Osteogenic proliferation and differentiation of canine bone marrow and adipose tissue derived mesenchymal stromal cells and the influence of hypoxia

Research in Veterinary Science 92 (2012) 66–75

Contents lists available at ScienceDirect

Research in Veterinary Science journal homepage: www.elsevier.com/locate/rvsc

Osteogenic proliferation and differentiation of canine bone marrow and adipose tissue derived mesenchymal stromal cells and the influence of hypoxia Dai-Jung Chung a, Kei Hayashi a, Chrisoula A. Toupadakis b, Alice Wong b, Clare E. Yellowley b,⇑ a b

Department of Surgical and Radiological Sciences, School of Veterinary Medicine, University of California Davis, Davis, CA 95616, United States Department of Anatomy, Physiology, and Cell Biology, School of Veterinary Medicine, University of California Davis, Davis, CA 95616, United States

a r t i c l e

i n f o

Article history: Received 30 June 2010 Accepted 5 October 2010

Keywords: Canine mesenchymal stromal cell Bone marrow Adipose tissue Proliferation Osteogenic differentiation Hypoxia

a b s t r a c t The aim of this study was to compare the osteogenic and proliferative potential of canine mesenchymal stromal cells (cMSCs) derived from bone marrow (BM-cMSCs) and adipose tissue (AT-cMSCs). Proliferation potential was determined under varying oxygen tensions (1%, 5%, and 21% O2). Effects of reduced oxygen levels on the osteogenic differentiation of AT-cMSCs were also investigated. AT-cMSCs proliferated at a significantly faster rate than BM-cMSCs, although both cell types showed robust osteogenic differentiation. Culture in 5% and 1% O2 impaired proliferation in cMSC from both sources and osteogenic differentiation in AT-cMSCs. Our data suggests that AT-cMSCs might be more suitable for use in a clinical situation, where large cell numbers are required for bone repair, due to their rapid proliferation combined with robust osteogenic potential. Our data also suggests that the inhibitory effects of hypoxia on both cell proliferation and differentiation should be considered when using MSCs in a potentially hypoxic environment such as a fracture site. Ó 2010 Elsevier Ltd. All rights reserved.

1. Introduction Many fractures can be repaired by the natural process of bone regeneration with appropriate fixation or coaptation. However, when substantial loss of host bone occurs as a result of trauma, tumor and non- or delayed unions, or when healing potential is compromised by disease or old age, other interventional therapies are required (Csaki et al., 2009; Kraus and Kirker-Head, 2006). Healing is complicated in more than 20% of fractures in human medicine and is a significant problem in veterinary medicine, especially in small breed dogs (Adamiak and Aleksiewicz, 2006; Welch et al., 1997). One option is to use autologous or allogenic cancellous bone grafts, although the quantity of autologous cancellous bone available might be insufficient due to the age or size of the patient, and allogenic bone graft might not be readily available (Kraus and Kirker-Head, 2006). Cell based therapies, in particular the use of mesenchymal stromal cells (MSCs), have recently emerged as an exciting alternative for treating orthopedic disorders and have provoked particular interest in veterinary medicine. MSCs are multipotential, capable of differentiating into a variety of cell lineages such as chondrocytes, adipocytes, osteoblasts, tenocytes, and myocytes under appropriate biochemical, hormonal, and mechanical stimuli in vitro and in vivo (Kadiyala et al., 1997; Neupane et al., 2008; Rebelatto et al., 2008). MSCs may contribute to ⇑ Corresponding author. E-mail address: [email protected] (C.E. Yellowley). 0034-5288/$ - see front matter Ó 2010 Elsevier Ltd. All rights reserved. doi:10.1016/j.rvsc.2010.10.012

tissue regeneration directly by incorporation as newly differentiated cells, or indirectly through production of trophic factors when transplanted at the lesion site (Das et al., 2009). Canine mesenchymal stromal cells (cMSCs) have been successfully isolated from bone marrow (BM) (Csaki et al., 2007; Jafarian et al., 2008; Kadiyala et al., 1997; Kamishina et al., 2008), adipose tissue (AT) (Neupane et al., 2008; Vieira et al., 2009), and other sources (Zucconi et al., 2010), and proved to have the capability of differentiating into osteoblasts in vitro and in vivo (Arinzeh et al., 2003; Jafarian et al., 2008; Kadiyala et al., 1997; Kraus et al., 1999; Neupane et al., 2008; Volk et al., 2005; Zucconi et al., 2010). In fact, BM and AT are presently the most important sources for cMSCs in veterinary medicine. In this study we sought to directly compare the proliferative and osteogenic differentiation capacity of MSCs from these two sources, derived from the same patient, and determine which source might be the most appropriate for therapeutic use. The oxygen environment appears to play a central role in maintaining the self renewal capacity of MSCs and their multipotency (Ma et al., 2009) and also strongly influences both MSC differentiation and proliferation. Given its critical role in MSC function, oxygen availability is often overlooked when considering the use of these cells for in vivo therapeutics or even in vitro tissue engineering. For example the hypoxic nature of the fracture site, which occurs as a direct result of disruption of blood vessels, has been well described (Brighton and Krebs, 1972a,b; Epari et al., 2008; Lu et al., 2008; Seekamp et al., 1995). In addition, cMSCs seeded in 3D scaffolds for tissue engineering may be hypoxic due to the uneven dis-

D.-J. Chung et al. / Research in Veterinary Science 92 (2012) 66–75

tribution of oxygen within the scaffold (Volkmer et al., 2008, 2010). Since MSCs may experience diverse oxygen environments during therapeutic use, we examined the influence of oxygen on proliferation and osteogenic differentiation of cMSCs. Our hypothesis was that cMSCs from different sources have different proliferative and osteogenic capacity. In addition, we hypothesized that lowering O2 tension would modulate cMSC proliferation and differentiation. Our aims were to (1) to compare the proliferative and osteogenic differentiation capacity of cMSCs from bone marrow and adipose tissue; (2) examine the effects of reducing O2 to 5% and 1% on cMSC proliferation and differentiation. 2. Materials and methods 2.1. Animals Paired BM and AT samples were collected from dogs diagnosed with cranial cruciate ligament (CCL) rupture and undergoing surgical treatment with informed consent. Apart from ligament rupture, the animals were otherwise healthy. Samples were collected from four adult dogs between the ages of 4 and 7 years (4.6 ± 1.3 years). 2.2. cMSC isolation from bone marrow BM samples were collected from the iliac crest by aspiration under ambient O2 conditions. Briefly, the iliac crest was palpated and aseptically prepared. A single stab incision was made with a No. 11 scalpel blade through the skin over the intended bone marrow aspiration site. An 18 gauge BM needle (Cardinal Health, USA) was inserted through the stab incision into the flat area of iliac crest. The BM needle was inserted into the bone, angled slightly medially with firm pressure in a twisting motion. When the marrow cavity was penetrated, the stylet was removed and a 50 mL syringe was placed onto the end of the needle (Townsend, 2008). 15–20 mL of BM was drawn into a 50 mL syringe containing 10 mL of heparin (1000 IU/mL, Abraxis, USA). BM-cMSCs were isolated and cultured as previously described (Toupadakis et al., 2010). Briefly, the BM was mixed with an equal volume of Hank’s Balanced Salt Solution (HBSS, Invitrogen, USA), agitated and centrifuged at 300g for 15 min. The supernatant was removed, and the pellet was resuspended in HBSS and centrifuged at 1000g for 5 min. The pellet was resuspended and put into a flask with standard growth media [RPMI (Invitrogen, USA), 10% Fetal Bovine Serum (FBS, Invitrogen, USA), 1% A/A (Antibiotic–Antimycotic, Invitrogen, USA)]. Cells were incubated for 2 days at 37 °C with 5% CO2 in a standard humidified incubator (HeraCell 150, Thermo Scientific, Walton, USA) at which point cultures were washed with HBSS to remove the non-adherent cells. Media was then changed every two days until first passage. 2.3. cMSC isolation from adipose tissue AT samples were collected from subcutaneous fat from the stifle area under ambient O2 conditions before closing the incision. ATcMSCs were isolated using a modification of published protocols (Toupadakis et al., 2010). Briefly, AT was minced with a razor blade into 1–5 mm pieces and washed 4–5 times in 3 volumes of phosphate buffered saline (PBS, Gibco, USA) to remove red blood cells. One volume of collagenase/BSA solution [0.1% collagenase type I (Worthington Biochemical Corporation, USA), 1% BSA (Sigma, USA) in PBS] was added and the tissue was put on a shaker for one hour at 37 °C. Following incubation, the collagenase was neutralized with an equal volume of standard culture media [RPMI, 10% FBS, 1% A/A]. Samples were centrifuged for 5 min, and floating fat was removed. The stromal vascular fraction (SVF) was resus-

67

pended in complete media and cells were plated and cultured for 2 days at 37 °C with 5% CO2 at which point cultures were washed with HBSS to remove the non-adherent cells. Media was then changed every two days until first passage. 2.4. Cell yield and proliferation studies Freshly isolated cells (P0) were expanded until 70% confluent in a humidified incubator at 37 °C with 5% CO2. Before replating for the 1st passage and subsequent expansion, total cell numbers from the primary cell expansion (P0) were counted using a digital automated cell counting system (Countess, Invitrogen, USA) and normalized to milliliter of bone marrow aspirated or gram of fat sample acquired. Passage 1 cMSCs were cultured at 1%, 5%, and 21% O2 at a seeding density of 5000 cells/cm2 in standard media and subcultured every 7 days for 3 weeks. For ambient (21%) O2 tension, cells were cultured in a standard humidified incubator at 37 °C with 5% CO2. For reduced O2 tension experiments, cells were cultured in standard humidified incubators at 37 °C with 5% CO2 and O2 reduced to 1% and 5% using supplemental N2 (Heracell 150, Kendro, USA). Cell number was determined using a digital automated cell counting system at passage 1, 2, and 3. Cells were divided into groups as follows: BM-cMSCs cultured in 1%, 5%, and 21% oxygen tension (BM-1%, BM-5%, and BM-21%), and AT-cMSCs cultured in 1%, 5%, and 21% oxygen tension (AT-1%, AT-5%, and AT-21%). 2.5. Osteogenic differentiation studies Osteogenic differentiation potential was assessed using modifications of previously described protocols (Kadiyala et al., 1997; Pittenger et al., 1999; Toupadakis et al., 2010) and the study design is outlined in Fig. 1. Initial studies examined osteogenic differentiation in the absence of dexamethasone (Sigma, USA). 1 paired BM and AT-cMSC sample was plated at 5000 cells/cm2 in standard media. After 24 h differentiation was initiated by adding osteogenic media (10 mM b-glycerophosphate (Sigma, USA), 50 mg/mL ascorbic acid (Wako, Japan) 10% FBS, 1% A/A in RPMI media) which was changed twice weekly. Cell morphology, Alizarin Red and Alkaline Phosphatase staining were examined up to week 4. To comprehensively compare the osteogenic potential of BM and AT-derived cMSCs, 4 Paired BM and AT-cMSCs (P3–5) samples were plated at 5000 cells/cm2 in standard media. After 24 h, differentiation was initiated by adding osteogenic media supplemented with 100 nM dexamethasone to enhance differentiation, which was changed twice weekly (Kadiyala et al., 1997). Alizarin Red staining was performed on day 1, week 1, and week 2. RNA was collected on day 0, day 1, week 1, and week 2. 2.6. To assess the influence of O2 on differentiation AT-cMSCs were seeded at 5000 cells/cm2 in standard media. After 24 h differentiation was initiated by adding osteogenic media preconditioned for 24 h in 1%, 5% or 21% O2 and cells transferred to incubators at 1%, 5% and 21% O2, respectively. Media changes were performed twice a week. Alizarin Red staining was performed, and RNA collected on days 1, 3, 7, 10, and 14. 2.7. Alizarin red staining for calcium deposition cMSC cultures were washed with PBS and fixed in a 5% paraformaldehyde solution for 15 min at room temperature. After fixation, cells were washed with deionized water and stained with a 0.5% Alizarin Red (Ali R, Sigma, USA) solution for 30 min at room temperature. Cells were washed with deionized water to remove excess stain, dried and imaged using a flatbed scanner.

68

D.-J. Chung et al. / Research in Veterinary Science 92 (2012) 66–75

Fig. 1. Study design for the osteogenic differentiation of cMSCs. Isolated cMSCs were plated at 5000 cells/cm2 and osteogenic media added 24 h later, D0. Total RNA for qRTPCR was collected on days 0, day 1, week 1, and week 2. Stains for alkaline phosphatase (AP) activity and calcium deposition (Alizarin Red, Ali R) were performed on day 1, week 1, and week 2.

Differences were considered statistically significant at a probability level of p < 0.05.

2.8. Alkaline phosphatase activity staining cMSC cultures were stained with a commercial alkaline phosphatase (AP) activity staining kit (Sigma, USA) according to the manufacturer’s instructions. Digital images of air dried plates were acquired using a flatbed scanner. 2.9. RNA collection and real time quantitative RT-PCR Total RNA was extracted from cells using RNeasy Mini Kit (Qiagen, USA) and treated with DNase I (Qiagen, USA). 250–1000 ng of total RNA was reverse-transcribed using Superscript First Strand Synthesis System (Invitrogen, USA) or QuantiTect Reverse Transcription Kit (Qiagen, USA) for RT-PCR in a BioRad iCycler (BioRad, USA). Primer sets were designed from known canine gene sequences or generated from previously published data (Neupane et al., 2008) (Table 1). RNA expression was determined by realtime quantitative RT-PCR. PCR reactions were carried out using Platinum Taq Polymerase (Invitrogen, USA) in an Eppendorf Mastercycler Realplex2 PCR machine (Eppendorf, USA). Samples were run in triplicate and fluorescence was monitored by SYBR Green intercalation into amplicons at each cycle. Cycle conditions were denaturation at 95 °C for 15 s, and 40 cycles of denaturation at 95°C for 15 s, annealing at 60 °C for 15 s, and elongation at 72 °C for 25 s. The final melting curve step was from 60–95 °C over 20 min, and 95 °C for 15 s. The level of expression of target gene, normalized to b2-microglobulin, was then calculated using the dCT method. In this study gene expression of the following were measured: collagen type 1 alpha1(COL1A1), OSTEOCALCIN, runt related transcription factor 2 (RUNX2) and OSTERIX.

3. Results 3.1. Comparison of cell yield We observed a significantly greater cMSC yield at first passage (P0) in AT compared to BM. The average cell yield of AT-cMSCs at P0 was 4.2 ± 1.7  105 cells per gram of tissue compared to 1.2 ± 0.2  105 cells per milliliter of bone marrow aspirate (p < 0.05) (Fig. 2) after 6–8 days in primary culture. 3.2. cMSC proliferation potential and the influence of hypoxia Overall, BM-cMSCs proliferated less rapidly than AT-cMSCs and lowering oxygen tension to 1% significantly reduced cell numbers in both groups, Fig. 3. Cell numbers in the AT-21% group were significantly higher than cell numbers in all other groups at all time points with the exception of AT-5% at P1 and P2. All BM groups showed significantly lower cell numbers compared to the AT-5%, and AT-21% groups at P1, and all BM groups showed significantly lower cell numbers compared to the AT-21% group at P2 and P3 (Fig. 3). MSCs from both BM and AT, at P1, showed significantly lower cell numbers at 1% oxygen tension compared to those grown at 21% oxygen though cell numbers in the 5% and 21% groups were not significantly different. From P4, cells in the 1%-BM and 5%-BM groups did not proliferate at a sufficient rate to continue the experiment.

2.10. Statistical analysis

3.3. Comparison of the osteogenic differentiation potential of BM and AT-cMSCs

Statistical analyses were performed by using SPSS (Version 15.0, IBM, USA). Differences among groups were assessed by unpaired Ttest, and one-way ANOVA test, followed by Tukey’s post hoc test.

In osteogenic media, both BM and AT-cMSCs became cuboidal, aggregated and formed mineralized nodules as evidenced by Alizarin Red staining, Fig. 4A and C. In control media, cells proliferated

Table 1 Primers for the real time RT-PCR. Gene RUNX2 COLLAGEN1A1 (COL1A1) Neupane et al. (2008) OSTERIX Neupane et al. (2008) OSTEOCALCIN Neupane et al. (2008) b2-MICROGLOBULIN Neupane et al. (2008)

Amplicon size Forward Reverse Forward Reverse Forward Reverse Forward Reverse Forward Reverse

TGGGAGAGGTACCAGATGGG TCTTGCCTCGTCCACTCCGG GTAGACACCACCCTCAAGAGC TTCCAGTCGGAGTGGCACATC ACGACACTGGGCAAAGCAG CATGTCCAGGGAGGTGTAGAC GAGGGCAGCGAGGTGGTGAG TCAGCCAGCTCGTCACAGTTGG TCTACATTGGGCACTGTGTCAC TGAAGAGTTCAGGTCTGACCAAG

152 119 285 134 136

D.-J. Chung et al. / Research in Veterinary Science 92 (2012) 66–75

69

Fig. 2. Comparison of cell yields from primary cell expansion of bone marrow (BM) and adipose tissue (AT). Bars represent mean total cMSC count ± SEM (n = 4). * represents a statistically significant difference, p < 0.05.

Fig. 3. Effects of oxygen on the proliferative capacity of AT-cMSCs and BM-cMSCs. Cells were divided into groups as follows: BM-cMSCs cultured in 1%, 5%, and 21% oxygen tension (BM-1%, BM-5%, and BM-21%), and AT-cMSCs cultured in 1%, 5%, and 21% oxygen tension (AT-1%, AT-5%, and AT-21%). Bars represent mean fold increase in cell number over initial seeding density ± SEM (n = 4). * represents a statistically significant difference, p < 0.05. Within each passage, bars with identical lower case letters are significantly different statistically from one another, p < 0.05.

until confluent, remained spindle-shaped or fibroblastic in morphology and nodule formation was not observed, Fig. 4 A. All cMSCs grown in osteogenic media showed positive Alizarin Red staining from week 1 compared to the controls and the intensity of staining increased at each subsequent time point, Fig. 4 B. Alkaline phosphatase activity staining could be visualized with the naked eye in BM-cMSC cultures and microscopically in ATcMSCs, Fig. 4C.

In subsequent differentiation assays 100 nM dexamethasone was added, which has been shown to enhance osteogenesis (Kadiyala et al., 1997) and resulted in stronger Alizarin Red staining in both BM and AT-cMSC cultures, Fig. 5. While all four paired samples showed robust Alizarin Red staining, AT-cMSCs showed stronger staining than their matched BM-cMSCs at weeks 1 and 2, Fig. 5. Inter-animal variability in Alizarin Red staining appeared to be greatest in BM-cMSCs.

70

D.-J. Chung et al. / Research in Veterinary Science 92 (2012) 66–75

Fig. 4. (A) Morphology of cMSCs in standard media (control) and in osteogenic media without dexamethasone (osteogenic). Scale bar: 200 lm. (B) Ali R and AP staining of BM and AT-cMSCs in standard and osteogenic media at day 1 (D1) and weeks 1–4 (W1, 2, 3 and 4). (C) Alizarin Red (Ali R) staining, and Alkaline Phosphatase (AP) activity staining of cMSCs in differentiation media at week 4. Scale bar: 200 lm.

Osteogenic gene expression was compared between BM and AT. Expression levels of OSTERIX were similar in both groups and showed a characteristic transient and early rise. RUNX2 and OSTEOCALCIN levels were also similar between groups, with OSTEOCALCIN levels showing a characteristic, though not significant, rise later in differentiation at week 2. COLLAGEN1A1 (COL1A1) mRNA expression was significantly higher in AT-cMSCs compared to BM-cMSCs at day 1 and week 1 (Fig. 6).

Expression levels of OSTERIX showed a characteristic transient and early rise at all oxygen tensions, with levels significantly higher in 21% O2 up to day 3 and on day 14, Fig. 7B. RUNX2 and OSTEOCALCIN showed the same pattern of expression in all O2 tensions. However there was a trend for elevated RUNX2 and OSTEOCALCIN levels in 21% O2 that reached significance for OSTEOCALCIN at days 1 and 3. COL1A1 mRNA expression was similar across all groups with the exception of significantly higher levels in 5% O2 than in 1% and 21% O2 on day 14, Fig. 7B.

3.4. Effects of hypoxia on osteogenic differentiation 4. Discussion and conclusions Culture of AT-cMSCs in 5% and 1% O2 reduced their osteogenic differentiation as evidenced by a decrease in intensity of Alizarin Red staining, Fig. 7A. Hypoxia also attenuated the proliferative capacity of AT-cMSCs under osteogenic growth conditions (data not shown) in a similar manner to cells grown in standard media, Fig. 3.

Mesenchymal stromal cells are a good resource for regenerative medicine because of their diverse cellular differentiation potential and trophic effects (Caplan and Dennis, 2006). Bone marrow has been studied extensively as a source of MSCs. Some data suggests it may be a superior source of MSCs for osteogenesis since BM-

D.-J. Chung et al. / Research in Veterinary Science 92 (2012) 66–75

Fig. 5. Ali R staining of BM and AT-cMSCs from four different dogs in osteogenic differentiation media containing dexamethasone at day 1 (D1), week 1 and 2 (W1, W2). AT sample ‘‘A’’ proliferated very rapidly and the cell layer peeled off by week 2 (shown by an ‘‘X’’).

MSCs are pre-committed towards the osteogenic lineage (Noel et al., 2008). Adipose tissue is another ideal source of MSCs because it is abundant and can be easily obtained from several regions of the body with minimal donor site morbidity (Neupane et al.,

71

2008; Tapp et al., 2009). In this study, we evaluated the proliferative and osteogenic potential of canine BM-MSCs and AT-MSCs isolated from the same patients. To our knowledge, this study is the first to present a direct comparison of cMSCs from BM and AT. In clinical practice, multiple injections coupled with large numbers of cells per dose results in a requirement for cells with a robust proliferative potential, as adequate cell numbers must be achieved by in vitro cell expansion. In our studies, AT-cMSCs proliferated significantly faster than BM-cMSCs resulting in a significantly higher cell yield at early passages. The proliferative capacity of AT-cMSCs remained significantly greater than that of BM-cMSCs through P3. In fact insufficient numbers of BM cells beyond this point precluded continuation of the study. Our data is consistent with studies that report higher proliferation potential of AT-MSCs than BM-MSCs in rabbits and rats (Lin et al., 2009; Yoshimura et al., 2007). Therefore, in the clinical arena where large cell numbers are required it appears that AT-cMSCs might be most useful. Indeed, serially passaged BM-MSCs have shown to decrease their osteogenic potential and alkaline phosphatase activity (Coelho et al., 2000; Kadiyala et al., 1997; Sugiura et al., 2004). While quantities of bone marrow are fairly limited, large number of stromal cells can be derived from a relatively small amount of adipose tissue. It has been demonstrated that AT-MSCs can be maintained in vitro for extended periods with stable population doubling and low levels of senescence (Kern et al., 2006; Lin et al., 2009; Zuk et al., 2002, 2001). In addition, cell proliferation

Fig. 6. mRNA expression of osteogenic markers OSTERIX, RUNX2, COL1A1 and OSTEOCALCIN in BM-cMSCs (BM) and AT-cMSCs (AT) at various time points in culture with osteogenic media containing dexamethasone (n = 4). Bars represent mean relative values normalized to b2-microglobulin expression ± SEM, Bars with identical letters are statistically different (p < 0.05). The relative values were calculated using the dCT method, expression of the target gene was normalized to b2-microglobulin.

72

D.-J. Chung et al. / Research in Veterinary Science 92 (2012) 66–75

Fig. 7. Effects of hypoxia on osteogenic differentiation of AT-cMSC. (A) AT-cMSC staining for calcium deposition (Alizarin Red) at each time points in 1%, 5%, and 21% oxygen tension (n = 3). (B) mRNA expression of the osteogenic markers OSTERIX, RUNX2, COL1A1, OSTEOCALCIN in induced AT-cMSCs. Data were normalized to b2-microglobulin and analyzed using the delta CT method. Data are expressed as a relative changes to day 0 expression; bars represent mean ± SEM, lower case letters represent a statistically significant differences between the three oxygen groups at each time point (a, b, c, d, e, f, g, h, i = p < 0.05).

rates and osteogenic potential were not affected by age differences in later life (Khan et al., 2009). Interestingly human AT-MSC yield

appears to be more highly correlated with body mass than with age (Aust et al., 2004). In contrast, increasing age appears to nega-

D.-J. Chung et al. / Research in Veterinary Science 92 (2012) 66–75

tively correlate with human BM-MSC yield (Caplan, 1994). AT-MSC proliferation capacity and primary cell yield may be site specific. In a study by Neupane and colleagues (Neupane et al., 2008) cell yield from subcutaneous fat was higher than cell yield from omental and inguinal fat. The cell yield from primary cultures of fat from a subcutaneous region (5.28  105 per gram) was similar to values we report from subcutaneous fat in the stifle (4.16  105 per gram). One of the limitations of this study is that adipose tissue and bone marrow were isolated from dogs diagnosed with CCL rupture. Altered cytokine profiles have been identified in joint tissues from dogs with osteoarthritis secondary to CCL rupture, including synovial membrane and CCL (Maccoux et al., 2007). There was also a trend for differential cytokine expression in adipose tissue taken from the infrapatellar fat pad (Maccoux et al., 2007). We isolated adipose tissue from the subcutaneous region of the affected stifle and bone marrow from the iliac crest of the same animal. Since the effects of joint pathology and severity of disease on the cytokine profile and MSC phenotype with increasing distance from the joint is unclear, the impact of this on our measured proliferation and differentiation profiles is unknown. Interestingly, an abnormality in the bone marrow MSC population was detected before the onset of disease in a murine model of rheumatoid arthritis (Mohanty et al., 2010) which suggests that at least in some cases MSC abnormality may even drive the disease. It is not clear whether MSCs aid tissue regeneration directly by in situ differentiation and matrix production or indirectly via production of trophic factors. For this reason we also compared the osteogenic capacity of cMSCs. Interestingly both sources underwent robust osteogenic differentiation, though AT-cMSCs showed stronger alizarin red staining than their matched BM-cMSCs at weeks 1 and 2 suggesting formation of more bone mineral in these cultures. Interestingly, levels of osteogenic gene expression were comparable between BM and AT-cMSCs with the exception of COL1A1, which was consistently elevated in AT-cMSCs. It is possible that the elevated mineral deposition observed in the AT-cMSC cultures is the result of the higher proliferative capacity of these cells and increased cell numbers in the AT-cMSC dishes. Gene expression, in contrast, is normalized to expression of a housekeeping gene, which indirectly takes into account cell number. It is possible then that these cells do have the same osteogenic potential but that higher cell numbers of AT-cMSCs enabled deposition of more mineral. Similar studies comparing the osteogenic capacity of BM and AT-MSCs have demonstrated greater osteogenic potential in BMMSCs (Hayashi et al., 2008; Im et al., 2005; Liu et al., 2007; Noel et al., 2008; Toupadakis et al., 2010; Vidal et al., 2007). In contrast, other studies indicate no significant differences between these two sources (De Ugarte et al., 2003; Rebelatto et al., 2008) or some even suggest greater osteogenic potential of AT-MSCs (Kern et al., 2006; Lin et al., 2009). Taken together our data suggest that in cases where large MSC numbers are required for an osteogenic application that adipose tissue may be the cell source of choice. Bone marrow and adipose tissue derived mesenchymal stromal cells are reported to deposit mineralized matrix when transplanted in vivo (Arinzeh et al., 2003; Jafarian et al., 2008; Pieri et al., 2010; Rhee et al., 2010). Direct comparisons of osteogenicity of BM and AT-MSCs in vivo are few and contradictory. In one study, BM-MSCs seeded in hydroxyapatite ceramics and implanted subcutaneously produced more bone than AT-MSCs (Hayashi et al., 2008). In a similar study, collagen sponges were seeded with BM-MSCs or ATMSCs and implanted into critical size defects in the ovine (Niemeyer et al., 2010). In this study again BM-MSCs were shown to produce more bone than AT-MSCs. In contrast, AT-MSCs and BMMSCs both failed to make bone in a calvarial defect model unless tranduced with vectors expressing bone morphogenetic protein 4 (BMP-4) (Lin et al., 2009). In contrast, the transduced cells were

73

able to produce mineralized matrix in vitro, with AT-MSCs depositing significantly more calcified extracellular matrix (Lin et al., 2009). Taken together these data suggest that osteogenic capacity in vitro does not always correlate with osteogenic capacity in vivo. Further studies are required to compare and optimize the performance of both of these cell types in in vivo environments. In our study, hypoxia attenuated both cell proliferation and osteogenic differentiation of AT-MSCs. The effects of hypoxia on MSC proliferation are contradictory. Some studies demonstrate that low O2 levels actually enhance cell proliferation (Grayson et al., 2007; Ma et al., 2009; Ren et al., 2006; Volkmer et al., 2010; Xu et al., 2007), while others, like our own suggest the opposite (Holzwarth et al., 2010). This may reflect the wide range of oxygen tensions that are considered to be hypoxic, cell source, seeding densities and time in culture among other things. Consistent with our findings, however, are several reports describing negative effects of hypoxia on the osteogenic differentiation capacity of MSCs (Fehrer et al., 2007; Malladi et al., 2006; Potier et al., 2007; Raheja et al., 2010). In a recent study by Volkmer et al. (2010) hypoxic pre-conditioning of human MSCs restored their osteogenic differentiation potential under hypoxic conditions. In contrast mouse MSCs expanded under hypoxic conditions and then differentiated under normoxic conditions still exhibited decreased osteogenesis (Xu et al., 2007). Another study suggests that MSCs are able to survive hypoxia for a short period of time (<48 h) without losing their osteogenic potential (Potier et al., 2007). Taken together these data suggest that lowering oxygen tension has an overall negative effect on osteogenic differentiation. In the bone marrow niche, stem cells reside in a low O2 environment between 1% and 7% (Watt and Hogan, 2000), and in adipose tissue O2 can be as low as 3% (Ma et al., 2009). It is thought that low O2 may contribute to maintenance of self renewal capacity and multipotency whilst inhibiting differentiation. Therefore it is maybe not surprising that lowering O2 tension had negative effects on osteogenesis. This may have significant consequences however for cells delivered directly to a hypoxic fracture site and their ability to survive and undergo osteogenic differentiation. If the goal is to retain the self renewal capacity and multipotency of MSCs in culture, then lower O2 tensions may be beneficial. However, the effects of O2 perturbation during cell isolation procedures, where bone marrow and adipose tissue are exposed to ambient air is unknown. Further studies to assess the effects of transient exposure to hyperoxia on MSC biology are warranted. The mechanisms by which hypoxia exerts its effects have been the focus of intense study. Hypoxia sensitive transcription factors such as hypoxia inducible factor 1-a (HIF-1a) are thought to regulate gene expression by binding to specific hypoxic response elements (HRE) (D’Angio and Finkelstein, 2000; Semenza, 2007). HIF-1a is ubiquinated and targeted for proteasomal degradation under conditions of normoxia, a process which is inhibited under hypoxia (D’Angio and Finkelstein, 2000; Semenza, 2007). It is likely that similar signal transduction pathways play a role in the inhibition of proliferation and differentiation that we have observed in this study. In summary, AT-cMSCs proliferate more rapidly in in vitro culture when compared to BM-cMSCs. Both cell types showed robust osteogenic potential, however more intense alizarin red staining in AT-cMSC cultures indicated more mineral deposition by these cells. As such, given the need for large cell numbers in a clinical situation, AT may be the superior choice for generation of cMSCs for tissue regeneration purposes. Lowering oxygen tension had negative effects on both cell proliferation and differentiation. This may be of significance when implanting cells into a hypoxic site such as a fracture. Further studies are required to assess the potential positive effects of hypoxic pre-conditioning under low oxygen conditions.

74

D.-J. Chung et al. / Research in Veterinary Science 92 (2012) 66–75

Supported by a private gift from Dick and Carolyn Randall. Also supported in part by the Center for Companion Animal Health and the Center for Equine Health, School of Veterinary Medicine, University of California, Davis. Conflict of interest There are no conflicts of interest to disclose. Acknowledgements We would like to acknowledge and thank Shinya Watari, DVM for his help with cell culture. References Adamiak, Z., Aleksiewicz, R., 2006. Treatment of fractured antebrachial bones in both thoracic extremities in toy breed dogs by application of reconstruction plates. Polish journal of veterinary sciences 9, 201–205. Arinzeh, T.L., Peter, S.J., Archambault, M.P., van den Bos, C., Gordon, S., Kraus, K., Smith, A., Kadiyala, S., 2003. Allogeneic mesenchymal stem cells regenerate bone in a critical-sized canine segmental defect. The Journal of bone and joint surgery. American volume 85-A, 1927–1935. Aust, L., Devlin, B., Foster, S.J., Halvorsen, Y.D., Hicok, K., du Laney, T., Sen, A., Willingmyre, G.D., Gimble, J.M., 2004. Yield of human adipose-derived adult stem cells from liposuction aspirates. Cytotherapy 6, 7–14. Brighton, C.T., Krebs, A.G., 1972a. Oxygen tension of healing fractures in the rabbit. Journal of Bone and Joint Surgery. American Volume 54, 323–332. Brighton, C.T., Krebs, A.G., 1972b. Oxygen tension of nonunion of fractured femurs in the rabbit. Surgery of gynecology and obstetrics 135, 379–385. Caplan, A.I., 1994. The mesengenic process. Clinics in Plastic Surgery 21, 429–435. Caplan, A.I., Dennis, J.E., 2006. Mesenchymal stem cells as trophic mediators. Journal of Cellular Biochemistry 98, 1076–1084. Coelho, M.J., Cabral, A.T., Fernande, M.H., 2000. Human bone cell cultures in biocompatibility testing. Part I: osteoblastic differentiation of serially passaged human bone marrow cells cultured in alpha-MEM and in DMEM. Biomaterials 21, 1087–1094. Csaki, C., Matis, U., Mobasheri, A., Ye, H., Shakibaei, M., 2007. Chondrogenesis, osteogenesis and adipogenesis of canine mesenchymal stem cells: a biochemical, morphological and ultrastructural study. Histochemistry and Cell Biology 128, 507–520. Csaki, C., Matis, U., Mobasheri, A., Shakibaei, M., 2009. Co-culture of canine mesenchymal stem cells with primary bone-derived osteoblasts promotes osteogenic differentiation. Histochemistry and Cell Biology 131, 251–266. D’Angio, C.T., Finkelstein, J.N., 2000. Oxygen regulation of gene expression: a study in opposites. Molecular Genetics and Metabolism 71, 371–380. Das, R., Jahr, H., van Osch, G., Farrell, E., 2009. The role of hypoxia in MSCs: considerations for regenerative medicine approaches. Tissue Engineering Part B Review. De Ugarte, D.A., Morizono, K., Elbarbary, A., Alfonso, Z., Zuk, P.A., Zhu, M., Dragoo, J.L., Ashjian, P., Thomas, B., Benhaim, P., Chen, I., Fraser, J., Hedrick, M.H., 2003. Comparison of multi-lineage cells from human adipose tissue and bone marrow. Cells Tissues Organs 174, 101–109. Epari, D.R., Lienau, J., Schell, H., Witt, F., Duda, G.N., 2008. Pressure, oxygen tension and temperature in the periosteal callus during bone healing – An in vivo study in sheep. Bone 43, 734–739. Fehrer, C., Brunauer, R., Laschober, G., Unterluggauer, H., Reitinger, S., Kloss, F., Gully, C., Gassner, R., Lepperdinger, G., 2007. Reduced oxygen tension attenuates differentiation capacity of human mesenchymal stem cells and prolongs their lifespan. Aging Cell 6, 745–757. Grayson, W.L., Zhao, F., Bunnell, B., Ma, T., 2007. Hypoxia enhances proliferation and tissue formation of human mesenchymal stem cells. Biochemical and Biophysical Research Communications 358, 948–953. Hayashi, O., Katsube, Y., Hirose, M., Ohgushi, H., Ito, H., 2008. Comparison of osteogenic ability of rat mesenchymal stem cells from bone marrow, periosteum, and adipose tissue. Calcified Tissue International 82, 238–247. Holzwarth, C., Vaegler, M., Gieseke, F., Pfister, S.M., Handgretinger, R., Kerst, G., Muller, I., 2010. Low physiologic oxygen tensions reduce proliferation and differentiation of human multipotent mesenchymal stromal cells. BMC Cell Biology 11, 11. Im, G.I., Shin, Y.W., Lee, K.B., 2005. Do adipose tissue-derived mesenchymal stem cells have the same osteogenic and chondrogenic potential as bone marrowderived cells? Osteoarthritis and Cartilage 13, 845–853. Jafarian, M., Eslaminejad, M.B., Khojasteh, A., Mashhadi Abbas, F., Dehghan, M.M., Hassanizadeh, R., Houshmand, B., 2008. Marrow-derived mesenchymal stem cells-directed bone regeneration in the dog mandible: a comparison between biphasic calcium phosphate and natural bone mineral. Oral Surgery, Oral Medicine, Oral Pathology, Oral Radiology and Endodontics 105, e14–24. Kadiyala, S., Young, R.G., Thiede, M.A., Bruder, S.P., 1997. Culture expanded canine mesenchymal stem cells possess osteochondrogenic potential in vivo and in vitro. Cell Transplantation 6, 125–134.

Kamishina, H., Farese, J.P., Storm, J.A., Cheeseman, J.A., Clemmons, R.M., 2008. The frequency, growth kinetics, and osteogenic/adipogenic differentiation properties of canine bone marrow stromal cells. In Vitro Cellular and Developmental Biology - Animal 44, 472–479. Kern, S., Eichler, H., Stoeve, J., Kluter, H., Bieback, K., 2006. Comparative analysis of mesenchymal stem cells from bone marrow, umbilical cord blood, or adipose tissue. Stem cells 24, 1294–1301. Khan, W.S., Adesida, A.B., Tew, S.R., Andrew, J.G., Hardingham, T.E., 2009. The epitope characterisation and the osteogenic differentiation potential of human fat pad-derived stem cells is maintained with ageing in later life. Injury 40, 150– 157. Kraus, K.H., Kirker-Head, C., 2006. Mesenchymal stem cells and bone regeneration. Veterinary Surgery 35, 232–242. Kraus, K.H., Kadiyala, S., Wotton, H., Kurth, A., Shea, M., Hannan, M., Hayes, W.C., Kirker-Head, C.A., Bruder, S., 1999. Critically sized osteo–periosteal femoral defects: a dog model. Journal of Investigative Surgery 12, 115–124. Lin, L., Shen, Q., Wei, X., Hou, Y., Xue, T., Fu, X., Duan, X., Yu, C., 2009. Comparison of osteogenic potentials of BMP4 transduced stem cells from autologous bone marrow and fat tissue in a rabbit model of calvarial defects. Calcified Tissue International 85, 55–65. Liu, T.M., Martina, M., Hutmacher, D.W., Hui, J.H., Lee, E.H., Lim, B., 2007. Identification of common pathways mediating differentiation of bone marrow- and adipose tissue-derived human mesenchymal stem cells into three mesenchymal lineages. Stem cells 25, 750–760. Lu, C., Rollins, M., Hou, H., Swartz, H.M., Hopf, H., Miclau, T., Marcucio, R.S., 2008. Tibial fracture decreases oxygen levels at the site of injury. Iowa Orthopaedic Journal 28, 14–21. Ma, T., Grayson, W.L., Frohlich, M., Vunjak-Novakovic, G., 2009. Hypoxia and stem cell-based engineering of mesenchymal tissues. Biotechnology Progress 25, 32– 42. Maccoux, L.J., Salway, F., Day, P.J., Clements, D.N., 2007. Expression profiling of select cytokines in canine osteoarthritis tissues. Veterinary Immunology and Immunopathology 118, 59–67. Malladi, P., Xu, Y., Chiou, M., Giaccia, A.J., Longaker, M.T., 2006. Effect of reduced oxygen tension on chondrogenesis and osteogenesis in adipose-derived mesenchymal cells. American journal of physiology. Cell physiology 290, C1139–1146. Mohanty, S.T., Kottam, L., Gambardella, A., Nicklin, M.J., Coulton, L., Hughes, D., Wilson, A.G., Croucher, P.I., Bellantuono, I., 2010. Alterations in the self-renewal and differentiation ability of bone marrow mesenchymal stem cells in a mouse model of rheumatoid arthritis. Arthritis research & therapy 12, R149. Neupane, M., Chang, C.C., Kiupel, M., Yuzbasiyan-Gurkan, V., 2008. Isolation and characterization of canine adipose-derived mesenchymal stem cells. Tissue Engineering Part A 14, 1007–1015. Niemeyer, P., Fechner, K., Milz, S., Richter, W., Suedkamp, N.P., Mehlhorn, A.T., Pearce, S., Kasten, P., 2010. Comparison of mesenchymal stem cells from bone marrow and adipose tissue for bone regeneration in a critical size defect of the sheep tibia and the influence of platelet-rich plasma. Biomaterials 31, 3572– 3579. Noel, D., Caton, D., Roche, S., Bony, C., Lehmann, S., Casteilla, L., Jorgensen, C., Cousin, B., 2008. Cell specific differences between human adipose-derived and mesenchymal-stromal cells despite similar differentiation potentials. Experimental Cell Research 314, 1575–1584. Pieri, F., Lucarelli, E., Corinaldesi, G., Aldini, N.N., Fini, M., Parrilli, A., Dozza, B., Donati, D., Marchetti, C., 2010. Dose-dependent effect of adipose-derived adult stem cells on vertical bone regeneration in rabbit calvarium. Biomaterials 31, 3527–3535. Pittenger, M.F., Mackay, A.M., Beck, S.C., Jaiswal, R.K., Douglas, R., Mosca, J.D., Moorman, M.A., Simonetti, D.W., Craig, S., Marshak, D.R., 1999. Multilineage potential of adult human mesenchymal stem cells. Science 284, 143–147. Potier, E., Ferreira, E., Andriamanalijaona, R., Pujol, J.P., Oudina, K., LogeartAvramoglou, D., Petite, H., 2007. Hypoxia affects mesenchymal stromal cell osteogenic differentiation and angiogenic factor expression. Bone 40, 1078– 1087. Raheja, L.F., Genetos, D.C., Yellowley, C.E., 2010. The effect of oxygen tension on the long-term osteogenic differentiation and MMP/TIMP expression of human mesenchymal stem cells. Cells Tissues Organs 191, 175–184. Rebelatto, C.K., Aguiar, A.M., Moretao, M.P., Senegaglia, A.C., Hansen, P., Barchiki, F., Oliveira, J., Martins, J., Kuligovski, C., Mansur, F., Christofis, A., Amaral, V.F., Brofman, P.S., Goldenberg, S., Nakao, L.S., Correa, A., 2008. Dissimilar differentiation of mesenchymal stem cells from bone marrow, umbilical cord blood, and adipose tissue. Bulletin of experimental biology and Medicine (Maywood) 233, 901–913. Ren, H., Cao, Y., Zhao, Q., Li, J., Zhou, C., Liao, L., Jia, M., Cai, H., Han, Z.C., Yang, R., Chen, G., Zhao, R.C., 2006. Proliferation and differentiation of bone marrow stromal cells under hypoxic conditions. Biochemical and Biophysical Research Communications 347, 12–21. Rhee, S.C., Ji, Y.H., Dhong, E.S., Park, S.H., Yoon, E.S., 2010. In vivo evaluation of mixtures of uncultured freshly-isolated adipose-derived stem cells and demineralized bone matrix for bone regeneration in a rat critically-sized calvarial defect model. Stem Cells and Development 2, 69–74. Seekamp, A., van Griensven, M., Ziegler, M., Gunderoth, M., 1995. Transcutaneous PO2 measurement in compound fractures. European journal of emergency medicine 2, 69–74. Semenza, G.L., 2007. Hypoxia-inducible factor 1 (HIF-1) pathway. Science STKE cm8.

D.-J. Chung et al. / Research in Veterinary Science 92 (2012) 66–75 Sugiura, F., Kitoh, H., Ishiguro, N., 2004. Osteogenic potential of rat mesenchymal stem cells after several passages. Biochemical and Biophysical Research Communications 316, 233–239. Tapp, H., Hanley Jr., E.N., Patt, J.C., Gruber, H.E., 2009. Adipose-derived stem cells: characterization and current application in orthopaedic tissue repair. Experimental biology and medicine (Maywood) 234, 1–9. Toupadakis, C.A., Wong, A., Genetos, D.C., Cheung, W.K., Borjesson, D.L., Ferraro, G.L., Galuppo, L.D., Leach, J.K., Owens, S.D., Yelloley, C.E., 2010. Comparison of the Osteogenic Potential of Equine Mesenchymal Stem Cells from Bone Marrow, Adipose Tissue, Umbilical Cord Blood, and Umbilical Cord Tissue. American journal of veterinary research 71, 1237–1245. Townsend 3rd, F.I., 2008. Bone marrow aspiration in dogs and cats. Laboratory Animal (NY) 37, 497–498. Vidal, M.A., Kilroy, G.E., Lopez, M.J., Johnson, J.R., Moore, R.M., Gimble, J.M., 2007. Characterization of equine adipose tissue-derived stromal cells: adipogenic and osteogenic capacity and comparison with bone marrow-derived mesenchymal stromal cells. Veterinary Surgery 36, 613–622. Vieira, N., Brandalise, V., Zucconi, E., Secco, M., Strauss, B., Zatz, M., 2009. Isolation, characterization and differentiation potential of canine adipose-derived stem cells. Cell Transplant. Volk, S.W., Diefenderfer, D.L., Christopher, S.A., Haskins, M.E., Leboy, P.S., 2005. Effects of osteogenic inducers on cultures of canine mesenchymal stem cells. American Journal of Veterinary Research 66, 1729–1737. Volkmer, E., Drosse, I., Otto, S., Stangelmayer, A., Stengele, M., Kallukalam, B.C., Mutschler, W., Schieker, M., 2008. Hypoxia in static and dynamic 3D culture systems for tissue engineering of bone. Tissue Engineering Part A 14, 1331– 1340.

75

Volkmer, E., Kallukalam, B.C., Maertz, J., Otto, S., Drosse, I., Polzer, H., Bocker, W., Stengele, M., Docheva, D., Mutschler, W., Schieker, M., 2010. Hypoxic preconditioning of human mesenchymal stem cells overcomes hypoxia-induced inhibition of osteogenic differentiation. Tissue Engineering Part A 16, 153–164. Watt, F.M., Hogan, B.L., 2000. Out of Eden: stem cells and their niches. Science 287, 1427–1430. Welch, J.A., Boudrieau, R.J., DeJardin, L.M., Spodnick, G.J., 1997. The intraosseous blood supply of the canine radius: implications for healing of distal fractures in small dogs. Veterinary Surgery 26, 57–61. Xu, Y., Malladi, P., Chiou, M., Bekerman, E., Giaccia, A.J., Longaker, M.T., 2007. In vitro expansion of adipose-derived adult stromal cells in hypoxia enhances early chondrogenesis. Tissue Engineering 13, 2981–2993. Yoshimura, H., Muneta, T., Nimura, A., Yokoyama, A., Koga, H., Sekiya, I., 2007. Comparison of rat mesenchymal stem cells derived from bone marrow, synovium, periosteum, adipose tissue, and muscle. Cell and Tissue Research 327, 449–462. Zucconi, E., Vieira, N.M., Bueno, D.F., Secco, M., Jazedje, T., Ambrosio, C.E., PassosBueno, M.R., Miglino, M.A., Zatz, M., 2010. Mesenchymal stem cells derived from canine umbilical cord vein-A novel source for cell therapy studies. Stem Cells and Development. Zuk, P.A., Zhu, M., Mizuno, H., Huang, J., Futrell, J.W., Katz, A.J., Benhaim, P., Lorenz, H.P., Hedrick, M.H., 2001. Multilineage cells from human adipose tissue: implications for cell-based therapies. Tissue Engineering 7, 211–228. Zuk, P.A., Zhu, M., Ashjian, P., De Ugarte, D.A., Huang, J.I., Mizuno, H., Alfonso, Z.C., Fraser, J.K., Benhaim, P., Hedrick, M.H., 2002. Human adipose tissue is a source of multipotent stem cells. Molecular Biology of the Cell 13, 4279–4295.