Advertisement

BMC Veterinary Research

, 15:201 | Cite as

The effect of hypoxia on chondrogenesis of equine synovial membrane-derived and bone marrow-derived mesenchymal stem cells

  • Alexis L. Gale
  • Renata M. Mammone
  • Michael E. Dodson
  • Renata L. Linardi
  • Kyla F. OrtvedEmail author
Open Access
Research article
Part of the following topical collections:
  1. Musculoskeletal disorders and orthopaedics

Abstract

Background

Joint injury is extremely common in equine athletes and post-traumatic osteoarthritis (PTOA), a progressive and debilitating disease, is estimated to affect 60% of horses in the USA. The limited potential for intrinsic healing of articular cartilage has prompted intense efforts to identify a cell-based repair strategy to prevent progression of PTOA. Mesenchymal stem cells (MSCs) have the potential to become an ideal source for cell-based treatment of cartilage lesions; however, full chondrogenic differentiation remains elusive. Due to the relatively low oxygen tension in articular cartilage, hypoxia has been proposed as a method of increasing MSC chondrogenesis. The objective of this study was to investigate the effect of hypoxic culture conditions on chondrogenesis in equine synovial membrane-derived MSCs (SM-MSCs) and bone marrow-derived MSCs (BM-MSCs). MSCs were isolated from synovial membrane and bone marrow collected from 5 horses. Flow cytometric analysis was used to assess cell surface marker expression including CD29, CD44, CD90, CD105, CD45, CD-79α, MHCI and MHCII. MSC pellets were cultured in normoxic (21% O2) or in hypoxic (5% O2) culture conditions for 28 days. Following the culture period, chondrogenesis was assessed by histology, biochemical analyses and gene expression of chondrogenic-related genes including ACAN, COL2b, SOX9, and COL10A1.

Results

Both cell types expressed markers consistent with stemness including CD29, CD44, CD90, CD105, and MHCI and were negative for exclusion markers (CD45, CD79α, and MHCII). Although the majority of outcome variables of chondrogenic differentiation were not significantly different between cell types or culture conditions, COL10A1 expression, a marker of chondrocyte hypertrophy, was lowest in hypoxic SM-MSCs and was significantly lower in hypoxic SM-MSCs compared to hypoxic BM-MSCs.

Conclusions

Hypoxic culture conditions do not appear to increase chondrogenesis of equine SM-MSCs or BM-MSCs; however, hypoxia may downregulate the hypertrophic marker COL10A1 in SM-MSCs.

Keywords

Mesenchymal stem cell Equine Synovial membrane Bone marrow Normoxia Hypoxia Chondrogenesis 

Abbreviations

ACAN

Aggrecan

ACI

Autologous chondrocyte implantation

BM-MSC

Bone marrow-derived mesenchymal stem cell

CD

Cluster of differentiation

COL10A1

Collagen type 10

COL2b

Collagen type 2

DMMB

Dimethylmethylene blue

FACS

Fluorescent-activated cell sorting

GAG

Glycosaminoglycan

HIF

Hypoxia-inducible factor

MHC

Major histocompatibility complex

MSC

Mesenchymal stem cell

NSAID

Non-steroidal anti-inflammatory drug

P2

Passage 2

PBS

Phosphate buffered saline

PTOA

Post-traumatic osteoarthritis

SM-MSC

Synovial membrane-derived mesenchymal stem cell

SOX9

Sex determining region Y-box 9

Background

Joint injury and cartilage damage are extremely common in equine athletes and often precipitate post-traumatic osteoarthritis (PTOA), a progressive and debilitating disease. Due to the poor intrinsic healing capabilities of cartilage, focal chondral defects are repaired with biomechanically inferior fibrocartilage following injury [1, 2]. In contrast to hyaline cartilage, fibrocartilage is unable to provide adequate compressive and tensile strength, thereby facilitating global joint degradation. Palliative care consists of non-steroidal anti-inflammatory drugs (NSAIDs) and corticosteroid therapy both of which have deleterious long-term side effects [3, 4]. Currently, no effective disease-modifying drugs that halt or reverse PTOA are available [5]. Repairing damaged articular cartilage following injury may prevent further joint degeneration and would be greatly beneficial.

Cell-based cartilage repair strategies have been intensely investigated; however, a suitable cell source for regeneration of hyaline cartilage remains elusive. In human orthopedic surgery, autologous chondrocyte implantation (ACI) has been the “gold-standard” for repair of large cartilaginous defects [6, 7]. Both autologous and allogeneic chondrocyte implantation have been described with some success in the horse [8, 9]. Despite improved clinical outcomes and healing, ACI has several limitations including the need for multiple surgical procedures, graft hypertrophy [10], and donor site morbidity [11]. Considering the limitations of chondrocyte implantation, an alternative cell source for resurfacing the articular surface, such as mesenchymal stem cells (MSCs), would be favorable.

MSCs are an ideal cell source as they are easily accessible, can be culture-expanded, and are multipotent with chondrogenic differentiation capabilities. The goal of MSC-based cartilage repair is for chondrogenesis of MSCs implanted into chondral defects facilitating replacement of hyaline cartilage. The vast majority of cell-based cartilage repair has been focused on bone marrow-derived MSCs (BM-MSCs); however, recently, synovial membrane-derived MSCs (SM-MSCs) have been investigated as a cell source due to demonstration of superior chondrogenesis in other species [12]. Synovium can be harvested in standing horses or during arthroscopic procedures, with SM-MSCs being isolated and expanded in the laboratory in preparation for chondrogenic differentiation.

Lack of complete MSC chondrogenic differentiation and progression towards the hypertrophic phenotype, with increased expression of collagen type X (COL10A1) remains a challenge in cell-based cartilage repair [13, 14]. Differences in culture conditions appear to be an important factor in the effectiveness of MSC chondrogenesis. Chondrocytes reside in a relatively hypoxic environment of 1–5% O2 (8-40mmHg) compared to other tissues in the body, including bone marrow which is at 7% O2 (50mmHg) [15, 16]. Traditional CO2 incubators are maintained at 21% O2 and 5% CO2, which may limit chondrogenesis in MSC cultures. Relative hypoxia has had variable effects on chondrogenesis thus far, with some studies demonstrating enhanced chondrogenesis in BM-MSCs and articular cartilage progenitor (ACP) pellet cultures with upregulation of collagen type II, aggrecan and SOX9 [17]. Hypoxia increases expression of hypoxia-inducible factors (HIFs), which play a significant role in signaling pathways of chondrogenesis, including SOX9, a key transcription factor of chondrogenesis [18, 19]. Ranera et al. (2013) demonstrated improved chondrogenesis in equine BM-MSCs cultured in hypoxic conditions [20]; however, studies investigating the effect of hypoxia on SM-MSCs are lacking.

Currently, there are no published studies evaluating the effects of hypoxia on equine SM-MSCs.

The main objective of this study was to compare the chondrogenic capabilities of equine SM-MSCs and BM-MSCs in hypoxic and normoxic culture conditions. We hypothesized that hypoxic culture conditions would increase chondrogenesis in both BM-MSCs and SM-MSCs but that SM-MSCs would have superior chondrogenesis compared to BM-MSCs.

Results

Immunophenotyping

The immunophenotypes of passage 2 (P2) BM-MSCs and SM-MSCs, as analyzed by flow cytometry, were similar between cell types with both cell types displaying cell surface antigen expression characteristic of MSCs (Fig. 1). BM-MSCs and SM-MSCs were strongly positive for expression of CD29, CD44, CD90, CD105 and MHCI. BM-MSCs and SM-MSCs were also negative for expression of the hematopoietic cell surface markers CD45RB and CD79α. As demonstrated in previous studies in our laboratory (unpublished data), BM-MSCs had variable expression of the exclusion marker, MHCII, with 14.48 ± 0.221% cells expressing MHCII, while expression of MHCII by SM-MSCs was negligible 3.02 ± 0.028%.
Fig. 1

Characterization of BM-MSCs and SM-MSCs using flow cytometric quantification of cell surface marker expression. a Expression of cell surface markers expected to be positive in MSC populations and b expression of cell surface markers expected to be negative in MSC populations. The white histograms represent isotype controls and black histograms represent respective cell surface marker staining. The mean ± SEM percentage of positive cells is in the corner of each histogram. Each histogram is a representative result of 5 horses

Chondrogenic differentiation

Chondrogenic differentiation potential of pellet cultures of SM-MSCs and BM-MSCs in normoxic and hypoxic conditions was compared by assessing MSC pellets at the end of a 28-day period. Grossly, normoxic BM-MSC pellets were larger and rounder than normoxic SM-MSC pellets, while hypoxic BM-MSC and SM-MSC pellets were similar in size and shape to normoxic BM-MSC pellets (Fig. 2). Histologically, BM-MSC pellets cultured under normoxic and hypoxic conditions exhibited more intense toluidine blue staining, consistent with proteoglycan deposition, than SM-MSC pellets cultured in either oxygen tension (Fig. 2).
Fig. 2

Photomicrographs of BM-MSC and SM-MSC pellets cultured in normoxic (21% O2) and hypoxic (5% O2) conditions for 28 days. Pellets were stained with H&E and toluidine blue (scale bar = 100 μm)

Glycosaminoglycan content, as quantified by the DMMB assay, was not significantly different between any of the treatment groups (Fig. 3). Additionally, DNA content and GAG/DNA ratio was not significantly different between any of the treatment groups (Fig. 3).
Fig. 3

Mean ± SEM a) GAG, b) DNA, and c) GAG:DNA content in BM-MSC and SM-MSC pellets cultured in normoxic (21% O2) and hypoxic (5% O2) conditions for 28 days

Expression of markers of chondrogenesis including SOX9, ACAN, and COL2b displayed variability as noted in Fig. 4. Expression of SOX9 was higher in normoxic and hypoxic BM-MSCs compared to SM-MSCs, whereas expression of ACAN and COL2b were both higher in normoxic and hypoxic SM-MSCs compared to BM-MSCs. There was a statistical trend for COL2b expression to be higher in hypoxic SM-MSCs compared to normoxic BM-MSCs (p = 0.0641). COL10A1 expression, a marker of hypertrophy, was significantly lower in hypoxic SM-MSCs compared to hypoxic BM-MSCs (p = 0.0339) (Fig. 4).
Fig. 4

Relative expression (mean ± SEM) of chondrogenic-related genes including SOX9, ACAN, COL2b and COL10A1 for BM-MSC and SM-MSC pellets cultured in normoxic (21% O2) and hypoxic (5% O2) conditions for 28 days. Expression is relative to normoxic BM-MSC pellets. 18S was used as a reference gene in all analyses. * p < 0.05

Adipogenic and osteogenic differentiation potential

Adipogenic differentiation was observed in both SM-MSCs and BM-MSCs with cells demonstrating lipid droplet deposition via positive staining with Oil Red O 14 days following adipogenic induction (Fig. 5). Control SM-MSCs and BM-MSCs that were not cultured in adipogenic media did not show evidence of adipogenic differentiation histologically. Osteogenic differentiation of SM-MSCs and BM-MSCs was evident following 14 days of culture in osteogenic media. Alizarin red staining was used to assess presence of calcium, with both cell types demonstrating positive staining compared to control cells (Fig. 5). Control cultures of SM-MSCs and BM-MSCs cultured in basal medium did not show any histologic evidence of differentiation.
Fig. 5

Photomicrographs of control and induced BM-MSCs and SM-MSCs stained with alizarin red (osteogenesis) and Oil Red O (adipogenesis). Osteogenically induced MSCs stained with alizarin red are positive for extracellular calcium consistent with osteogenesis. Adipogenically induced MSCs demonstrated positive Oil Red O staining of lipid droplets consistent with adipogenesis. (Bar = 100 μm)

Discussion

The aim of this study was to investigate the effects of hypoxia on chondrogenesis of equine SM-MSCs and BM-MSCs. Although many studies have evaluated chondrogenesis of equine MSCs derived from different sources, including bone marrow, synovial fluid, and adipose tissue, consistent and complete chondrogenic differentiation remains elusive [21, 22, 23]. In attempts to improve chondrogenesis, different culture conditions have been investigated, including relative hypoxia as chondrocytes reside in a low oxygen environment in the body(~ 1–5% O2), especially compared to standard incubator oxygen tension (~ 21% O2) [16, 24, 25]. To date, no studies have evaluated the effects of hypoxia on equine SM-MSCs. In this study, we demonstrated that overall chondrogenic differentiation was not significantly different between SM-MSCs and BM-MSCs cultured in either normoxic or hypoxic conditions. However, we found that COL10A1 expression, a marker of chondrocyte hypertrophy, was significantly downregulated in SM-MSCs cultured in hypoxia.

In order to evaluate our cell populations for cell surface markers consistent with MSCs prior to chondrogenesis in hypoxic conditions, we evaluated the immunophenotypes of SM-MSCs and BM-MSCs using flow cytometry. We found that both populations of cells were positive for cell surface markers consistent with MSCs including CD29, CD44, CD90, and CD105 and negative for exclusion markers including CD45, CD79α, and MHCII. As demonstrated in previous studies in our laboratory (unpublished data), expression of MHCII was negligible in SM-MSCs (mean 3.02%), while interestingly BM-MSCs had more variable expression of MHCII (mean 14.48%) with 47.24% of cells expressing MHCII in one horse. MHCII is generally considered an exclusion marker; however, variability of MHCII expression in equine BM-MSCs has been shown repeatedly by our laboratory (unpublished data) and others [26]. These findings suggest that equine BM-MSCs may exhibit more variability in MHCII expression than equine SM-MSCs, which may be important when considering the clinical application of allogeneic MSCs. Increased immunogenicity and cell rejection has been associated with increased expression of MHCII by equine BM-MSCs and has been associated with increased immunogenicity due to allorecognition [26]. In this study we did not investigate the effect of hypoxia on immunophenotype or cell viability as hypoxia has been previously shown to have no significant effect on either factor in equine MSCs [27].

Collagen type X expression is often increased in osteoarthritis and is used as a marker of the undesirable hypertrophic phenotype in chondrocytes and chondrogenically differentiating MSCs. The majority of studies have demonstrated that hypoxia is effective at downregulating COL10A1 expression and protein synthesis [28, 29, 30], although upregulation has been less frequently reported [31, 32]. In the study reported here, we found that COL10A1 expression was lowest in SM-MSCs cultured in hypoxia and this was significantly downregulated compared to BM-MSCs cultured in hypoxia. This could have important implications for future studies looking to optimize culture conditions for SM-MSCs in which chondrogenesis is desirable.

Despite decreased COL10A1 expression in SM-MSCs, there did not appear to be a significant difference in chondrogenesis between cell types or culture conditions. In fact, only moderate chondrogenesis was apparent at the end of the 28-day culture period when differentiated pellets were evaluated histologically and biochemically. Lack of complete chondrogenesis of equine MSCs continues to be significant hurdle with many studies investigating the effects of different culture conditions including oxygen tension, cell type, and growth factors [24, 25]. Interestingly, Anderson et al.(2016) recently demonstrated that chondrogenic potential and lack of the hypertrophic response were present in low oxygen tension; however, this only held true for cells with high intrinsic chondrogenic capacity at baseline prior to differentiation [17]. Similar variability in chondrogenic capacity of cells has been demonstrated by others [33, 34], highlighting the importance of cell-to-cell variation in MSC cultures. Although different culture conditions, such as hypoxia, may slightly alter chondrogenic capacity, pre-sorting cells using fluorescent-activated cell sorting (FACS) such that cells with high intrinsic chondrogenic potential are selected may be a far more useful tool. For example, enhanced chondrogenesis has been previously demonstrated by selecting for LNGFR+THY-1+ [34] cells or CD105+ cells [35].

Hypoxia is thought to promote chondrogenesis through hypoxia-inducible factor pathways including HIF-2α-mediated induction of SOX-9 [36] and HIF-1α-mediated inhibition of COL1A1 [37]. In this study, there was increased expression of SOX-9 and COL2b in BM-MSCs and SM-MSCs cultured in hypoxia compared to normoxia although these increases did not reach statistical significance. This may be in part due to the intra- and inter-animal variability of MSC populations. Additionally, culturing MSCs in pellet form may represent an inherent issue with oxygen tension as there is a natural gradient created across the pellet, despite constant incubator conditions, leading to cells within the pellet being exposed to variable oxygen tensions. For example, Markway et al. (2010) found that micropellets (~ 170 cells/micropellet) demonstrated superior chondrogenesis in hypoxic conditions compared to larger pellets (~ 2 × 105 cells/pellet) [38].

Another important factor involved in chondrogenesis appears to be pre-differentiation MSC expansion conditions. Ranera et al. (2013) compared the effect of MSC expansion conditions on future chondrogenesis in normoxic conditions and found that cells expanded in hypoxic conditions had increased ECM formation during 21-day pellet culture when compared to pellets formed from cells expanded in normoxic conditions. Pellets formed from hypoxia-expanded cells demonstrated increased GAG content and more intense Alcian blue and Safranin O staining [20]. Similar results have been shown in human BM-MSCs [39] and ovine BM-MSCs [40], with cells that were expanded in hypoxic conditions prior to pellet culture displaying more robust chondrogenesis regardless of culture conditions (normoxia or hypoxia) during pellet culture.

In the study reported here, MSCs were expanded under normoxic conditions prior to pellet formation, however, hypoxic expansion of cells may be indicated in future studies.

Few studies have investigated the effect of oxygen tension on chondrogenesis of SM-MSCs or synovial fluid-derived MSCs (SF-MSCs). As described for other cell types, the effect of hypoxia on chondrogenesis of these cell types appears to be variable. Both Bae et al. (2018) and Li et al. (2011) showed improved chondrogenesis in human SM-MSCs cultured under hypoxic conditions [41, 42]. However, Ohara et al. (2016) found no effect of hypoxia on the chondrogenic potential of human SM-MSCs and Neybecker et al. (2018) found minimal effects of hypoxia on chondrogenesis of human SF-MSCs obtained from OA joints [43, 44]. To the authors’ knowledge, this is the first study describing the effect of hypoxia on chondrogenesis of equine SM-MSCs. Similar to other studies, we did not detect a major effect of hypoxia on chondrogenesis of SM-MSCs.

Overall, considerable variability appears to exist within and between equine MSC populations. Although hypoxia may inhibit COL10A1 expression in SM-MSCs, further refinement of culture conditions including pre-sorting and selection of cells with high chondrogenic potential and MSC expansion in hypoxic conditions should be considered to optimize chondrogenesis. Additionally, culturing MSCs in three-dimensional scaffolds could be considered as this has shown to improve chondrogenesis in previous studies [45].

Conclusions

Enhanced cartilage repair using chondrogenically differentiated MSCs would be an ideal clinical resource; however, chondrogenesis of equine MSC cultures continues to represent a significant challenge. Synovial membrane-derived MSCs did not demonstrate improved chondrogenesis under hypoxic conditions. Further optimization of culture conditions is indicated for equine MSCs with efforts focused on pre-selection of MSCs with superior chondrogenic differentiation capabilities using FACS and expansion of MSCs in hypoxic conditions prior to induction of chondrogenesis.

Methods

Animals

Five systemically healthy adult horses (2–7 years) being euthanized at the University of Pennsylvania for reasons unrelated to the study were used to tissue collection. The study was performed following approval by the University of Pennsylvania Institutional Animal Care and Use Committee (IACUC #805973).

MSC isolation and culture

Bone marrow was collected under sterile conditions from the sternebrae of horses immediately following euthanasia. Using an 11-gauge Jamshidi bone marrow biopsy needle (VWR Scientific, Bridgeport, NJ) containing 10,000 U of heparin, 40 mL of bone marrow was aspirated. Bone marrow samples were processed via density centrifugation with Ficoll-Paque Plus (GE Healthcare, Chicago, IL, USA) prior to seeding into flasks containing medium consisting of Dulbecco’s Modified Eagle Medium (DMEM) with 1 g/L of D-glucose, 2 mM L-glutamine, and 1 mM sodium pyruvate (ThermoFisher Scientific, Hampton, NH), penicillin (100 U/mL)-streptomycin (100 μg/mL) solution (Invitrogen, Carlsbad, CA), 10% fetal bovine serum (FBS) (VWR Life Science Seradigm, VWR, Radnor, PA), and basic fibroblastic growth factor (bFGF, 1 ng/mL) (Invitrogen, Carlsbad, CA). Media was changed every 48 h.

Synovial membrane was collected from the same horses immediately following bone marrow aspiration. All synovial membrane was collected aseptically from the dorsal aspect of the antebrachiocarpal and middle carpal joint of grossly normal carpi. Following harvest, synovial membrane was rinsed in phosphate buffered saline (PBS) with penicillin (100 U/mL) and streptomycin (100 μg/mL). Synovial membrane (~ 400 mg) was then finely cut into small pieces with a #10 scalpel blade and incubated at 37 °C in 200 μL FBS for 20 min, as previously described [46]. Samples were re-suspended and cultured in DMEM with 4.5 g/L D-glucose, 2 mM L-Glutamine, and 1 mM sodium pyruvate, penicillin (100 U/mL)-streptomycin (100 μg/mL) solution, and 10% FBS. Media was changed every 48 h. Synovial membrane pieces were maintained in the flask until migration of MSCs was confirmed by the presence of MSC colonies attached to the tissue culture flasks.

Both BM-MSCs and SM-MSCs were passaged when they reached ~ 80% confluency using Trypsin-EDTA Cell Dissociation Reagent (ThermoFisher Scientific, Waltham, MA). Passage 2 (P2) cells used for all differentiation assays. Cell number and viability was determined using the Cellometer Auto 2000 Cell Viability Counter (Nexcelom Bioscience, Lawrence, MA) and ViaStain™ AOPI staining solution (Nexcelom Bioscience LLC, Lawrence, MA).

Immunophenotyping of MSCs

Passage 2 BM-MSCs and SM-MSCs were immunophenotyped using flow cytometry. Following trypsinization, cells (6 × 104) were placed in 96-well round bottom plates and washed twice with PBS. Cell pellets were resuspended in 100 μL of PBS with 0.5% bovine serum albumin (BSA) (Sigma Aldrich, St. Louis, MO) and 0.02% sodium azide (ThermoFisher scientific, Waltham, MA) and incubated at 4 °C for 20 min. Cells were then incubated with 50 μL of the appropriate primary antibody at 4 °C for 45 min, rinsed twice with PBS, and then resuspended in the secondary antibody (50 μL) when appropriate and incubated at 4 °C for 45 min. After the final PBS rinse, the pellets were re-suspended in 200 μL of PBS containing 7-AAD (7-Aminoactinomycin D, ThermoFisher scientific, Waltham, MA) as a viability stain. Cells were stained with anti-CD29, CD44, CD90, CD105, CD45, CD-79α, MHCI and MHCII antibodies and isotype controls were used to establish fluorescent gates (Table 1). Flow cytometry and subsequent analysis were performed using the Cytoflex S Benchtop Flow Cytometer and CytExpert Software, version 1.0 (Beckman Coulter Inc., Brea, CA).
Table 1

Antibodies used for flow cytometric analysis of equine cell surface markers

Antibody

Clone/ Isotype

Host Species

Target Species

Fluorophore

2o Antibody

Company

Dilution for 1o Antibody

CD29

TMD29/IgG1

Mouse

Human

APC

Yes

Goat anti-mouse IgG

EMD Millipore

1:100

CD44

IM7/IgG2b

Rat

Human

FITC

No

Thermo IM7

1:80

CD90

?/IgM

Mouse

Canine, Equine

RPE

No

WSU Monoclonal Antibody Center

1:200

CD105

SN6/IgG1

Mouse

Human

Alexa 488

No

Bio Rad

1:10

CD45RB

?/IgM

Mouse

Equine

RPE

No

WSU Monoclonal Antibody Center

1:200

CD79α

HM57/IgG1

Mouse

Human

Alexa 647

No

Bio Rad

1:200

MHCI

cz3/IgG2b

Mouse

Equine

APC

Yes

Goat anti-mouse IgG

Gifta

1:100

MHCII

cz11/IgG1

Mouse

Equine

APC

Yes

Goat anti-mouse IgG

Gifta

1:200

Isotype Control

Corresponding MAB

 

Target Species

Fluorophore

 

Company

Dilution

IgG1

To CD29

 

Mouse

APC

 

Abcam

1:100

IgG2b

To CD44

 

Rat

Alexa 488

 

Abcam

1:100

IgM

To CD90

 

Mouse

PE

 

Abcam

1:200

IgG1

To CD105

 

Mouse

Alexa 488

 

Abcam

1:200

IgM

To CD45RB

 

Mouse

PE

 

Abcam

1:200

IgG1

To CD79α

 

Mouse

Alexa 647

 

Abcam

1:400

IgG2b

To MHCI

 

Mouse

APC

 

Abcam

1:100

IgG1

To MHCII

 

Mouse

APC

 

Abcam

1:100

aGifts from Dr. Doug Antczak, Cornell University, Ithaca, New York, USA

Chondrogenic differentiation assay

For chondrogenic differentiation, 500,000 P2 cells were pelleted in 15 mL conical tubes via centrifugation at 400 g for 5 min. After 48 h in the appropriate basal media for the cell type, chondrogenesis was induced with chondrogenic media containing of DMEM with 4.5 g/L D-glucose with 1% sodium pyruvate and L-Glutamine (4 mM), HEPES buffer (25 mM), penicillin (100 U/mL)-streptomycin (100 μg/mL) solution supplemented with transforming growth factor-β3 (0.01 μg/mL) (ThermoFisher Scientific, Waltham, MA), dexamethasone (0.4 μg/mL), 2- phospho-L-ascorbic acid (0.05 μg/mL), proline (0.04 mg/mL) (ThermoFisher Scientific, Waltham, MA), 1% insulin-transferrin-selenium solution (ThermoFisher Scientific, Waltham, MA), and 1% FBS. Pellets were maintained in normoxic (21% O2) or in hypoxic (5% O2) culture conditions for 28 days. At the end of the 28-day culture period, pellets were fixed in a 10% formalin solution prior to paraffin embedding and sectioning. Pellet sections at the thickness of 5 μm obtained from the center of the pellet were then stained with hematoxylin and eosin (H&E) and toluidine blue.

Adipogenic and osteogenic differentiation assays

Adipogenic and osteogenic differentiation assays were performed in normoxic (21% O2) only to demonstrate multipotency of both cell types. For adipogenic differentiation, cells were seeded into 6-well tissue culture plates containing basal medium at a density of 5100 cells/ cm2. After 48 h, the medium in the treatment wells was changed to adipogenic induction medium consisting of the basal differentiation medium outlined above supplemented with biotin (8 μg/mL) (Sigma-Aldrich, St. Louis, MO), calcium pantothenate (4 μg/mL) (Sigma-Aldrich, St. Louis, MO), insulin (5.8 μg/mL) (Sigma-Aldrich, Stl Louis, MO), dexamethasone (4 μg/mL), isobutylmethylxanthine (0.1 mg/mL) (Sigma-Aldrich, St. Louis, MO), rosiglithizone (0.0178 mg/mL) (Sigma-aldrich, St. Louis, MO), 5% rabbit serum (ThermoFisher Scientific, Waltham, MA), and 3% FBS. Medium was changed every 48 h. After 6 days in induction medium, the medium was changed to adipogenic maintenance medium using the same reagents except rosiglithisone or isobutylmethylxanthine. For each horse, control SM-MSCs and BM-MSCs were maintained in the cell-type specific basal medium for the duration of the culture. Following 14 days of culture, cells were rinsed with PBS and fixed with 10% formalin before staining with Oil Red O (Sigma-Aldrich Corp., St. Louis, MO) for confirmation of lipid droplet accumulation in the cytoplasm of cells.

For osteogenic differentiation, cells were seeded into 6-well culture plates in SM-MSC or BM-MSC medium at a seeding density of 2900 cells/cm2. After 48 h, osteogenic differentiation medium was added containing basal differentiation medium consisting of Advanced DMEM/F12, 1% sodium pyruvate (Gibco Life Technologies, Carlsbad, CA), 25 mM HEPES buffer, 4 mM L-glutamine (ThermoFisher Scientific, Waltham, MA), and penicillin (100 U/mL)-streptomycin (100 μg/mL) solution. The basal medium was supplemented with β-glycerophosphate (2.2 μg/mL) (Sigma Aldrich, St. Louis, MO), dexamethasone (8 μg/mL), 2-phospho-L-ascorbic acid (0.05 mg/mL) (Sigma-Aldrich, St. Louis, MO), and 10% FBS. Cells are cultured in osteogenic medium for 14 days. Media was changed every 48 h. For each horse, control SM-MSCs and BM-MSCs were maintained in basal medium appropriate to the cell type for the duration of the culture. Following 14 days of culture, cells were rinsed with PBS and fixed with 10% formalin before staining with 2% Alizarin Red (Sigma-Aldrich, St. Louis, MO) at pH 4.2 for confirmation of extra-cellular calcium matrix.

Gene expression

For assessment of chondrogenic differentiation, two pellets were collected from each treatment group. Pellets were biopulverized in liquid nitrogen using a multiple sample stainless steel biopulverizer and hammer (BioSpec Products, Inc., Bartlesville, OK). RNA was extracted using the Qiagen RNeasy Fibrous Tissue Mini Kit (Qiagen, Germantown, MD). RNA concentration and purity were quantified with a UV microspectrophotometer (NanoDrop™ One, ThermoFisher Scientific, Waltham, MA). Complementary DNA was prepared using a High Capacity cDNA Reverse Transcription kit (ThermoFisher Scientific, Waltham, MA) and an Eppendorf master cycler (Hamburg, Germany). Real-time quantitative PCR was performed using TaqMan™ Master mix and the Applied Biosystems™ QuantStudio™ 6 Flex Real-Time PCR System (Applied Biosystem, Foster City, CA). The following genes were analyzed: aggrecan (ACAN), collagen type II (COL2b), SRY-box 9 (SOX9), and collagen type X (COL10A1) for chondrogenesis. Primers and probes for ACAN, COL2b, and SOX9 were designed using NCBI Primer-BLAST and Integrated DNA Technologies (IDT) PrimerQuest Tool software and synthesized by IDT (Coralville, IA) (Table 2). Primers and probes for COL10A1 were obtained from ThermoFisher Scientific’s proprietary equine-specific gene expression assay database (ARCE46U). All samples were run in triplicate using 18S as a reference gene. The cycle threshold (CT) values for triplicates were averaged and data were analyzed using the ΔCt method where fold change is expressed as 2-ΔΔCt using normoxic BM-MSCs as the calibrator.
Table 2

Equine primer and probe sequences used for gene expression analyses

Gene

Primer and probe sequences

18S, 18 small ribonucleic acid

Forward, 5′- GCCGCTAGAGGTGAAATTCT-3′

Reverse, 5′- TCGGAACTACGACGGTATCT −3′

Probe, 5′- AAGACGGACCAGAGCGAAAGCAT-3′

ACAN, aggrecan

Forward, 5′-GAGGAGATGGAGGGTGAGGT −3′

Reverse, 5′-GATGGTGATGTCCTCCTCGC-3′

Probe, 5′-TTCACCTGTGTAGCAGATGGCGTC-3’

COL2b, type II collagen

Forward, 5’-GCTACACTCAAGTCCCTCAAC-3′

Reverse, 5′-ATCCAGTAGTCTCCGCTCTT-3′

Probe, 5′-ACCTGAAACTCTGCCACCCTGAAT-3’

SOX9, SRY-box 9

Forward, 5’-CTGGAGACTGCTGAACGAGA-3′

Reverse, 5′-GAGATGTGTGTCTGCTCCGT − 3′

Probe, 5′-AGAAGGACCACCCGGACTACAAGTA-3’

Biochemical analyses

BM-MSC and SM-MSC pellets were collected and stored at -20 °C in medium prior to biochemical assays. The dimethylmethylene blue (DMMB) spectrophotometric assay (Sigma-Aldrich, St. Louis, MO) was used to quantify proteoglycan content in pellets digested in 0.5 mg/mL papain (Sigma Aldrich St. Louis, MO). Chondroitin-4 sulfate (Sigma-Aldrich, St. Louis, MO) was used to establish a standard curve and the optical density determined at 525 nm [47]. Total DNA content was determined using 0.5 mg/mL papain digested pellets incubated for 24 h at 65 °C. Digested samples were then mixed bisbenzimide compound (Hoescht, Sigma-Aldrich, Burlington, MA) and DNA was quantified using a fluorometric assay with an excitation wavelength of 348 nm and an emission wavelength of 456 nm. Calf thymus DNA (Sigma-Aldrich, St. Louis, MO) was used to establish a standard curve. Proteoglycan concentration was normalized to the quantity of DNA in that sample.

Statistical analysis

Continuous values are expressed as means ± SEM. A mixed effects model was used to analyze all continuous data including cell surface marker expression, fold change gene expression, and GAG content. Horse was considered as a random effect. Statistically significant differences between treatments were determined using a Wilcoxon rank sum test. All data were analyzed using JMP14 (SAS, Cary, NC). Significance was set at p < 0.05.

Notes

Acknowledgements

The authors would like to thank Karie Durynski for her technical assistance with histology.

Authors’ contributions

ALG contributed to study design, acquisition, analysis and interpretation of data, and preparation of the final manuscript. RLL, RMM and MED contributed to acquisition of data. KO contributed to study design, acquisition, analysis and interpretation of data, and preparation of the manuscript. All authors approved the final manuscript.

Funding

This work was supported by the Raymond Firestone Trust Research Grant at the University of Pennsylvania. The grant supported the cost of consumables and assays. The funding agency did not play any role in the design of the study, collection, analysis and interpretation of data, or writing the manuscript.

Ethics approval

The Institutional Animal Care and Use Committee of the University of Pennsylvania approved the use of horses in these studies. All horses were privately owned and diagnosed with chronic neurological or musculoskeletal diseases. Horses were donated to the University of Pennsylvania following written consent for use.

Consent for publication

Not applicable.

Competing interests

The authors declare that they have no competing interests.

References

  1. 1.
    Mankin HJ. The response of articular cartilage to mechanical injury. J Bone Jt Surgery Am. 1982;64(3):460–6.CrossRefGoogle Scholar
  2. 2.
    Strauss EJ, Goodrich LR, Chen CT, Hidaka C, Nixon AJ. Biochemical and biomechanical properties of lesion and adjacent articular cartilage after chondral defect repair in an equine model. Am J Sports Med. 2005;33(11):1647–53.CrossRefGoogle Scholar
  3. 3.
    Maiden L, Thjodleifsson B, Seigal A, et al. Long-term effects of nonsteroidal anti-inflammatory drugs and cyclooxygenase-2 selective agents on the small bowel: a cross-sectional capsule enteroscopy study. Clin Gastroenterol Hepatol. 2007;5(9):1040–5.CrossRefGoogle Scholar
  4. 4.
    Forget H, Lacroix A, Bourdeau I, Cohen H. Long-term cognitive effects of glucocorticoid excess in Cushing’s syndrome. Psychoneuroendocrinology. 2016;65:26–33.CrossRefGoogle Scholar
  5. 5.
    Brown TD, Johnston RC, Saltzman CL, Marsh JL, Buckwalter JA. Posttraumatic osteoarthritis: a first estimate of incidence, prevalence, and burden of disease. J Orthop Trauma. 2006;20(10):739–44.CrossRefGoogle Scholar
  6. 6.
    Brittberg M, Lindahl A, Nilsson A, Ohlsson C, Isaksson O, Peterson L. Treatment of deep cartilage defects in the knee with autologous chondrocyte transplantation. N Engl J Med. 1994;331(14):889–941.CrossRefGoogle Scholar
  7. 7.
    Bartlett W, Skinner JA, Gooding CR, et al. Autologous chondrocyte implantation versus matrix-induced autologous chondrocyte implantation for osteochondral defects of the knee: a prospective, randomised study. J bone Jt Surgery Br. 2005;87(5):640–5.CrossRefGoogle Scholar
  8. 8.
    Ortved KF, Nixon AJ, Mohammed HO, Fortier LA. Treatment of subchondral cystic lesions of the medial femoral condyle of mature horses with growth factor enhanced chondrocyte grafts: a retrospective study of 49 cases. Equine Vet J. 2012;44(5):606–13.CrossRefGoogle Scholar
  9. 9.
    Hendrickson DA, Nixon AJ, Grande DA, et al. Chondrocyte-fibrin matrix transplants for resurfacing extensive articular cartilage defects. J Orthop Res. 1994;12(4):485–97.CrossRefGoogle Scholar
  10. 10.
    Peterson L, Minas T, Brittberg M, Nilsson A, Sjogren-Jansson E, Lindahl A. Two- to 9-year outcome after autologous chondrocyte transplantation of the knee. Clin Orthop Relat Res. 2000;374(374):212–34.CrossRefGoogle Scholar
  11. 11.
    Pearce SG, Hurtig MB, Clarnette R, Kalra M, Cowan B, Miniaci A. An investigation of 2 techniques for optimizing joint surface congruency using multiple cylindrical osteochondral autografts. Arthroscopy. 2001;17(1):50–5.CrossRefGoogle Scholar
  12. 12.
    Koga H, Muneta T, Nagase T, et al. Comparison of mesenchymal tissues-derived stem cells for in vivo chondrogenesis: suitable conditions for cell therapy of cartilage defects in rabbit. Cell Tissue Res. 2008;333(2):207–15.CrossRefGoogle Scholar
  13. 13.
    Johnstone B, Hering TM, Caplan AI, Goldberg VM, Yoo JU. In vitro chondrogenesis of bone marrow-derived mesenchymal progenitor cells. Exp Cell Res. 1998;238(1):265–72.CrossRefGoogle Scholar
  14. 14.
    Deng Y, Lei G, Lin Z, Yang Y, Lin H, Tuan RS. Engineering hyaline cartilage from mesenchymal stem cells with low hypertrophy potential via modulation of culture conditions and Wnt/β-catenin pathway. Biomaterials. 2019;192:569–78.CrossRefGoogle Scholar
  15. 15.
    Lund-Olesen K. Oxygen tension in synovial fluids. Arthritis Rheum. 1970;13(6):769–76.CrossRefGoogle Scholar
  16. 16.
    Brighton CT, Heppenstall RB. Oxygen tension in zones of the epiphyseal plate, the metaphysis and diaphysis. An in vitro and in vivo study in rats and rabbits. J Bone Joint Surg Am. 1971;53(4):719–28.CrossRefGoogle Scholar
  17. 17.
    Anderson DE, Markway BD, Bond D, McCarthy HE, Johnstone B. Responses to altered oxygen tension are distinct between human stem cells of high and low chondrogenic capacity. Stem Cell Res Ther. 2016;7(1):154.CrossRefGoogle Scholar
  18. 18.
    Lafont JE. Lack of oxygen in articular cartilage: consequences for chondrocyte biology. Int J Exp Pathol. 2010;91(2):99–106.CrossRefGoogle Scholar
  19. 19.
    Murphy CL, Thoms BL, Vaghjiani RJ, Hypoxia LJE. HIF-mediated articular chondrocyte function: prospects for cartilage repair. Arthritis Res Ther. 2009;11(1):213.CrossRefGoogle Scholar
  20. 20.
    Ranera B, Remacha AR, Álvarez-Arguedas S, et al. Expansion under hypoxic conditions enhances the chondrogenic potential of equine bone marrow-derived mesenchymal stem cells. Vet J. 2013;195(2):248–51.CrossRefGoogle Scholar
  21. 21.
    Kisiday JD, Kopesky PW, Evans CH, Grodzinsky AJ, McIlwraith CW, Frisbie DD. Evaluation of adult equine bone marrow- and adipose-derived progenitor cell chondrogenesis in hydrogel cultures. J Orthop Res. 2008;26(3):322–31.CrossRefGoogle Scholar
  22. 22.
    Watts AE, Ackerman-Yost JC, Nixon AJ. A comparison of three-dimensional culture systems to evaluate in vitro chondrogenesis of equine bone marrow-derived mesenchymal stem cells. Tissue Eng Part A. 2013;19(19–20):2275–83.CrossRefGoogle Scholar
  23. 23.
    Prado AAF, Favaron PO, da Silva LCLC, Baccarin RYA, Miglino MA, Maria DA. Characterization of mesenchymal stem cells derived from the equine synovial fluid and membrane. BMC Vet Res. 2015;11(1):281.CrossRefGoogle Scholar
  24. 24.
    Desancé M, Contentin R, Bertoni L, et al. Chondrogenic differentiation of defined equine mesenchymal stem cells derived from umbilical cord blood for use in cartilage repair therapy. Int J Mol Sci. 2018;19(2):537.CrossRefGoogle Scholar
  25. 25.
    Branly T, Contentin R, Desancé M, et al. Improvement of the chondrocyte-specific phenotype upon equine bone marrow mesenchymal stem cell differentiation: influence of culture time, transforming growth factors and type I collagen siRNAs on the differentiation index. Int J Mol Sci. 2018;19(2):435.CrossRefGoogle Scholar
  26. 26.
    Schnabel LV, Pezzanite LM, Antczak DF, Felippe MJ, Fortier LA. Equine bone marrow-derived mesenchymal stromal cells are heterogeneous in MHC class II expression and capable of inciting an immune response in vitro. Stem Cell Res Ther. 2014;5(1):13.CrossRefGoogle Scholar
  27. 27.
    Ranera B, Remacha A, Álvarez-Arguedas S, et al. Effect of hypoxia on equine mesenchymal stem cells derived from bone marrow and adipose tissue. BMC Vet Res. 2012;8(1):142.CrossRefGoogle Scholar
  28. 28.
    Leijten J, Georgi N, Moreira Teixeira L, van Blitterswijk CA, Post JN, Karperien M. Metabolic programming of mesenchymal stromal cells by oxygen tension directs chondrogenic cell fate. Proc Natl Acad Sci. 2014;111(38):13954–9.CrossRefGoogle Scholar
  29. 29.
    Henrionnet C, Liang G, Roeder E, et al. * Hypoxia for mesenchymal stem cell expansion and differentiation: the best way for enhancing TGFß-induced chondrogenesis and preventing calcifications in alginate beads. Tissue Eng Part A. 2017;23(17–18):913–22.CrossRefGoogle Scholar
  30. 30.
    Pattappa G, Johnstone B, Zellner J, Docheva D, Angele P. The importance of physioxia in mesenchymal stem cell chondrogenesis and the mechanisms controlling its response. Int J Mol Sci. 2019;20(3):484.CrossRefGoogle Scholar
  31. 31.
    Meretoja VV, Dahlin RL, Wright S, Kasper FK, Mikos AG. The effect of hypoxia on the chondrogenic differentiation of co-cultured articular chondrocytes and mesenchymal stem cells in scaffolds. Biomaterials. 2013;34(17):4266–73.CrossRefGoogle Scholar
  32. 32.
    Müller J, Benz K, Ahlers M, Gaissmaier C, Mollenhauer J. Hypoxic conditions during expansion culture prime human mesenchymal stromal precursor cells for chondrogenic differentiation in three-dimensional cultures. Cell Transplant. 2011;20(10):1589–602.CrossRefGoogle Scholar
  33. 33.
    Mabuchi Y, Morikawa S, Harada S, et al. LNGFR(+)THY-1(+)VCAM-1(hi+) cells reveal functionally distinct subpopulations in mesenchymal stem cells. Stem Cell Rep. 2013;1(2):152–65.CrossRefGoogle Scholar
  34. 34.
    Ogata Y, Mabuchi Y, Yoshida M, et al. Purified human synovium mesenchymal stem cells as a good resource for cartilage regeneration. PLoS One. 2015;10(6):e0129096.CrossRefGoogle Scholar
  35. 35.
    Harvanová D, Tóthová T, Sarišský M, Amrichová J, Rosocha J. Isolation and characterization of synovial mesenchymal stem cells. Folia Biol (Praha). 2011;57(3):119–24.Google Scholar
  36. 36.
    Lafont JE, Talma S, Murphy CL. Hypoxia-inducible factor 2α is essential for hypoxic induction of the human articular chondrocyte phenotype. Arthritis Rheum. 2007;56(10):3297–306.CrossRefGoogle Scholar
  37. 37.
    Duval E, Bouyoucef M, Leclercq S, Baugé C, Boumédiene K. Hypoxia inducible factor 1 alpha down-regulates type i collagen through Sp3 transcription factor in human chondrocytes. IUBMB Life. 2016;68(9):756–63.CrossRefGoogle Scholar
  38. 38.
    Markway BD, Tan G-K, Brooke G, Hudson JE, Cooper-White JJ, Doran MR. Enhanced Chondrogenic differentiation of human bone marrow-derived mesenchymal stem cells in low oxygen environment micropellet cultures. Cell Transplant. 2010;19(1):29–42.CrossRefGoogle Scholar
  39. 39.
    Adesida AB, Mulet-Sierra A, Jomha NM. Hypoxia mediated isolation and expansion enhances the chondrogenic capacity of bone marrow mesenchymal stromal cells. Stem Cell Res Ther. 2012;3(2):9.CrossRefGoogle Scholar
  40. 40.
    Bornes TD, Jomha NM, Mulet-Sierra A, Adesida AB. Hypoxic culture of bone marrow-derived mesenchymal stromal stem cells differentially enhances in vitro chondrogenesis within cell-seeded collagen and hyaluronic acid porous scaffolds. Stem Cell Res Ther. 2015;6(1):84.CrossRefGoogle Scholar
  41. 41.
    Li J, He F, Pei M. Creation of an in vitro microenvironment to enhance human fetal synovium-derived stem cell chondrogenesis. Cell Tissue Res. 2011;345(3):357–65.CrossRefGoogle Scholar
  42. 42.
    Bae HC, Park HJ, Wang SY, Yang HR, Lee MC, Han H-S. Hypoxic condition enhances chondrogenesis in synovium-derived mesenchymal stem cells. Biomater Res. 2018;22(1):28.CrossRefGoogle Scholar
  43. 43.
    Ohara T, Muneta T, Nakagawa Y, et al. Hypoxia enhances proliferation through increase of colony formation rate with chondrogenic potential in primary synovial mesenchymal stem cells. J Med Dent Sci. 2016;63(4):61–70.PubMedGoogle Scholar
  44. 44.
    Neybecker P, Henrionnet C, Pape E, et al. In vitro and in vivo potentialities for cartilage repair from human advanced knee osteoarthritis synovial fluid-derived mesenchymal stem cells. Stem Cell Res Ther. 2018;9(1):329.CrossRefGoogle Scholar
  45. 45.
    Rodenas-Rochina J, Kelly DJ, Gómez Ribelles JL, Lebourg M. Influence of oxygen levels on chondrogenesis of porcine mesenchymal stem cells cultured in polycaprolactone scaffolds. J Biomed Mater Res Part A. 2017;105(6):1684–91.CrossRefGoogle Scholar
  46. 46.
    Fülber J, Maria DA, da Silva LCLC, Massoco CO, Agreste F, Baccarin RYA. Comparative study of equine mesenchymal stem cells from healthy and injured synovial tissues: an in vitro assessment. Stem Cell Res Ther. 2016;7:35.CrossRefGoogle Scholar
  47. 47.
    Farndale RW, Sayers CA, Barrett AJ. A direct spectrophotometric microassay for sulfated glycosaminoglycans in cartilage cultures. Connect Tissue Res. 1982;9:247–8.CrossRefGoogle Scholar

Copyright information

© The Author(s). 2019

Open AccessThis article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated.

Authors and Affiliations

  1. 1.Department of Clinical Studies, New Bolton Center, School of Veterinary MedicineUniversity of PennsylvaniaKennett SquareUSA

Personalised recommendations