance between mother and offspring [12,29].

Among stem cell types, MSCs are widely used for cell therapy due to their advantages, including self-renewal, multi-lineage differentiation, low risk of teratoma formation, and low immunogenicity [30,31]. MSCs can be derived from various tissues, such as bone marrow [32], adipose tissue [33], Wharton’s jelly [34], placenta [35], amniotic membrane [36], and amniotic fluid [37]. In 2006, the International Society for Cellular Therapy (ISCT) established the "golden criteria" for MSCs characterization. These criteria include fibroblast-like plastic-adherent morphology, high expression of mesenchymal biomarkers (CD73, CD90, CD105), minimal expression of hematopoietic or immune biomarkers (e.g., CD31, CD34, CD45, HLA-DR), and the ability to differentiate into adipocytes, osteoblasts, and chondrocytes [38]. Moreover, in response to stimuli, MSCs secrete a plethora of bioactive molecules with anti-inflammatory [39,40], immunomodulatory [39,41,42,43], and other therapeutic effects, such as inhibiting cell death, reducing fibrosis, promoting vascularization, and exhibiting antimicrobial properties [42,44,45].

Over the past two decades, studies have extensively characterized human milk-derived stem cells [5,6,7,10,14,16,18,22,23,46,47,48,49,50,51,52,53] and their therapeutic potential across various pathological conditions [54,55,56,57,58,59].

In veterinary species, studies have primarily focused on isolating and characterizing MSC-like cells from bovine, swine, and rabbit milk [19,60,61], highlighting their regenerative, pro-angiogenic [62], and antimicrobial [61] properties with potential applications in neonatal health and veterinary medicine.

However, research on equine mammary and milk-derived stem cells remains limited. To date, only one study has successfully isolated and characterized mammary stem cells (MaSCs) from equine mammary gland tissue [63]. Subsequent investigations have explored the role of extracellular vesicles [64] and microRNAs [65] in MaSC regulation, the development of equine mammary organoids for comparative research [66], and species-specific mechanisms of mammary cancer resistance [67].

Nevertheless, the presence and therapeutic potential of MSCs in equine colostrum remain largely unexplored. This study aims to isolate and characterize MSCs for the first time from equine colostrum collected immediately after delivery, laying the groundwork for future applications in veterinary regenerative medicine.

2. Materials and Methods

2.1. Materials

Chemicals were obtained from Sigma-Aldrich (Merck KGaA, Darmstadt, Germany). FBS (Fetal Bovine Serum) is branded GIBCO™ (Thermo Fisher Scientific, Waltham, MA, USA). Plastics were from Falcon™ (Corning Inc., NY, USA) and SarstedtTM (Sarstedt AG & Co. KG, Nümbrecht, Germany) unless otherwise stated.

2.2. Samples

Colostrum samples (n=6) were collected from mares hospitalized at the Equine Perinatology and Reproduction Unit of the Equine Clinical Service, Department of Veterinary Medical Sciences, University of Bologna (Italy), for delivery management. Samples were aseptically collected into sterile 15 mL tubes immediately after foaling. Colostrum quality, indicative of its IgG concentration [68], was assessed using a refractometer equipped with a Brix scale [68,69]. The portion of each sample not utilized for quality assessment was allocated for cell isolation. It was stored at +4°C for a maximum of 12 hours before being transported to the Laboratory of Animal Reproduction and Biotechnologies (Equine Clinical Service, Department of Veterinary Medical Sciences, University of Bologna, Italy). All procedures were conducted in compliance with the guidelines approved by the Animal Welfare Committee (CoBA) of the University of Bologna (protocol no. 4630/23).

2.3. Equine Colostrum-Derived MSCs Isolation and Culture

Equine colostrum-derived MSCs (C-MSCs) were isolated as previously described by Widjaja et al. [70] with some modifications. Briefly, colostrum was mixed and diluted 1:1 with Dulbecco’s polyphosphate-buffered saline (DPBS) plus antibiotics (100 IU/mL penicillin, 100 g/mL streptomycin) and antimycotics (2.5 mg/L Amphotericin B) in 50 mL conical tube and centrifuged at 1881×g for 20 min at 20°C. After centrifugation, the supernatant and the fat layer were discarded and the cell pellet was re-suspended in 2 mL DPBS, then transferred into a sterile 15 mL conical tube. The cell pellet was re-suspended and centrifuged at 1881×g for 10 min at 20°C. This operation was repeated twice. Finally, the cellular pellet was re-suspended in complete culture medium (DMEM F-12 + 10% FBS + 100 IU/mL penicillin + 100 g/mL streptomycin). Cells were plated into 25 cm² culture flasks and incubated in 5% CO₂ at 38.5°C, in a humidified atmosphere (Passage 0). After 24 h, the culture medium was completely replaced, and non-adherent cells were removed. The culture medium was changed every 3 days until reaching a confluence of 80-90%. At this time, cells were dissociated using a 0.25% Trypsin-EDTA (Ethylenediaminetetraacetic acid) solution, counted using a Neubauer improved chamber, and cryopreserved as described by Merlo et al. [71]. Briefly, cells in 0.5 mL of FBS were put in a 1.5 mL cryogenic tube at 4°C. After 10 min, the cell suspension was diluted 1:1 with FBS + 16% DMSO (Dimethyl sulfoxide, final concentration 8% DMSO) and maintained for a further 10 min at 4°C. Then the cryogenic tube was set to −80°C for 24 h in Mr. Frosty (Nalgene) and finally stored in liquid nitrogen.

Equine C-MSCs were thawed at 37°C in 20 mL DMEM + 10% FBS, then centrifuged at 470×g at 25°C for 10 min. The pellet was re-suspended in 1 mL of complete culture medium and cell concentration and viability were evaluated by staining cells with Trypan blue (0.4%) solution. Cells were seeded in a 25 cm² flask (5000 cells/cm²) as “Passage 1” (P1). Third-passage (P3) cells were used for in vitro tests.

2.4. Characterization of Colostrum-Derived MSCs

When the confluence of 80–90% at P2 was reached, cells from all six colostrum samples were detached from the flask, and viability and concentrations were determined as described above. The cells were subsequently seeded to evaluate their growth rate through Population Doubling Time (PDT) analysis and for characterization via the colony-forming unit (CFU) assay, adhesion and migration assay, and tri-lineage in vitro differentiation. Furthermore, CD marker expression was evaluated by RT-PCR. For all samples, each test was carried out in three replicates.

2.4.1. Population Doubling Time (PDT) Analysis

To determine growth rate, C-MSCs were seeded in 6-well plate at a cell density of 5000 cells/cm². Cell number was assessed every day for 7 days in triplicate. The mean number of cells was calculated and plotted on a semi-log curve against culture time to generate a growth curve. The population doubling time (PDT) was calculated from the log phase of the growth curve using the equation described by Bezerra et al. (2022) [72]:

PDT=(t×log2) / log (Nt / N0)

(1)

where t represents the period of cultivation in days, N0 represents the initial number of cells, and Nt represents the number of cells in a specific period of culture.

2.4.2. CFU (Colony-Forming Unit) Assay

For determining the ability of cells to form colonies, 1 × 10² cells were cultured for 4 weeks in 6-well plates. Colonies were then fixed in 4% paraformaldehyde at RT for 1 h and stained with Giemsa 0.1% stain (30 min). Using an inverted light microscope (Eclipse TE 2000-U, Nikon Instruments Inc., Tokyo, Japan), the operator counted colonies formed by at least 16–20 nucleate cells.

2.4.3. Adhesion and Migration Assays

To determine whether C-MSCs preserved their adhesion capability, a spheroid formation assay was performed. Differently from the cell-substratum adhesion, performed on monolayer cultures adherent to rigid substrates, this test gives information about the direct cell-cell adhesion architecture found in normal tissues. Cells were cultured through the ‘hanging-drop’ method in Corning spheroid microplates (96-well, black/clear bottom round, Ultra-Low Attachment surface, Corning Inc., NY, USA, 5000 cells/25 µL drop). Bright-field images were acquired by a CCD camera (DS-Fi2, Nikon, Tokyo, Japan) mounted on an inverted light microscope (Eclipse TE 2000-U, Nikon Instruments Inc., Tokyo, Japan) using 4× magnification every 24 h from the seeding to the complete spheroid formation after 96 h. Starting from the binary masks obtained by Fiji software (ImageJ, v. 1.8.0_345/1.54g), the volume of each spheroid was computed using ReViSP (sourceforge.net/projects/revisp/, v. 2.3) [73], a software specifically designed to accurately estimate the volume of spheroids and to render an image of their 3D surface.

To assess cell migration potential, a scratch assay (or wound-healing assay) was carried out, as previously described by Liang et al. [74]. Briefly, P3 cells were seeded (5000 cells/cm²) in 6-well plates. At 80 to 90% confluence the cell monolayer was scraped using a 200 μL pipette tip. The cells were then washed twice with DPBS to remove any detached cells, and the plate was incubated under standard culture conditions. Images were captured at the same two locations of each well, both immediately after the tip-scratch (time 0 = t0) and every 24 h over a 96 h period (0, 24, 48, 72, 96 h). The distances of each scratch closure were measured by Fiji software (ImageJ, v. 1.8.0_345/1.54g), and the migration percentages (Wound Closure %) were calculated using the following formula [75]:

[(distance at t0−distance at tn *100]/distance at t0

(2)

where tn corresponds to the distance at each specific time point analyzed.

2.4.4. Multi-Lineage In Vitro Differentiation

In vitro differentiation potential toward osteogenic, adipogenic, and chondrogenic lineages of C-MSCs was also evaluated. Cells were seeded in 24-well plates at a density of 5000 cells/cm² and cultured until subconfluency. Then, the culture medium was replaced, and cells were cultured for 3 weeks under specific induction media (Table 1) [76]. As a negative control, an equal number of cells was cultured in expansion medium. The specific induction media and expansion medium were replaced twice a week. To assess differentiation, cells were fixed with 4% paraformaldehyde at room temperature (RT) for 1 h. Differentiation was visualized using Oil Red O for adipogenic vacuoles, Alcian Blue for sulfated proteoglycans, and Alizarin Red S for calcium deposits. Bright-field images were captured using a CCD camera (DS-Fi2, Nikon, Tokyo, Japan) mounted on an inverted light microscope (Eclipse TE 2000-U, Nikon Instruments Inc., Tokyo, Japan) at 10× magnification.

Quantification of positively stained areas was performed using Fiji software (ImageJ, v. 1.8.0_345/1.54g) following the methodology described by Heyman et al. (2022) [77]. The "ColourDeconvolution2" plugin was applied to separate the stain and background channels, which were then converted to 8-bit grayscale images and binarized using auto-thresholding. The positively stained areas were measured, and the differentiation ratio (DR) was calculated as the ratio between the areas of the stain-positive signal and the background.

To compare differentiation across samples, a normalized DR was determined for each sample by dividing the mean DR of the differentiated wells by the mean DR of the non-induced controls.

2.4.5. Molecular Characterization

For the molecular characterization of C-MSCs, total RNA was extracted from snap-frozen cells using the NucleoSpin® RNA Kit (Macherey-Nagel, Düren, Germany) following the manufacturer’s protocol. Residual genomic DNA was removed by treating the RNA with Amplification-Grade DNase I (Invitrogen, Thermo Fisher Scientific, Waltham, MA, USA). Complementary DNA (cDNA) was synthesized from the purified RNA using the RevertAid Reverse Transcriptase Kit (Thermo Fisher Scientific, Waltham, MA, USA).

The resulting cDNA was directly subjected to PCR using the 2× EpiArt™ HS Taq Master Mix (Vazyme Biotech Co., Ltd., Nanjing, China). The expression of genes encoding mesenchymal stromal cell markers (CD90, CD73), hematopoietic markers (CD45, CD34), and major histocompatibility complex markers (MHC-I, MHC-II) was analyzed. To ensure the proper expression of samples, GAPDH was used as a housekeeping gene. The specific primers employed for amplification are listed in Table 2 [78,79,80].

PCR products were resolved on a 2% agarose gel stained with ethidium bromide. Electrophoresis was conducted to verify the expected band sizes, and gels were visualized and photographed under ultraviolet light using a UVP PhotoDoc-It Imaging System (UVP, LLC, Upland, CA, USA). The presence of discrete bands corresponding to the expected sizes confirmed positive expression, while the absence of bands indicated a negative result. A 50 bp DNA Ladder (Thermo Fisher Scientific, Waltham, MA, USA) was used as a molecular size marker.

2.4.6. Statistical Analysis

Data are reported as mean ± standard deviation (SD). Statistical analyses and graph generation were performed using GraphPad Prism software (version 9.0.0). Data were analyzed, for normal distribution, using a Shapiro–Wilk test. Population doubling times (PDTs) and colony numbers were analyzed using one-way ANOVA, followed by Tukey’s post-hoc test for pairwise comparisons. Proliferation rates and wound closure percentages were evaluated with two-way ANOVA, followed by Tukey’s post-hoc test for comparisons between groups. Volumes of 3D spheroids at 96 hours and differentiation ratio (DRs) were analyzed using the Kruskal-Wallis test, followed by Dunn's multiple comparisons test. Results were considered significant when the probability of their occurrence due to chance alone was less than 5% (p < 0.05).

3. Results

3.1. Isolation and Morphological Characterization of Equine Colostrum-Derived Cell

Equine colostrum cells (n=6; ~20 mL/sample) were successfully isolated following the protocol detailed in the Materials and Methods section. Isolation was based solely on the cells' ability to adhere to plastic. Phase-contrast microscopy (Figure 1) revealed a morphologically heterogeneous population, comprising cells with a round/polyhedral, epithelial-like appearance, alongside spindle-shaped cells characteristic of Mesenchymal Stromal cells (MSCs). Cell colonies became visible within the first 24 hours and reached confluence in 10.17 ± 1.17 days, with a cell yield ranging from 10.4 to 36.4 × 10³ cells/mL of colostrum.

A detailed summary of the colostrum samples, including volume, Brix Index at collection, number of cells isolated and time required to reach confluence, is reported in Table 3.

Undifferentiated cells were cultured up to passage three (P3) across all samples, with no observable changes in morphology during the culture period or after cryopreservation and thawing. Post-thaw viability, determined by Trypan Blue stain, at P3 exceeded 70%, while the initial viability was over 95%.

3.2. Characterization of Colostrum-Derived MSCs

3.4.1. Population Doubling Time (PDT) Analysis

To assess the growth efficiency and proliferation rate of successfully established colostrum-derived MSC (C-MSC) cultures, population doubling times (PDTs) were calculated. The semi-logarithmic growth curves (Figure 2) revealed variability in the growth patterns across the samples. Three samples (4-5-6) exhibited a brief lag phase of 1-2 days, followed by a significant and linear increase in cell concentration, while three samples demonstrated prolonged lag phases (1-2) or a slower overall growth rate (3). The mean PDT values, which confirm the variability between samples, are summarized in Table 3 and illustrated in Figure 2. One sample (3) exhibited the longest PDT (6.21± 1.84 days), indicating reduced proliferative efficiency, which was significantly different from the mean PDT of the other sample (2 ± 0.7 days, p < 0.0001; Figure 2). In contrast, 2/6 samples (2-4) showed the shortest PDTs, reflecting a higher proliferation rate (1.42 ± 0.28 and 1.22 ± 0.08, respectively). Overall, the sample effect on both growth rate and PDT was highly significant (p < 0.0001).

3.4.2. CFU (Colony-Forming Unit) Assay

The Colony Forming Unit (CFU) assay was performed to evaluate the self-renewal capacity of C-MSCs. At P3, MSCs from all six samples were seeded in 6-well plates at a density of 1 × 10² cells/well and cultured for 4 weeks. The resulting colonies were categorized as "dispersed," characterized by a vague macroscopic appearance and scattered cells under the microscope, as opposed to dense, compact colonies with the typical "fingerprint" pattern (Figure 3) [81]. Colonies were counted in triplicate, and the average number of colonies formed is summarized in Table 3 and presented in Figure 3, revealing significant sample-related variability (p < 0.0001). Two samples (2-4) demonstrated the highest clonogenic potential, forming significantly more CFUs than the other samples (71 ± 16.7 and 59.3 ± 7.37, respectively; p < 0.0001). Conversely, one sample (6) exhibited the lowest CFU count (13 ± 3.6), showing a statistically significant difference compared to samples 2, 4, and 5 (p < 0.0001, Figure 3). No significant differences were detected among samples 1, 3, and 5 (21 ± 2, 34.7 ± 11, and 40.3 ± 5.5, respectively; Figure 3).

3.4.3. Adhesion and Migration Assays

To assess the cell-cell adhesion capacity of C-MSCs, at P3 of in vitro culture, cells were cultured using the hanging-drop method in ultra-low attachment microplates (5000 cells/25 µL drop). Except for samples 1 and 3, all tested MSC populations demonstrated the ability to form three-dimensional spheroid-like structures. Spheroid formation was evident after 24 hours, and by the fourth day of incubation, most C-MSCs had developed stable, compact, and rigid spheroids with a spherical morphology (Figure 4).

In contrast, samples 1 and 3 failed to generate spheroids, instead forming loosely compact cellular sheets that remained easily dissociable, suggesting a lack of spheroid-forming capacity (Figure 4). Bright-field images were acquired at multiple time points (24, 48, 72, and 96 hours) and analyzed using ReViSP, an open-source voxel-based 3D image analysis tool capable of estimating spheroid volume starting from binary masks and rendering a three-dimensional surface representation (Figure 4).

Quantitative analysis of spheroid volume revealed dynamic structural changes over time, characterized by progressive compaction and a corresponding reduction in volume between 48 and 96 hours. By the end of the incubation period (4 days), spheroid volumes had stabilized, with no significant differences detected between the samples.

Given the growing interest in the role of MSCs in tissue repair [82], we aimed to evaluate their migratory capacity using the scratch assay, a highly quantifiable, adaptable, and cost-effective method. The percentage of wound closure was assessed at different time points (24, 48, 72, and 96 hours) and plotted against time, as shown in Figure 5.

Quantitative analysis revealed significant differences in wound closure percentages among the groups (Two-way ANOVA, p < 0.0001). Notably, sample 2 demonstrated the highest migratory capacity, achieving complete wound closure (100%) at 72 hours, significantly outperforming all other groups. In contrast, samples 1, 4, 5, and 6 exhibited a more gradual increase in wound closure over time, without exceeding approximately 30% at 96 h (Figure 5).

Inter-group comparisons at individual time points revealed intermediate wound closure dynamics for sample 3. Although its performance showed modest but non-significant improvements over the slower-migrating groups (1, 4, 5, and 6), it remained significantly lower than sample 2 (p < 0.001, Figure 5).

3.4.4. Multi-Lineage In Vitro Differentiation

According to the criteria established by the International Society for Cell & Gene Therapy (ISCT) [38], at P3 of in vitro culture, C-MSCs were evaluated for their potential to differentiate into osteogenic, chondrogenic and adipogenic lineages under specific induction conditions.

After 3 weeks of adipogenic induction, all isolated cells exhibited adipogenic differentiation, as demonstrated by a morphological transition from spindle-shaped to round or oval-shaped cells. This was accompanied by the presence of intracytoplasmic lipid droplets, confirmed by positive staining with Oil Red O (Figure 6). In contrast, control MSCs maintained their spindle-shaped morphology, forming a monolayer without staining with Oil Red O, confirming the absence of adipogenic differentiation (Figure 6).

Chondrogenic differentiation was observed in C-MSCs from all samples after 21 days of culture in a chondrogenic medium. This was marked by a morphological change from spindle-shaped cells to larger, round cell aggregates, along with the secretion of an extracellular matrix enriched in sulfated proteoglycans, a defining trait of cartilage tissue. Positive Alcian Blue staining (Figure 6) confirmed the presence of sulfated proteoglycans, while no staining was detected in the control wells (Figure 6), indicating a lack of chondrogenic differentiation.

Following osteogenic induction, C-MSCs from all samples exhibited matrix calcium deposition, as demonstrated by positive Alizarin Red S staining, a key marker of osteogenic differentiation (Figure 6). No calcium deposition was observed in the control groups (Figure 6).

Quantification of positively stained areas using ImageJ software showed no statistically significant differences in adipogenic, chondrogenic, or osteogenic differentiation capacities between the groups (p > 0.005).

3.4.5. Molecular Characterization

Following the guidelines of the International Society for Cell & Gene Therapy (ISCT) [38], the molecular characterization of C-MSCs was performed by analyzing the expression of genes encoding MSC markers (CD90, CD73), hematopoietic markers (CD45, CD34), and major histocompatibility complex markers (MHC-I, MHC-II). To ensure the proper expression of samples, GAPDH was used as a housekeeping gene (Figure 7). The PCR results are summarized in Table 4. Qualitative PCR analysis demonstrated that MSCs isolated from all six colostrum samples expressed the MSC markers CD90 and CD73, while hematopoietic markers CD34 and CD45 were absent at P3 of in vitro culture (Figure 7). Additionally, MHC-I expression was weakly positive in all samples except for sample 1, where it was absent, while MHC-II expression was consistently negative across all samples (Figure 7).

4. Discussion

The expanding field of regenerative medicine has intensified the search for innovative and advantageous stem cell sources. Recent studies have demonstrated that milk, including colostrum, harbors a heterogeneous population of cells, including stem and progenitor cells with significant regenerative potential [5,6,7,10,14,16,18,22,23,46,47,48,49,50,51,52,53].

While interest in the therapeutic applications of milk- and colostrum-derived stem cells in veterinary medicine is increasing, little is known about the presence and characteristics of mesenchymal stem/stromal cells in equine colostrum.

This study aimed to isolate and characterize equine colostrum-derived MSCs (C-MSCs) to validate colostrum as a promising, non-invasive stem cell source. Building on our previous findings on fetal- and adipose-derived MSCs [83,84], we sought to compare C-MSCs with these well-characterized sources, assessing their potential for veterinary and translational medicine.

Our findings confirmed the successful isolation of C-MSCs, with a cell yield ranging from 10.4 to 36.4 × 10³ cells/mL. These cells exhibited plastic adherence and a fibroblast-like morphology, consistent with prior reports on MSCs isolated from human breast milk [47,85]. The presence of a heterogeneous cell population in colostrum, including both fibroblast-like and round-shaped cells, aligns with studies on bovine and human milk-derived MSCs, where an epithelial-to-mesenchymal transition (EMT) has been observed during culture [10,47,60].

As reported in studies on human breast milk MSCs, the lactation stage significantly influences stem cell quantity and viability, with a higher concentration found in early lactation[6,47,48]. This variation may explain discrepancies in isolation success across studies. Like findings by Fan et al. [7], who encountered difficulties in culturing breast milk-derived MSCs from later lactation stages, our results suggest that the early postpartum period is optimal for MSC isolation.

MSC characterization was performed at passage three, when the culture reached a homogeneous population, in accordance with previous studies [86,87,88]. Population doubling time (PDT) is a key parameter for assessing MSC proliferative potential in therapeutic applications [89,90], exhibited notable inter-sample variability. While some C-MSC samples exhibited a rapid expansion, others displayed a slower proliferation rate, with an average PDT of 2.7 ± 0.6 days. As previously reported, PDT is highly dependent on the tissue of origin and culture conditions [91,92]. Our findings align with those observed before for Wharton’s jelly MSCs (WJ-MSCs, 2.7 ± 0.9 days) [83], while C-MSCs proliferated faster than bone marrow MSCs (BM-MSCs, 3.6 ± 1.4 days) [83] but slower than amniotic fluid MSCs (AF-MSCs, 27.13 ± 2.6 hours) [91], adipose tissue MSCs (AT-MSCs, 2.2 ± 1.1 days), and umbilical cord blood MSCs (UCB-MSCs, 2.4 ± 1.3 days) [83] . These results emphasize the variability in MSC proliferation dynamics based on tissue source and culture conditions, even when carried out in the same laboratory.

The colony-forming unit (CFU) assay confirmed the self-renewal capacity of C-MSCs. Our results contrast with those reported by Fan et al.; indeed, authors failed to obtain CFU formation from human breast milk MSCs, possibly due to differences in lactation stage, culture conditions, or intrinsic biological differences between species [7]. The ability of C-MSCs to form colonies suggests a robust self-renewal potential, comparable to that of MSCs derived from equine fetal annexes and other postnatal sources [93]. Further comparative studies are required to elucidate the specific factors influencing CFU efficiency across different species and MSC sources.

Spheroid formation was assessed using the hanging-drop method, with most samples forming compact spheroids indicative of strong cell-cell adhesion properties. Notably, smaller spheroids correlated with increased adhesion ability, a characteristic previously described in fetal-derived MSCs, such as WJ-MSCs, compared to adult tissue-derived MSCs [83]. This suggests that C-MSCs share adhesion properties similar to those observed in fetal MSCs.

The migratory potential of MSCs is a key characteristic that influences their capacity for systemic application and integration into host tissues during therapeutic interventions [30]. The scratch assay demonstrated efficient C-MSC migration, achieving approximately 30% wound closure at 96 hours. Comparative studies in our laboratory showed similar migration efficiencies among MSCs from various sources, including amniotic membrane (AM-MSCs, 34.14 ± 4.51%) and Wharton’s jelly (WJ-MSCs, 38.20 ± 2.88%) [84]. Comparable migration rates were observed among MSCs from WJ, UCB, BM, and AT, though AT-MSCs exhibited notable inter-sample variability [83]. These findings indicate that C-MSCs possess migration capabilities like fetal annex-derived MSCs, reinforcing their regenerative potential.

C-MSCs exhibited a multipotent differentiation capacity, successfully differentiating into osteogenic, chondrogenic, and adipogenic lineages, as confirmed by specific staining performed after 21 days of culture in the appropriate differentiation medium. These results align with prior studies on MSCs from human breast milk [10,16] and bovine milk [60], which demonstrated similar tri-lineage differentiation potential. The mesodermal differentiation ability of C-MSCs further validates their identity as MSCs, in accordance with the criteria established by the International Society for Cellular Therapy (ISCT) [38]. Notably, unlike MSCs from adult equine tissues [94] and our previous findings on equine AF-MSCs, UCB-MSCs, and WJ-MSCs [87], C-MSCs displayed minimal donor-to-donor variability.

To confirm the mesenchymal identity of equine C-MSCs, we evaluated MSC surface marker expression. These cells expressed CD90 and CD73 while lacking hematopoietic markers CD34 and CD45, consistent with findings on MSCs from AF, CB, and WJ [87]. Similar expression patterns have been reported in MSCs from milk across species, including rabbits, bovines, and humans, with high levels of CD29, CD166, CD44, and CD105, and an absence of CD45 and CD34 [10,19,22,47,59,60,95].

A key feature of MSCs is their immunomodulatory potential, which is largely influenced by the expression of major histocompatibility complex (MHC) molecules. In our study, MHC-I expression varied among samples, with some exhibiting weak expression and one lacking it entirely. Previous studies on equine BM-MSCs have shown MHC-I expression with heterogeneous MHC-II expression [96,97]. In horses, MSCs that express MHC-I but not MHC-II have been shown to avoid triggering T-cell proliferation [97]. The absence of MHC-II expression is therefore a crucial factor for potential allogeneic applications, as it enables MSCs to evade immune rejection [97,98]. Our findings confirm that equine C-MSCs did not express MHC-II, in agreement with our previous studies on equine WJ- and AT-MSCs [84] and recent findings on AF-MSCs [91].

The successful isolation and characterization of equine C-MSCs highlight their potential as a novel, non-invasive source of regenerative cells for veterinary medicine. Their ease of collection, multipotency, and immunomodulatory properties position them as strong candidates for equine regenerative therapies, with possible translational implications for human medicine. However, the variability observed among samples from different donors necessitates validation with a larger sample size. Future studies should also investigate their in vivo therapeutic potential and interactions with host tissues.

5. Conclusions

This study provides the first comprehensive characterization of mesenchymal stromal cells isolated from equine colostrum (C-MSCs), confirming their mesenchymal phenotype, self-renewal ability, migration potential, and multipotency. These findings pave the way for further investigations into their applications in veterinary regenerative medicine, with promising translational implications for human therapies.