Sheep Placenta Cotyledons: A Noninvasive Source of Ovine Mesenchymal Stem Cells

Abstract

Sheep are one of the most frequently used large animal models in stem cell research. However, minimal invasive or noninvasive sources of mesenchymal stem cells (MSCs) in sheep are scarce. In the light of the principles of the 3Rs (reduce, refine, replace), it would therefore be desirable to identify a minimally invasive or noninvasive ovine MSC source. In humans, the chorionic villi of the placenta, which can be noninvasively harvested as part of the afterbirth, have been identified as a rich source of MSCs. Therefore, in the present study, ovine placenta cotyledons, which have similar function and structure to human chorionic villi, were tested for their potential use as a noninvasive source of ovine MSCs. Through mincing of the placental cotyledons, collagenase digestion, and Ficoll density gradient centrifugation, combined with plastic adherence selection, MSCs were successfully isolated. Their morphological, immunophenotypical, and cellular growth characteristics, as well as their proliferation, differentiation, and migration potential, were evaluated and compared to the currently best-researched MSC source, bone marrow-derived stem cells. Ovine cotyledons were shown to be a reliable, abundant source for the noninvasive, pain- and risk-free harvest of MSCs. The collection procedure does not interfere with partum or the initial bonding phase between ewes and lambs and is therefore exempt from ethical debate. Ovine placenta cotyledon-derived MSCs exhibit multipotential characteristics and can be cryopreserved for later use.

Introduction

Mesenchymal stem cells (MSCs) have tremendous therapeutic potential, which is harnessed in regenerative medicine to repair damaged tissues and treat degenerative diseases for which there are currently no or only limited therapeutic options. While already many clinical trials exploring the use of MSCs for the treatment of a broad variety of diseases are ongoing, their intrinsic properties and mechanism of action as well as the role of their microenvironment in modulating their behavior and function are not yet fully understood.¹ Therefore, before launching new therapeutic approaches, the efficacy and safety of new treatments need to be tested in preclinical and clinical trials using animal models.² Sheep are among the most frequently used large animal models for a wide range of indications. They are large animals but docile, easy to handle and house, relatively inexpensive, and available in large numbers. Due to their relatively big size, they lend themselves to longitudinal analyses and repeated sampling from individual animals. Based on similarities with humans in weight, bone and joint structure, and bone/cartilage regeneration,^3–17 the use of sheep for orthopedic and stem cell research continues to increase and is even advised by the European Medicines Agency (EMA)¹⁸ and the U.S. Food and Drug Administration (FDA)¹⁹ for testing a variety of orthopedic cellular and biologic therapies.

Bone marrow and adipose-derived stem cells are the best researched stem cell sources of humans and animals. However, a disadvantage of bone marrow-derived MSCs (bmMSCs) is the invasive harvesting methodology.^3,20–26 Adipose tissue is routinely gained through a less but still invasive surgical procedure.^21,26–31

Minimal invasive or noninvasive MSC sources in sheep are scarce. Peripheral blood would be a largely pain- and risk-free source of stem cells. However, as in other species,^17,32–35 the amount of stem cells contained in ovine peripheral blood is relatively low.^34,35 Therefore, there is a need for an abundantly available, reliable, noninvasive MSC source for this frequently used model animal. Perinatal tissues, such as cord blood, umbilical cord tissue, amniotic membranes, or amniotic fluid, are often used as MSC sources.^{13,15,18,19,36–69} They are abundantly available and can be harvested in a noninvasive, ethically inoffensive manner.^40,45,70,71 The cell populations derived from these sources express all typical MSC markers and produce mesodermal progenitor cells with multilineage differentiation potential.^37,72 Access to perinatal MSC sources is pain free and bears no risk for the mother or the newborn.^70,73–75 Above that, juvenile MSCs derived from perinatal tissues provide several advantages over adult MSCs. They are characterized by a higher proliferation rate, longer life span, better quality, and lower immunogenicity.^{45,70–72,76} In addition, juvenile MSCs have a broader differentiation capability toward cell types of endo- or ectodermal origin.^45,63

A disadvantage of perinatal tissues especially from large animal donors may be the higher risk of bacterial or mycotic contamination due to the nonsterile birth canal and the delivery in a farm environment.^44,61 This makes the use of disinfectants on tissue harvest and antibiotics in the cell culture medium mandatory.⁶¹ Therefore, to date, ovine cord blood in sheep was mainly collected during preterm cesarean sections.⁴³ Also, amnion-derived and amniotic fluid-derived MSCs, which were successfully obtained in sheep, were mainly collected during aseptic surgical approaches.^{13,15,18,19,36,41,51,52,59,67}

In humans the chorionic villi of the placenta (Fig. 1A) have been identified as rich source of MSCs.^77–79 It is nowadays agreed that MSCs are of perivascular origin.^80–82 They reside in the so-called stem cell niche, which is typically located around blood vessels and offers an environment for MSCs, in which they remain quiescent until internal or external signals stimulate their replication, migration, differentiation, or secretory activity to repair or replace tissue.^83–86 Chorionic villi serve as junction between the maternal and fetal blood circulation.⁸⁷ Therefore, they contain a lot of blood vessels, which makes them a potentially valuable MSC source. De facto MSCs were confirmed to reside in a vascular/perivascular niche in the placenta chorionic villi.⁷⁷

FIG. 1.

Human and ovine placenta. Schematic drawing of a human placenta (A) and an ovine placenta (B). In ruminants, the chorionic villi are organized into distinct structures called cotyledons (B–F). All drawings by Manuela Miksic/Vetmeduni Vienna. The chorionic villi in cotyledons largely correspond to the human counterparts. Cotyledons (fetal part of the placenta) and caruncles (maternal part of the placenta) form specialized units known as placentomes. Following parturition, the chorionic villi are withdrawn from the maternal caruncles (C) and the fetal membranes are expulsed (D). Hematoxylin and eosin-stained sections of a cotyledon (E, F) showing a high number of blood vessels (potential perivascular niche) containing residual blood (F). Anatomically equivalent structures in B, C and D, are indicated by equal colored boxes. F is a close-up of the boxed area in E. Scale bar = 1.5 cm (D), 4 mm (E), and 200 μm (F), respectively.

The placenta of ruminants is cotyledonary, nondeciduate, and epitheliochorial in contrast to the discoid, deciduate, and hemochorial human placenta. However, the function and structure of chorionic villi largely correspond to the human counterparts.⁸⁷ In ruminants, the chorionic villi are organized in distinct structures called cotyledons (Fig. 1B–F). Cotyledons (fetal part of the placenta) and caruncles (maternal part of the placenta) (Fig. 1B, C) form specialized units known as placentomes.^87,88 Following parturition, the chorionic villi are withdrawn from the maternal caruncles and the fetal membranes are expulsed (Fig. 1C).^87,88

Ovine placental cotyledons obtained on full-term delivery were, to our knowledge, not yet described as MSC source. As part of the afterbirth, they are abundantly available and harvesting them is not associated with any ethical concerns because they are usually discarded as trash. MSCs from placenta cotyledons would be an easily and truly noninvasively accessible source of sheep MSCs. Therefore, the aim of the present study was to test ovine placenta cotyledons for their potential use as noninvasive source of ovine MSCs and to compare the obtained MSCs to bone marrow-derived MSCs, the currently best-researched stem cell source.

Materials and Methods

Collection of the afterbirth and preparation of cotyledons

All afterbirth samples were collected during the same breading season on full-term delivery of the lambs according to the Vetmeduni Vienna Guidelines for Good Scientific Practice. The afterbirths were collected post partum from Merino ewes of the Vetmeduni Vienna teaching herd. They were left in situ until they were spontaneously passed (one to 3 h post partum). After delivery of the lambs, and after the important first bonding phase between the ewes and lambs, the ewe's tails were covered with a rectal glove and the vulva was cleaned and washed with a soap solution containing iodine. When the afterbirth was passed, it was collected directly into a sterile pot with a lid to protect it from the farm environment and potential fungal spores in the air. For sterility reasons, the part of the afterbirth, which had been hanging out of the vulva after delivery, was discarded. All further sample processing was done in a laboratory environment. The afterbirths were transferred into a 1% iodine solution for 2 min before their dissection with sterile scissors and sterile forceps. Between 8 and 12 cotyledons were harvested per afterbirth (n = 5). In case of twins or triplets, the secundines were not divided and assigned to individual lambs but the afterbirth of the entire litter was harvested in toto. The harvested cotyledons were washed twice with sterile saline (Braun, 0.9% NaCl for infusion) and stored at 4°C in phosphate-buffered saline (DPBS+ Ca, Mg; Lonza) containing 1% pen/strep (10,000 U penicillin/10 mg streptomycin/mL; Sigma Aldrich) and 1% amphotericin B (250 μg/mL; Biochrom) for a maximum of 18 h post partum.

One cotyledon of each afterbirth was immediately placed in buffered 4% formalin (ACM, Herba Chemosan Apotheker AG) for histologic analysis. It was embedded in paraffin and 4 μm slices were cut and subjected to hematoxylin (Thermo Scientific) and eosin staining (Merck).⁸⁹

Isolation and culture of cotyledon-derived MSCs

Under a laminar airflow workbench, the cotyledons (between 7 and 11 per afterbirth) were transferred into a Petri dish and all remaining tissues attached to the cotyledons were carefully removed. Then, the cotyledons were minced into small pieces and digested in a 1 mg/mL collagenase (type I; Sigma Aldrich) solution (Dulbecco's modified Eagle's medium [DMEM] low-glucose, without L-glutamine; Lonza) for 1 h under constant stirring at 37°C. The obtained viscous and bloody solution was mixed with medium to stop the collagenase activity and filtered through a cell strainer (100 μm; Greiner BioOne). A density gradient filtration was performed using Ficoll (Ficoll Paque Premium, Ficoll density 1.077 g/mL; GE Healthcare) to separate the mononuclear cell fraction. For this purpose, the filtered suspension was loaded onto the Ficoll in 50-mL falcon tubes (2:1 ratio of cotyledon solution to Ficoll), and centrifuged for 30 min at room temperature at 330 g (without brake). The buffy coat containing the mononuclear cell fraction was carefully transferred into a new 15-mL falcon tube. It was washed with phosphate-buffered saline (PBS) and centrifuged at 400 g for 5 min at room temperature twice. After resuspension in standard culture medium consisting of DMEM (low glucose, without L-glutamine; Lonza), 10% fetal calf serum (FCS, low endotoxin; Sigma Aldrich), 1% L-glutamine (L-alanyl-L-glutamine 200 mM; Biochrom), 1% pen/strep (100 × ; Sigma Aldrich), and 1% amphotericin B (250 μg/mL; Biochrom), the cells were seeded onto T75 culture dishes and cultured in a humidified atmosphere at 37°C and 5% CO₂. The following day the cells were washed to remove nonadherent cells and tissue debris. Medium was further changed twice weekly. Passaging was performed on colony confluence for the first passage and later at 80–90% confluence by trypsinization (trypsin 0.05%/EDTA 0.02%; Biochrom). Cells were frozen and stored in liquid nitrogen until further use at the lowest possible passage (P1–2).

Harvest, isolation, and culture of bmMSCs

Bone marrow was harvested postmortem from the iliac crest of three adult sheep, aged 2–4 years, which were euthanized for reasons unrelated to this study. The sheep were placed in sternal recumbency. After surgical skin preparation, the iliac crest was punctured using a Jamshidi biopsy needle (13G; Greiner BioOne). Bone marrow (17.5 mL) was aspirated carefully with a preheparinized syringe (2.5 mL heparin in 20 mL syringe) and taken to the laboratory for further processing. To isolate the mononuclear cell fraction, a density gradient filtration was performed using Ficoll as described above. MSCs were cultured, frozen, and stored in liquid nitrogen until further use at the lowest possible passage (P1–2) as described above.

Cell proliferation

Population doubling rates per day and generation times were evaluated for cotyledon-derived MSC (cotMSCs, n = 5) and bmMSCs (n = 3) samples. For that purpose, cells were seeded at a defined density (3000 cells/cm²). Care was taken that the proliferation assay was carried out in the same passage (P1) for cotMSCs and bmMSCs. On 80–90% confluence, cells were trypsinized and counted using an automatic cell counter (Countess II FL; Invitrogen). Population doubling rates (PD) and generation times (GT) were calculated according to the following formulas^37,90:

PD: (cell count harvest/cell count seeding)/ln (2)

PD/day: PD/day in culture

GT: 1/(PD/day in culture)

For statistical analysis, a t-test for two groups with different variances was performed using IBM SPSS Statistics 24.

Migration assay

To confirm their migration potential, a scratch migration assay was performed for cotMSCs (n = 5) and bmMSCs (n = 3). On a six-well plate, 3 × 10⁵ cells/well were seeded in triplicate and cultured until subconfluence. Care was taken that the cotMSCs and bmMSCs used were in the same passage (P3). On subconfluence, a scratch was created using a pipette tip (20–200 μL). Pictures of the scratch area were taken every 6 h for 48 h using an automatic cell imaging system (EVOS^® Auto FL Imaging System; Thermo Fisher Scientific). Based on the pictures, the time until complete closure of the scratch was determined and the migration capability of cotMSC and bmMSC samples was confirmed.

MSC differentiation

To prove their trilineage differentiation potential, cotMSCs (n = 5) and bmMSCs (n = 3) were differentiated into the chondrogenic, adipogenic, and osteogenic lineage. Care was taken that the cotMSCs and bmMSCs used were in the same passage (P3). cotMSCs cultured for the same time period but in standard culture medium (described above) served as control.

Chondrogenic differentiation

For chondrogenic differentiation, 350,000 cells were suspended in 3 mL chondrogenic differentiation medium (StemPro Chondrogenesis Differentiation Kit; Gibco) and centrifuged for 5 min at 280 g, 4°C to form a pellet. The cell pellets were cultured in duplicate in 15-mL Falcon tubes and were daily shaken carefully to prevent attachment to the plastic tube. Medium was changed twice weekly. After 3 weeks of cultivation in differentiation medium, the pellets were fixed with buffered 4% formalin (ACM, Herba Chemosan Apotheker AG) and subjected to paraffin embedding and sectioning. The sections were stained with Alcian blue (Chroma, Waldeck GmbH) according to a standard staining protocol⁸⁹ to confirm the production of acidic mucosubstances.

Osteogenic differentiation

Osteogenic differentiation was performed in monolayer using the osteogenic differentiation medium (StemPro Osteogenesis Differentiation Kit; Gibco). First, 1000 cells/cm² were seeded on 24-well plates in standard culture medium. After 3 days, the medium was replaced by osteogenic differentiation medium. Medium was changed twice weekly. After 3 weeks, the cells formed three-dimensional structures (cell sheets or nodules), which were fixed with buffered 4% formalin, harvested, paraffin embedded, and sectioned. To demonstrate extracellular calcium deposition, as sign for osteogenic differentiation, von Kossa staining (silver nitrate, Merck; Natriumthiosulfat, Sigma Aldrich) was performed.⁸⁹

Adipogenic differentiation

Adipogenic differentiation was performed in monolayer culture. After cell seeding (1000 cells/cm²) in standard culture medium and culturing for 3 days, adipogenic differentiation was induced using adipogenic differentiation medium (StemPro Adipogenesis Differentiation Kit; Gibco). Medium was changed twice a week. After 3 weeks, the cells were fixed with buffered 4% formalin and stained for intracellular lipid vacuoles with Oil red O (Sigma-Aldrich) according to a standard staining protocol.⁸⁹

Flow cytometry analysis

Cell surface antigen expression of cotMSCs (n = 5) and bmMSCs (n = 3) was analyzed using positive and negative surface markers to define their stem cell character (Table 1). The markers have been selected according to Boos et al.⁹¹ For flow cytometry, cells at P2 were detached from the flask using trypsin. MSCs were washed with FACS buffer containing PBS, 3% FCS, and 0.1% NaN₃ (Sigma Aldrich) and pelleted by centrifugation for 5 min at 400 g. Subsequently, 1 × 10⁵ cells per sample were incubated with a specific primary antibody at 4°C. After 30 min, unbound antibody was removed by washing with 1 mL FACS buffer. The cell pellets were resuspended in 200 μL FACS buffer and analyzed with a BD FACSCanto II (BD Biosciences). Per sample, 1 × 10⁴ events were recorded. Unstained and isotype controls were performed. The results were analyzed by FlowJo (10.0.7) software.

Table 1.

Monoclonal Antibodies Used to Characterize Cell Surface Marker Expression by Flow Cytometry

Antibody	Clone	Concentration (v/v)	Source	Isotype	Clone	Source
CD29	MAR4	1/10	PD Pharmingen	IgG1_K	MOPC-21	BD Pharmingen
CD31	CO.3E1D4	1/50	BioRad AbD Serotec	IgG2_a	G155–178	BD Pharmingen
CD44	25.32	1/200	BioRad AbD Serotec	IgG1	MOPC-21	BD Pharmingen
CD45	01.11.1932	1/50	BioRad AbD Serotec	IgG1	MOPC-21	BD Pharmingen
CD166	3A6	1/10	BD Pharmingen	IgG1_k	MOPC-21	BD Pharmingen

Results

Collection of cotyledons

Collection of the afterbirth was easy and not associated with any harm to the ewe or the newborn lamb. After mincing and digesting the cotyledons, a viscous bloody solution was obtained. Despite the tissue harvest in a farm environment, no bacterial or mycotic contamination of the cells was noticed in culture.

MSC isolation and culture

MSCs could be reliably isolated from all cotyledon (n = 5) and bone marrow (n = 3) samples with a reproducible cell yield (Table 2). MSCs from both cell sources fulfilled the minimal criteria of MSCs defined by the International Society for Cellular Therapy (spindle-shaped morphology, adherence to plastic, and colony formation).⁹² The morphology of cotMSCs at different stages of colony confluence in P0 is presented in Figure 2. It was similar to the widely and well-known morphology of the “gold standard” bmMSCs.

FIG. 2.

CotMSCs in P0 showing spindle-shaped morphology, adherence to plastic, and colony formation at different stages of colony confluence. Scale bar 100 μm. cotMSCs, cotyledon-derived mesenchymal stem cells.

Table 2.

MSC Isolation Efficiency: Cell Yield per Donor at First Passage

Cell type	Raw material	Cell count at P1	Days in culture until P1 (days)
cotMSC1	10 Cotyledons	1,030,000	6
cotMSC2	11 Cotyledons	1,530,000	7
cotMSC3	11 Cotyledons	1,400,000	5
cotMSC4	9 Cotyledons	1,070,000	8
cotMSC5	7 Cotyledons	677,000	7
bmMSC1	40 mL bone marrow	2,180,000	10
bmMSC2	50 mL bone marrow	2,530,000	9
bmMSC3	40 mL bone marrow	3,700,000	9

bmMSCs, bone marrow-derived MSCs; cotMSCs, cotyledon-derived MSCs.

Cell proliferation

Cell proliferation was evaluated for cotMSCs and bmMSCs (Fig. 3). The mean population doubling rate per day of bmMSCs was 1.672. The individual values were 0.920, 1.789, and 2.308. The mean cell doubling rate of the cotMSCs was 0.729. The individual values were 0.294, 0.635, 0.877, 0.879, and 0.960) (Fig. 3A, B). Accordingly, the generation times for bmMSCs (mean: 0.693 days, individual values: 0.433, 0.559, and 1.086) were shorter than for cotMSCs (mean: 1.659 days, individual values: 1.041, 1.137, 1.140, 1.575, and 3.405). Due to the low sample number, the proliferation differences between cotMSCs and bmMSCs did not reach statistical significance (population doubling per day: p = 0.135; generation time: p = 0.101).

FIG. 3.

Cell proliferation. Population doubling rates per day of bmMSCs and cotMSCs (A). Mean population doubling rates per day of bmMSCs and cotMSCs (B). Due to the low sample number, the proliferation differences between cotMSCs and bmMSCs did not reach statistical significance (population doubling per day: p = 0.135). bmMSCs, bone marrow-derived mesenchymal stem cells.

Migration assay

After seeding cotMSCs and bmMSCs for the migration assay, they reached confluence after 3 days and a scratch was performed (Fig. 4A, B). The scratch was photographed every 6 h to show cell migration into the cell-free area. Although bmMSCs showed subjectively faster migration at all time points (Fig. 4C–F), after 48 h, both cell types had closed the gap confirming their migration potential (Fig. 4G, H).

FIG. 4.

Migration assay. To assess migration capacity of cotMSCs (A, C, E, G) and bone marrow-derived MSCs (B, D, F, H), a scratch was created in a cell monolayer using a pipette tip (A, B). Example pictures after 12 h (C, D) and 24 h (E, F) are presented. After 48 h (G, H), both cell types had closed the gap. Scale bar = 100 μm. MSC, mesenchymal stem cell.

MSC differentiation

Both cotMSCs (Fig. 5, line 1–5) and adult bmMSCs (Fig. 5, line 6) showed chondrogenic differentiation potential with a subjectively higher chondrogenic potential inherent to cotMSCs (Fig. 5, row 1, line 1–5) compared to the bmMSCs (Fig. 5, row 1, line 6). After 3 weeks in culture, the cotMSCs showed subjectively more production of extracellular acidic polysaccharides and mucopolysaccharides than bmMSCs. Also, lacunae formation around the cells, which is indicative for cartilage tissue, appeared more distinct for the cotMSCs than the bmMSCs. The control samples showed no indication of chondrogenic differentiation.

FIG. 5.

MSC differentiation. Chondrogenic (row 1), osteogenic (row 2), and adipogenic (row 3) differentiation of cotMSC (line 1–5) and bone marrow-derived MSC (line 6). Acidic mucosubstances produced during chondrogenic differentiation were demonstrated as extracellular blue substrate with Alcian blue staining. After osteogenic differentiation, von Kossa staining displayed black extracellular calcium deposition. Oil red O-positive droplets inside the cells confirmed adipogenic differentiation of bone marrow-derived MSCs (row 3, line 6), whereas cotMSCs showed no positive staining (row 3, line 1–5) indicating a lack of adipogenic differentiation potential. cotMSCs cultured for the same time period in standard culture medium served as control (line7). Scale bar = 100 μm (row 1 and 2) and 50 μm (row 3), respectively.

Osteogenic differentiation was demonstrated for both cell sources. Extracellular calcium deposition was confirmed by van Kossa staining (Fig. 5, row 2). After 3 weeks in adipogenic differentiation medium, cotMSCs showed no positive staining (Fig. 5, row 3, line 1–5) indicating a lack of adipogenic differentiation potential. In contrast, bmMSCs showed obvious and numerous intracellular lipid vacuoles that stained intensively for Oil red O (Fig. 5, row 3, line 6). cotMSCs cultured for the same time period in standard culture medium served as control (Fig. 5, line7).

Flow cytometry analysis

FACS analysis confirmed the expression of MSC markers on the protein level. In accordance to Boos et al.,⁹¹ cotMSCs and bmMSCs were positive for the surface markers CD29, CD44, and CD166 and negative for CD31 and CD45, which are typical hematopoietic cell markers. Isotype controls (IgG1 and IgG2a) were negative. FACS histograms are depicted in Figure 6.

FIG. 6.

FACS analysis of cotMSCs and bmMSCs. Both MSC types were positive for the surface markers CD29, CD44, and CD166 and negative for the hematopoietic markers CD31 and CD45. Black = histograms primary antibodies, gray = histograms controls. Percentages of live, positively stained cells in relation to controls are given in the right upper corner of each histogram.

Discussion

Sheep are a frequently used animal model in regenerative medicine. However, truly noninvasive sources of MSCs in sheep are scarce. Fetal/perinatal tissues are typical noninvasive sources of MSCs in many species.^{13,15,18,19,36–69} However, in sheep, most studies using perinatal tissues such as cord blood, amniotic membranes, or amniotic fluid obtained from these tissues preterm using surgical harvesting procedures.^{13,18,19,36,41,43,59} In the current study, ovine placental cotyledons obtained on full-term delivery were tested for their suitability as noninvasive MSC source.

Semisterile harvesting of the ovine afterbirth was easy to perform. Although perinatal tissues bear a high risk of bacterial or mycotic contamination^44,61 due to the nonsterile birth canal and the delivery in a farm environment, which make the use of antibiotic in culture mandatory,⁶¹ no problems with either bacterial or mycotic contamination were encountered. In this study, 1% iodine solution was used as disinfectant to clean the obtained afterbirth from potential bacterial or mycotic contamination immediately following harvest. In addition, the cotyledons were kept in a PBS solution containing antibiotics until further processing. The obtained MSCs performed well in culture and during differentiation and showed no negative effects of the initial cleaning step or the antibiotics. Whether this cell source would also be feasible for harvest and culture without the addition of disinfectants and antibiotics requires further investigation.

During processing of the cotyledons, residual blood, which was contained in the blood vessels of the chorionic villi, resulted in a blood-like appearance of the solution obtained after mincing and digesting the villi. This finding supports the theory that the isolated MSCs do not solely originate from the perivascular niche in the placental chorionic villi (Fig. 1E) as reported for humans⁷⁷ but also from the residual fetal blood contained in the blood vessels of the chorionic villi (Fig. 1F). The fetal part of the ovine placenta (the cotyledon) and the maternal part disconnect without injury to the maternal tissue. Due to this nonbloody separation, the blood contained in the cotyledon is exclusively of fetal origin. Therefore, it stands to reason that the isolated MSCs derive from the perivascular niche in the chorionic villi and the residual fetal blood. For a more detailed characterization of the exact origin of cotMSCs, further studies will be necessary. With the isolation technique described in this article, fibroblasts are likely contained in the early passage cell population in addition to the desired MSCs. However, it was previously shown that fibroblast contamination did not adversely influence MSCs but even led to better differentiation potential than shown for fibroblast-depleted MSC populations.⁹³

In the current study, we demonstrated that cotMSCs have good osteogenic and chondrogenic but very limited to no adipogenic differentiation potential. It is well known that MSCs have different degrees of phenotypic plasticity depending on the tissue source.^37,72,94,95 Despite similar morphology and immunophenotypic signature during expansion and growth, the differentiation potential toward the adipogenic lineage can be greatly influenced by the tissue of origin and the developmental stage (adult or fetal).⁹⁶ Adipogenic differentiation potential relies on the activation of the peroxisome proliferator–activated receptor γ (PPAR-γ) pathway, which is reduced or delayed in fetal MSCs, resulting in strong impairment of adipogenesis.⁹⁶ Since the cotMSCs used in the current study were obtained from perinatal tissue of fetal origin, the reduced PPAR-γ pathway may be a reason for the lack of adipogenic differentiation observed in these MSCs. This finding is also in accordance to other studies, which reported a low adipogenic differentiation potential for equine cord blood-derived MSCs.^{63,94,95,97,98} Age-related differences in adipogenic differentiation potential were also reported for muscle stem cells. It was shown that muscle stem cells have a greater propensity to adipogenic differentiation with advanced age.⁹⁹

Despite the reduced adipogenic differentiation, the cotMSCs met all minimum criteria required for the characterization of MSCs.⁹² They were spindle shaped, showed formation of colonies in culture, adherence to plastic, and expression or lack of expression of specific surface antigens detected by FACS (positive for CD29, CD33, and CD166, negative for CD31 and CD45). The MSC yield was reliable as MSCs were obtained from all five afterbirths included into the study. CotMSCs and bmMSCs showed good migration potential confirmed by scratch assays. Although cell proliferation and migration of cotMSCs seemed overall slower than for adult bmMSCs, which is in contrast to findings of other authors who reported a higher proliferation rate of juvenile compared to adult MSCs.^70,71

However, after further fine-tuning of the harvesting, isolation, and culture conditions, some of the shortcomings of the cotMSCs in comparison to bmMSCs may be overcome. For instance, the cotMSC yield may be raised considerably as the average ovine afterbirth comprises about 100 cotyledons^87,88 and in the current study only 7–11 per donor were processed. Due to the noninvasive harvesting modalities, sheep cotyledons may lend themselves as a valuable source of MSCs to consider in the light of the principles of the 3Rs (reduce, refine, replace) for studies performed on these frequently used model animals.

In summary, in this study, MSCs have been successfully harvested from ovine placenta cotyledons. They exhibit morphological, immunophenotypical, and multipotential characteristics similar to those observed in bone marrow-derived ovine or human MSCs.^3,92 Ovine cotyledons were shown to be a reliable, abundant source for the truly noninvasive, pain- and risk-free harvest of MSCs. They can be harvested easily and without a bigger risk of bacterial or mycotic contamination than any other perinatal tissue obtained on full-term delivery. The collection procedure does not interfere with partum or the initial bonding phase between ewe and lamb and is therefore exempt from ethical debate. The isolated MSCs can be cryopreserved for later use in in vitro studies or for autologous as well as allogeneic in vivo applications. Further characterization of sheep cotMSCs may be beneficial for translational studies using MSCs for particular therapeutic applications. For example, gene expression analysis of the master regulators of differentiation (Sox9, Runx2, PPARγ) and of differentiation markers during chondro-, osteo-, and adipogenesis would be interesting, to investigate if a different basal expression of the transcription factors can explain the differences in the differentiation potential.

Footnotes

Acknowledgments

We acknowledge Manuela Miksic, Brigitte Machac, Eva Haltmayer, Anna Bauer, Astrid Kohl, Michaela Pinisch, Stephanie Schmidt, Öczan Korkmaz, and the Management and Staff of the Vetmeduni Vienna Teaching and Research farm Kremesberg for their technical support.

Disclosure Statement

No competing financial interests exist.

References

Herberts

C.A.

, Kwa

M.S.

, and Hermsen

H.P.

Risk factors in the development of stem cell therapy. J Transl Med, 9, 29, 2011.

Prabhakar

Translational research challenges: finding the right animal models. J Investig Med, 60, 1141, 2012.

McCarty

R.C.

, Gronthos

, Zannettino

A.C.

, Foster

B.K.

, and Xian

C.J.

Characterisation and developmental potential of ovine bone marrow derived mesenchymal stem cells. J Cell Physiol, 219, 324, 2009.

Martini

, Fini

, Giavaresi

, and Giardino

Sheep model in orthopedic research: a literature review. Comp Med, 51, 292, 2001.

Entrican

, Wattegedera

S.R.

, and Griffiths

D.J.

Exploiting ovine immunology to improve the relevance of biomedical models. Mol Immunol, 66, 68, 2015.

Nuss

K.M.

, Auer

J.A.

, Boos

, and von Rechenberg

An animal model in sheep for biocompatibility testing of biomaterials in cancellous bones. BMC Musculoskelet Disord, 7, 67, 2006.

Zarrinkalam

M.R.

, Beard

, Schultz

C.G.

, and Moore

R.J.

Validation of the sheep as a large animal model for the study of vertebral osteoporosis. Eur Spine J, 18, 244, 2009.

Porada

G.A.

, Porada

, and Zanjani

E.D.

The fetal sheep: a unique model system for assessing the full differentiative potential of human stem cells. Yonsei Med J, 45 Suppl, 7, 2004.

Welin

A.K.

, Svedin

, Lapatto

, Sultan

, Hagberg

, Gressens

, Kjellmer

, and Mallard

Melatonin reduces inflammation and cell death in white matter in the mid-gestation fetal sheep following umbilical cord occlusion. Pediatr Res, 61, 153, 2007.

10.

Emmert

M.Y.

, Weber

, Wolint

, Frauenfelder

, Zeisberger

S.M.

, Behr

, Sammut

, Scherman

, Brokopp

C.E.

, Schwartlander

, Vogel

, Vogt

, Grunenfelder

, Alkadhi

, Falk

, Boss

, and Hoerstrup

S.P.

Intramyocardial transplantation and tracking of human mesenchymal stem cells in a novel intra-uterine pre-immune fetal sheep myocardial infarction model: a proof of concept study. PLoS One, 8, e57759, 2013.

11.

Airey

J.A.

, Almeida-Porada

, Colletti

E.J.

, Porada

C.D.

, Chamberlain

, Movsesian

, Sutko

J.L.

, and Zanjani

E.D.

Human mesenchymal stem cells form Purkinje fibers in fetal sheep heart. Circulation, 109, 1401, 2004.

12.

Porada

C.D.

, Park

, Almeida-Porada

, and Zanjani

E.D.

The sheep model of in utero gene therapy. Fetal Diagn Ther, 19, 23, 2004.

13.

Barboni

, Russo

, Curini

, Mauro

, Martelli

, Muttini

, Bernabo

, Valbonetti

, Marchisio

, Di Giacinto

, Berardinelli

, and Mattioli

Achilles tendon regeneration can be improved by amniotic epithelial cell allotransplantation. Cell Transplant, 21, 2377, 2012.

14.

Barboni

, Mangano

, Valbonetti

, Marruchella

, Berardinelli

, Martelli

, Muttini

, Mauro

, Bedini

, Turriani

, Pecci

, Nardinocchi

, Zizzari

V.L.

, Tete

, Piattelli

, and Mattioli

Synthetic bone substitute engineered with amniotic epithelial cells enhances bone regeneration after maxillary sinus augmentation. PLoS One, 8, e63256, 2013.

15.

Caminal

, Fonseca

, Peris

, Moll

, Rabanal

R.M.

, Barrachina

, Codina

, Garcia

, Cairo

J.J.

, Goodia

, Pla

, and Vives

Use of a chronic model of articular cartilage and meniscal injury for the assessment of long-term effects after autologous mesenchymal stromal cell treatment in sheep. N Biotechnol, 31, 492, 2014.

16.

Martinello

, Bronzini

, Perazzi

, Testoni

, De Benedictis

G.M.

, Negro

, Caporale

, Mascarello

, Iacopetti

, and Patruno

Effects of in vivo applications of peripheral blood-derived mesenchymal stromal cells (PB-MSCs) and platlet-rich plasma (PRP) on experimentally injured deep digital flexor tendons of sheep. J Orthop Res, 31, 306, 2013.

17.

Muttini

, Valbonetti

, Abate

, Colosimo

, Curini

, Mauro

, Berardinelli

, Russo

, Cocciolone

, Marchisio

, Mattioli

, Tosi

, Vulpiani

M.P.

, and Barboni

Ovine amniotic epithelial cells: in vitro characterization and transplantation into equine superficial digital flexor tendon spontaneous defects. Res Vet Sci, 94, 158, 2013.

18.

Reflection paper on in-vitro cultured chondrocyte containing products for cartilage repair of the knee.. European Medicines Agency, EMA/CAT/CPWP/568181/2009, 2010. www.ema.europa.eu/docs/en_GB/document_library/Scientific_guideline/2010/05/WC500090887.pdf (Accessed February 2, 2017).

19.

Guidance for Industry. Preparation of IDEs and INDs for products intended to repair or replace knee cartilage. U.S. Department of Health and Human Services Food and Drug Administration Center for Biologics Evaluation and Research Center for Devices and Radiological Health, December 2011. www.fda.gov/downloads/Biologics BloodVaccines/GuidanceCompliance RegulatoryInformation/Guidances/CellularandGeneTherapy/UCM288011.pdf (Accessed February 2, 2017).

20.

Delling

, Lindner

, Ribitsch

, Julke

, and Brehm

Comparison of bone marrow aspiration at the sternum and the tuber coxae in middle-aged horses. Can J Vet Res, 76, 52, 2012.

21.

Heidari

, Shirazi

, Akhondi

M.M.

, Hassanpour

, Behzadi

, Naderi

M.M.

, Sarvari

, and Borjian

Comparison of proliferative and multilineage differentiation potential of sheep mesenchymal stem cells derived from bone marrow, liver, and adipose tissue. Avicenna J Med Biotechnol, 5, 104, 2013.

22.

Kasashima

, Ueno

, Tomita

, Goodship

A.E.

, and Smith

R.K.W.

Optimisation of bone marrow aspiration from the equine sternum for the safe recovery of mesenchymal stem cells. Equine Vet J, 43, 288, 2011.

23.

Kassem

, and Abdallah

B.M.

Human bone-marrow-derived mesenchymal stem cells: biological characteristics and potential role in therapy of degenerative diseases. Cell Tissue Res, 331, 157, 2008.

24.

Prockop

D.J.

Marrow stromal cells as stem cells for continual renewal of nonhematopoietic tissues and as potential vectors for gene therapy. J Cell Biochem, 30–31, 284, 1998.

25.

Rentsch

, Hess

, Rentsch

, Hofmann

, Manthey

, Scharnweber

, Biewener

, and Zwipp

Ovine bone marrow mesenchymal stem cells: isolation and characterization of the cells and their osteogenic differentiation potential on embroidered and surface-modified polycaprolactone-co-lactide scaffolds. In Vitro Cell Dev Biol Anim, 46, 624, 2010.

26.

Yoshimura

, Muneta

, Nimura

, Yokoyama

, Koga

, and Sekiya

Comparison of rat mesenchymal stem cells derived from bone marrow, synovium, periosteum, adipose tissue, and muscle. Cell Tissue Res, 327, 449, 2007.

27.

Barberini

D.J.

, Freitas

N.P.P.

, Magnoni

M.S.

, Maia

, Listoni

A.J.

, Heckler

M.C.

, Sudano

M.J.

, Golim

M.A.

, Landim-Alvarenga

F.D.

, and Amorim

R.M.

Equine mesenchymal stem cells from bone marrow, adipose tissue and umbilical cord: immunophenotypic characterization and differentiation potential. Stem Cell Res Ther, 5, 25, 2014.

28.

Grzesiak

, Krzysztof

, Karol

, and Joanna

Isolation and morphological characterisation of ovine adipose-derived mesenchymal stem cells in culture. Int J Stem Cells, 4, 99, 2011.

29.

Ranera

, Ordovas

, Lyahyai

, Bernal

M.L.

, Fernandes

, Remacha

A.R.

, Romero

, Vazquez

F.J.

, Osta

, Cons

, Varona

, Zaragoza

, Martin-Burriel

, and Rodellar

Comparative study of equine bone marrow and adipose tissue-derived mesenchymal stromal cells. Equine Vet J, 44, 33, 2012.

30.

Vidal

M.A.

, Kilroy

G.E.

, Lopez

M.J.

, Johnson

J.R.

, Moore

R.M.

, and Gimble

J.M.

Characterization of equine adipose tissue-derived stromal cells: adipogenic and osteogenic capacity and comparison with bone marrow-derived mesenchymal stromal cells. Vet Surg, 36, 613, 2007.

31.

Zuk

P.A.

, Zhu

, Mizuno

, Huang

, Futrell

J.W.

, Katz

A.J.

, Benhaim

, Lorenz

H.P.

, and Hedrick

M.H.

Multilineage cells from human adipose tissue: implications for cell-based therapies. Tissue Eng, 7, 211, 2001.

32.

Giovannini

, Brehm

, Mainil-Varlet

, and Nesic

Multilineage differentiation potential of equine blood-derived fibroblast-like cells. Differentiation, 76, 118, 2008.

33.

Koerner

, Nesic

, Romero

J.D.

, Brehm

, Mainil-Varlet

, and Grogan

S.P.

Equine peripheral blood-derived progenitors in comparison to bone marrow-derived mesenchymal stem cells. Stem Cells, 24, 1613, 2006.

34.

Wang

, and Moutsoglou

Osteogenic and adipogenic differentiation potential of an immortalized fibroblast-like cell line derived from porcine peripheral blood. In Vitro Cell Dev Biol Anim, 45, 584, 2009.

35.

Lyahyai

, Mediano

D.R.

, Ranera

, Sanz

, Remacha

A.R.

, Bolea

, Zaragoza

, Rodellar

, and Martin-Burriel

Isolation and characterization of ovine mesenchymal stem cells derived from peripheral blood. BMC Vet Res, 8, 169, 2012.

36.

Barboni

, Curini

, Russo

, Mauro

, Di Giacinto

, Marchisio

, Alfonsi

, and Mattioli

Indirect co-culture with tendons or tenocytes can program amniotic epithelial cells towards stepwise tenogenic differentiation. PLos One, 7, e30974, 2012.

37.

Burk

, Ribitsch

, Gittel

, Juelke

, Kasper

, Staszyk

, and Brehm

Growth and differentiation characteristics of equine mesenchymal stromal cells derived from different sources. Vet J, 195, 98, 2013.

38.

Coli

, Nocchi

, Lamanna

, Iorio

, Lapi

, Urciuoli

, Scatena

, Giannessi

, Stornelli

M.R.

, and Passeri

Isolation and characterization of equine amnion mesenchymal stem cells. Cell Biol Int Rep (2010), 18, e00011, 2011.

39.

De Coppi

, Bartsch

, Siddiqui

M.M.

, Xu

, Santos

C.C.

, Perin

, Mostoslavsky

, Serre

A.C.

, Snyder

E.Y.

, Yoo

J.J.

, Furth

M.E.

, Soker

, and Atala

Isolation of amniotic stem cell lines with potential for therapy. Nat Biotechnol, 25, 100, 2007.

40.

De Schauwer

, Goossens

, Piepers

, Hoogewijs

M.K.

, Govaere

J.L.

, Smits

, Meyer

, Van Soom

, and Van de Walle

G.R.

Characterization and profiling of immunomodulatory genes of equine mesenchymal stromal cells from non-invasive sources. Stem Cell Res Ther, 5, 6, 2014.

41.

Di Tomo

, Pipino

, Lanuti

, Morabito

, Pierdomenico

, Sirolli

, Bonomini

, Miscia

, Mariggio

M.A.

, Marchisio

, Barboni

, and Pandolfi

Calcium sensing receptor expression in ovine amniotic fluid mesenchymal stem cells and the potential role of R-568 during osteogenic differentiation. PLoS One, 8, e73816, 2013.

42.

Erices

, Conget

, and Minguell

J.J.

Mesenchymal progenitor cells in human umbilical cord blood. Br J Haematol, 109, 235, 2000.

43.

Fuchs

J.R.

, Hannouche

, Terada

, Zand

, Vacanti

J.P.

, and Fauza

D.O.

Cartilage engineering from ovine umbilical cord blood mesenchymal progenitor cells. Stem Cells, 23, 958, 2005.

44.

Gulati

B.R.

, Kumar

, Mohanty

, Kumar

, Somasundaram

R.K.

, and Yadav

P.S.

Bone morphogenetic protein-12 induces tenogenic differentiation of mesenchymal stem cells derived from equine amniotic fluid. Cells Tissues Organs, 198, 377, 2013.

45.

Hoynowski

S.M.

, Fry

M.M.

, Gardner

B.M.

, Leming

M.T.

, Tucker

J.R.

, Black

, Sand

, and Mitchell

K.E.

Characterization and differentiation of equine umbilical cord-derived matrix cells. Biochem Biophys Res Commun, 362, 347, 2007.

46.

Iacono

, Brunori

, Pirrone

, Pagliaro

P.P.

, Ricci

, Tazzari

P.L.

, and Merlo

Isolation, characterization and differentiation of mesenchymal stem cells from amniotic fluid, umbilical cord blood and Wharton's jelly in the horse. Reproduction, 143, 455, 2012.

47.

In ’t Anker

P.S.

, Scherjon

S.A.

, Kleijburg-van der Keur

, de Groot-Swings

G.M.

, Claas

F.H.

, Fibbe

W.E.

, and Kanhai

H.H.

Isolation of mesenchymal stem cells of fetal or maternal origin from human placenta. Stem Cells, 22, 1338, 2004.

48.

Kim

, Lee

, Kim

, Hwang

K.J.

, Kwon

H.C.

, Kim

S.K.

, Cho

D.J.

, Kang

S.G.

, and You

Human amniotic fluid-derived stem cells have characteristics of multipotent stem cells. Cell Prolif, 40, 75, 2007.

49.

Koch

T.G.

, Heerkens

, Thomsen

P.D.

, and Betts

D.H.

Isolation of mesenchymal stem cells from equine umbilical cord blood. BMC Biotechnol, 7, 26, 2007.

50.

Koike

, Zhou

, Takeda

, Fathy

, Okabe

, Yoshida

, Nakamura

, Kato

, and Nikaido

Characterization of amniotic stem cells. Cell Reprogram, 16, 298, 2014.

51.

Kunisaki

S.M.

, Jennings

R.W.

, and Fauza

D.O.

Fetal cartilage engineering from amniotic mesenchymal progenitor cells. Stem Cells Dev, 15, 245, 2006.

52.

Kunisaki

S.M.

, Fuchs

J.R.

, Steigman

S.A.

, and Fauza

D.O.

A comparative analysis of cartilage engineered from different perinatal mesenchymal progenitor cells. Tissue Eng, 13, 2633, 2007.

53.

Lange-Consiglio

, Corradetti

, Meucci

, Perego

, Bizzaro

, and Cremonesi

Characteristics of equine mesenchymal stem cells derived from amnion and bone marrow: in vitro proliferative and multilineage potential assessment. Equine Vet J, 45, 737, 2013.

54.

Lange-Consiglio

, Rossi

, Tassan

, Perego

, Cremonesi

, and Parolini

Conditioned medium from horse amniotic membrane-derived multipotent progenitor cells: immunomodulatory activity in vitro and first clinical application in tendon and ligament injuries in vivo. Stem Cells Dev, 22, 3015, 2013.

55.

Lange-Consiglio

, Tassan

, Corradetti

, Meucci

, Perego

, Bizzaro

, and Cremonesi

Investigating the efficacy of amnion-derived compared with bone marrow-derived mesenchymal stromal cells in equine tendon and ligament injuries. Cytotherapy, 15, 1011, 2013.

56.

Lovati

A.B.

, Corradetti

, Lange Consiglio

, Recordati

, Bonacina

, Bizzaro

, and Cremonesi

Comparison of equine bone marrow-, umbilical cord matrix and amniotic fluid-derived progenitor cells. Vet Res Commun, 35, 103, 2011.

57.

Marcus

A.J.

, and Woodbury

Fetal stem cells from extra-embryonic tissues: do not discard. J Cell Mol Med, 12, 730, 2008.

58.

Martino

N.A.

, Reshkin

S.J.

, Ciani

, and Dell'Aquila

M.E.

Calcium-sensing receptor-mediated osteogenic and early-stage neurogenic differentiation in umbilical cord matrix mesenchymal stem cells from a large animal model. PLoS One, 9, e111533, 2014.

59.

Mauro

, Turriani

, Ioannoni

, Russo

, Martelli

, Di Giacinto

, Nardinocchi

, and Berardinelli

Isolation, characterization, and in vitro differentiation of ovine amniotic stem cells. Vet Res Commun, 34 Suppl 1, S25, 2010.

60.

Montemurro

, Andriolo

, Montelatici

, Weissmann

, Crisan

, Colnaghi

M.R.

, Rebulla

, Mosca

, Peault

, and Lazzari

Differentiation and migration properties of human foetal umbilical cord perivascular cells: potential for lung repair. J Cell Mol Med, 15, 796, 2011.

61.

Passeri

, Nocchi

, Lamanna

, Lapi

, Miragliotta

, Giannessi

, Abramo

, Stornelli

M.R.

, Matarazzo

, Plenteda

, Urciuoli

, Scatena

, and Coli

Isolation and expansion of equine umbilical cord-derived matrix cells (EUCMCs). Cell Biol Int, 33, 100, 2009.

62.

Perin

, Giuliani

, Jin

, Sedrakyan

, Carraro

, Habibian

, Warburton

, Atala

, and De Filippo

R.E.

Renal differentiation of amniotic fluid stem cells. Cell Prolif, 40, 936, 2007.

63.

Reed

S.A.

, and Johnson

S.E.

Equine umbilical cord blood contains a population of stem cells that express Oct4 and differentiate into mesodermal and endodermal cell types. J Cell Physiol, 215, 329, 2008.

64.

Sarugaser

, Lickorish

, Baksh

, Hosseini

M.M.

, and Davies

J.E.

Human umbilical cord perivascular (HUCPV) cells: a source of mesenchymal progenitors. Stem Cells, 23, 220, 2005.

65.

Secco

, Zucconi

, Vieira

N.M.

, Fogaca

L.L.Q.

, Cerqueira

, Carvalho

M.D.F.

, Jazedje

, Okamoto

O.K.

, Muotri

A.R.

, and Zatz

Mesenchymal stem cells from umbilical cord: do not discard the cord!. Neuromuscular Disord, 18, 17, 2008.

66.

Seshareddy

, Troyer

, and Weiss

M.L.

Method to isolate mesenchymal-like cells from wharton's jelly of umbilical cord. Methods Cell Biol, 86, 101, 2008.

67.

Shaw

S.W.S.

, Bollini

, Nader

K.A.

, Gastadello

, Mehta

, Filppi

, Cananzi

, Gaspar

H.B.

, Qasim

, De Coppi

, and David

A.L.

Autologous transplantation of amniotic fluid-derived mesenchymal stem cells into sheep fetuses. Cell Transplant, 20, 1015, 2011.

68.

Schuh

E.M.

, Friedman

M.S.

, Carrade

D.D.

, Li

, Heeke

, Oyserman

S.M.

, Galuppo

L.D.

, Lara

D.J.

, Walker

N.J.

, Ferraro

G.L.

, Owens

S.D.

, and Borjesson

D.L.

Identification of variables that optimize isolation and culture of multipotent mesenchymal stem cells from equine umbilical-cord blood. Am J Vet Res, 70, 1526, 2009.

69.

Tsai

M.S.

, Lee

J.L.

, Chang

Y.J.

, and Hwang

S.M.

Isolation of human multipotent mesenchymal stem cells from second-trimester amniotic fluid using a novel two-stage culture protocol. Hum Reprod, 19, 1450, 2004.

70.

Hass

, Kasper

, Bohm

, and Jacobs

Different populations and sources of human mesenchymal stem cells (MSC): a comparison of adult and neonatal tissue-derived MSC. Cell Commun Signal, 9, 12, 2011.

71.

Rogers

, and Casper

R.F.

Umbilical cord blood stem cells. Best Pract Res Clin Obstet Gynaecol, 18, 893, 2004.

72.

Zhang

, Khan

, Delling

, and Tobiasch

Mechanisms underlying the osteo- and adipo-differentiation of human mesenchymal stem cells. ScientificWorldJournal, 2012, 793823, 2012.

73.

Bartholomew

, Owens

S.D.

, Ferraro

G.L.

, Carrade

D.D.

, Lara

D.J.

, Librach

F.A.

, Borjesson

D.L.

, and Galuppo

L.D.

Collection of equine cord blood and placental tissues in 40 thoroughbred mares. Equine Vet J, 41, 724, 2009.

74.

Valle

, Screnci

, Murgi

, Capozzi

, and Girelli

Collection of umbilical cord blood for banking: collection rate and factors influencing collection. Blood Transfus, 2016 [Epub ahead of print]; DOI: 10.2450/2016.0262-16.

75.

Moretti

, Hatlapatka

, Marten

, Lavrentieva

, Majore

, Hass

, and Kasper

Mesenchymal stromal cells derived from human umbilical cord tissues: primitive cells with potential for clinical and tissue engineering applications. Adv Biochem Eng Biotechnol, 123, 29, 2010.

76.

Barlow

, Brooke

, Chatterjee

, Price

, Pelekanos

, Rossetti

, Doody

, Venter

, Pain

, Gilshenan

, and Atkinson

Comparison of human placenta- and bone marrow-derived multipotent mesenchymal stem cells. Stem Cells Dev, 17, 1095, 2008.

77.

Castrechini

N.M.

, Murthi

, Gude

N.M.

, Erwich

J.J.H.M.

, Gronthos

, Zannettino

, Brennecke

S.R.

, and Kalionis

Mesenchymal stem cells in human placental chorionic villi reside in a vascular niche. Placenta, 31, 203, 2010.

78.

Igura

, Zhang

, Takahashi

, Mitsuru

, Yamaguchi

, and Takahashi

T.A.

Isolation and characterization of mesenchymal progenitor cells from chorionic villi of human placenta. Cytotherapy, 6, 543, 2004.

79.

Zhang

X.H.

, Mitsuru

, Igura

, Takahashi

, Ichinose

, Yamaguchi

, and Takahashi

T.A.

Mesenchymal progenitor cells derived from chorionic villi of plus human placenta for cartilage tissue engineering. Biochem Biophys Res Commun, 340, 944, 2006.

80.

da Silva Meirelles

, Caplan

A.I.

, and Nardi

N.B.

In search of the in vivo identity of mesenchymal stem cells. Stem Cells, 26, 2287, 2008.

81.

da Silva Meirelles

, Sand

T.T.

, Harman

R.J.

, Lennon

D.P.

, and Caplan

A.I.

MSC frequency correlates with blood vessel density in equine adipose tissue. Tissue Eng Part A, 15, 221, 2009.

82.

Crisan

, Yap

, Casteilla

, Chen

C.W.

, Corselli

, Park

T.S.

, Andriolo

, Sun

, Zheng

, Zhang

, Norotte

, Teng

P.N.

, Traas

, Schugar

, Deasy

B.M.

, Badylak

, Buhring

H.J.

, Giacobino

J.P.

, Lazzari

, Huard

, and Peault

A perivascular origin for mesenchymal stem cells in multiple human organs. Cell Stem Cell, 3, 301, 2008.

83.

Fuchs

, Tumbar

, and Guasch

Socializing with the neighbors: stem cells and their niche. Cell, 116, 769, 2004.

84.

Jones

D.L.

, and Wagers

A.J.

No place like home: anatomy and function of the stem cell niche. Nat Rev Mol Cell Biol, 9, 11, 2008.

85.

Scadden

D.T.

The stem-cell niche as an entity of action. Nature, 441, 1075, 2006.

86.

Shi

, and Gronthos

Perivascular niche of postnatal mesenchymal stem cells in human bone marrow and dental pulp. J Bone Miner Res, 18, 696, 2003.

87.

Schnorr

Embryologie der Haustiere. Stuttgart, Germany: Enke Verlag, 2001.

88.

Rüsse

Lehrbuch der Embryologie der Haustiere. Berlin and Hamburg, Germany: Paul Parey Verlag, 1991.

89.

Romeis

Mikroskopische Technik. Munich, Germany: Urban & Schwarzenberg, 1989.

90.

Gittel

, Brehm

, Burk

, Juelke

, Staszyk

, and Ribitsch

Isolation of equine multipotent mesenchymal stromal cells by enzymatic tissue digestion or explant technique: comparison of cellular properties. BMC Vet Res, 9, 2013.

91.

Boos

A.M.

, Loew

J.S.

, Deschler

, Arkudas

, Bleiziffer

, Gulle

, Dragu

, Kneser

, Horch

R.E.

, and Beier

J.P.

Directly auto-transplanted mesenchymal stem cells induce bone formation in a ceramic bone substitute in an ectopic sheep model. J Cell Mol Med, 15, 1364, 2011.

92.

Dominici

, Le Blanc

, Mueller

, Slaper-Cortenbach

, Marini

, Krause

, Deans

, Keating

, Prockop

, and Horwitz

Minimal criteria for defining multipotent mesenchymal stromal cells. The international society for cellular therapy position statement. Cytotherapy, 8, 315, 2006.

93.

J.W.

, Kim

J.A.

, and Ha

C.W.

Do the fibroblasts contained in early passage msc population adversely affect the characteristics of stem cell population obtained from human placenta? Int J Stem Cells, 5, 89, 2012.

94.

Kern

, Eichler

, Stoeve

, Kluter

, and Bieback

Comparative analysis of mesenchymal stem cells from bone marrow, umbilical cord blood, or adipose tissue. Stem Cells, 24, 1294, 2006.

95.

Shetty

, Cooper

, and Viswanathan

Comparison of proliferative and multilineage differentiation potentials of cord matrix, cord blood, and bone marrow mesenchymal stem cells. Asian J Transfus Sci, 4, 14, 2010.

96.

Ragni

, Vigano

, Parazzi

, Montemurro

, Montelatici

, Lavazza

, Budelli

, Vecchini

, Rebulla

, Giordano

, and Lazzari

Adipogenic potential in human mesenchymal stem cells strictly depends on adult or foetal tissue harvest. Int J Biochem Cell Biol, 45, 2456, 2013.

97.

Rebelatto

C.K.

, Aguiar

A.M.

, Moretao

M.P.

, Senegaglia

A.C.

, Hansen

, Barchiki

, Oliveira

, Martins

, Kuligovski

, Mansur

, Christofis

, Amaral

V.F.

, Brofman

P.S.

, Goldenberg

, Nakao

L.S.

, and Correa

Dissimilar differentiation of mesenchymal stem cells from bone marrow, umbilical cord blood, and adipose tissue. Exp Biol Med (Maywood), 233, 901, 2008.

98.

Wagner

, Wein

, Seckinger

, Frankhauser

, Wirkner

, Krause

, Blake

, Schwager

, Eckstein

, Ansorge

, and Ho

A.D.

Comparative characteristics of mesenchymal stem cells from human bone marrow, adipose tissue, and umbilical cord blood. Exp Hematol, 33, 1402, 2005.

99.

Redshaw

, Loughan

P.T.

Oxygen concentration modulates the differentiation of muscle stem cells toward myogenic and adipogenic fates. Differentiation, 84, 193, 2012.