• View in gallery
    Figure 1—

    Photomicrographs of immunofluorescence labeling of canine PMSCs. Notice strong staining for stem cell surface–specific marker CD90 and lack of staining for hematopoietic stem cell surface markers CD34 and CD45 and for CD146. Immunofluorescent stain; bar = 100 μm.

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    Figure 2—

    Results of RT-PCR analysis and gel electrophoresis for mRNA expression of pluripotency-associated transcription factors SOX2, OCT4, and NANOG and the housekeeping gene β2 MICROGLOBULIN in each of 4 canine tissue–derived MSCs (BMSCs, AMSCs, MMSCs, and PMSCs). The 4 bands above each transcription factor label correspond (from left to right) with MSCs derived from bone marrow, adipose tissue, muscle, and periosteum, respectively. The DNA ladders are labeled L, and each band corresponds to 100 bp, with the lowest band representing 100 bp.

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    Figure 3—

    Photomicrographs of canine BMSCs, AMSCs, MMSCs, and PMSCs cultured in adipogenic or standard medium for 21 days. Notice staining of lipids in MSCs cultured in adipogenic medium and lack of staining in MSCs cultured in standard medium. Oil red O stain; bar = 40 μm.

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    Figure 4—

    Photomicrographs of canine MMSCs cultured in chondrogenic medium with (+) and without (−) hTGF-β1 for 28 days. Notice that there is no appreciable difference in the cellular morphology or staining characteristics between the 2 culture conditions and that there is no evidence of chondrogenic differentiation. A large central area of tissue necrosis with a mesenchymal population in the periphery of each photomicrograph is visible. H&E, Alcian blue, and Safranin O stains; bar = 100 μm.

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    Figure 5—

    Photomicrographs of canine BMSCs, AMSCs, MMSCs, and PMSCs cultured for 8 weeks in osteogenic or standard medium. Notice stained nodules in MSCs cultured in osteogenic medium and no staining in MSCs cultured in standard medium. Von Kossa stain; bar = 40 μm.

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    Figure 6—

    Photomicrographs of canine BMSCs, AMSCs, MMSCs, and PMSCs cultured for 8 weeks in osteogenic or standard medium. Notice stained nodules in MSCs cultured in osteogenic medium and no staining in MSCs cultured in standard medium. Alizarin red stain; bar = 40 μm.

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    Figure 7—

    Results of RT-PCR analysis and gel electrophoresis for mRNA expression of osteogenic transcription factors ALP (b), RUNX2 (c), OSTERIX (d), and OSTEOPONTIN (e) in canine MSCs derived from bone marrow (B), adipose tissue (A), muscle (M), and periosteum (P) cultured in osteogenic medium for 4 weeks (OM4), 6 weeks (OM6), and 8 weeks (OM8). Notice that basal expression of these transcription factors was not seen in any of the 4 tissue-derived sources of MSCs cultured in standard medium, for 4 weeks (SM4). Bone marrow–derived MSCs cultured in SM4 had nonspecific gene expression that was inconsistent with OSTEOPONTIN. The housekeeping gene β2-microglobulin (a) was expressed in all tissue-derived MSCs regardless of culture conditions. The positive control tissue (c), cortical bone cultured in SM4, reveals mRNA expression of ALP, RUNX2, OSTERIX, and OSTEOPONTIN as expected. See Figure 2 for remainder of key.

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Isolation, characterization, and in vitro proliferation of canine mesenchymal stem cells derived from bone marrow, adipose tissue, muscle, and periosteum

Agatha H. KisielCompanion Animal Department, Atlantic Veterinary College, University of Prince Edward Island, Charlottetown, PE C1A 4P3, Canada.

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Laurie A. McDuffeeComparative Orthopaedic Research Laboratory, Department of Health Management, Atlantic Veterinary College, University of Prince Edward Island, Charlottetown, PE C1A 4P3, Canada.

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Elmabrok MasaoudCentre of Veterinary Epidemiological Research, Atlantic Veterinary College, University of Prince Edward Island, Charlottetown, PE C1A 4P3, Canada.
Department of Statistics, Faculty of Science, University of Al-Jabal Al-Garbi, Zawia, Libya.

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Trina R. BaileyCompanion Animal Department, Atlantic Veterinary College, University of Prince Edward Island, Charlottetown, PE C1A 4P3, Canada.

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Blanca P. Esparza GonzalezComparative Orthopaedic Research Laboratory, Department of Health Management, Atlantic Veterinary College, University of Prince Edward Island, Charlottetown, PE C1A 4P3, Canada.

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Rodolfo Nino-FongComparative Orthopaedic Research Laboratory, Department of Health Management, Atlantic Veterinary College, University of Prince Edward Island, Charlottetown, PE C1A 4P3, Canada.

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Abstract

Objective—To isolate and characterize mesenchymal stem cells (MSCs) from canine muscle and periosteum and compare proliferative capacities of bone marrow-, adipose tissue-, muscle-, and periosteum-derived MSCs (BMSCs, AMSCs, MMSCs, and PMSCs, respectively).

Sample—7 canine cadavers.

Procedures—MSCs were characterized on the basis of morphology, immunofluorescence of MSC-associated cell surface markers, and expression of pluripotency-associated transcription factors. Morphological and histochemical methods were used to evaluate differentiation of MSCs cultured in adipogenic, osteogenic, and chondrogenic media. Messenger ribonucleic acid expression of alkaline phosphatase, RUNX2, OSTERIX, and OSTEOPONTIN were evaluated as markers for osteogenic differentiation. Passage-1 MSCs were counted at 24, 48, 72, and 96 hours to determine tissue-specific MSC proliferative capacity. Mesenchymal stem cell yield per gram of tissue was calculated for confluent passage-1 MSCs.

Results—Successful isolation of BMSCs, AMSCs, MMSCs, and PMSCs was determined on the basis of morphology; expression of CD44 and CD90; no expression of CD34 and CD45; mRNA expression of SOX2, OCT4, and NANOG; and adipogenic and osteogenic differentiation. Proliferative capacity was not significantly different among BMSCs, AMSCs, MMSCs, and PMSCs over a 4-day culture period. Periosteum provided a significantly higher MSC yield per gram of tissue once confluent in passage 1 (mean ± SD of 19,400,000 ± 12,800,000 of PMSCs/g of periosteum obtained in a mean ± SD of 13 ± 1.64 days).

Conclusions and Clinical Relevance—Results indicated that canine muscle and periosteum may be sources of MSCs. Periosteum was a superior tissue source for MSC yield and may be useful in allogenic applications.

Abstract

Objective—To isolate and characterize mesenchymal stem cells (MSCs) from canine muscle and periosteum and compare proliferative capacities of bone marrow-, adipose tissue-, muscle-, and periosteum-derived MSCs (BMSCs, AMSCs, MMSCs, and PMSCs, respectively).

Sample—7 canine cadavers.

Procedures—MSCs were characterized on the basis of morphology, immunofluorescence of MSC-associated cell surface markers, and expression of pluripotency-associated transcription factors. Morphological and histochemical methods were used to evaluate differentiation of MSCs cultured in adipogenic, osteogenic, and chondrogenic media. Messenger ribonucleic acid expression of alkaline phosphatase, RUNX2, OSTERIX, and OSTEOPONTIN were evaluated as markers for osteogenic differentiation. Passage-1 MSCs were counted at 24, 48, 72, and 96 hours to determine tissue-specific MSC proliferative capacity. Mesenchymal stem cell yield per gram of tissue was calculated for confluent passage-1 MSCs.

Results—Successful isolation of BMSCs, AMSCs, MMSCs, and PMSCs was determined on the basis of morphology; expression of CD44 and CD90; no expression of CD34 and CD45; mRNA expression of SOX2, OCT4, and NANOG; and adipogenic and osteogenic differentiation. Proliferative capacity was not significantly different among BMSCs, AMSCs, MMSCs, and PMSCs over a 4-day culture period. Periosteum provided a significantly higher MSC yield per gram of tissue once confluent in passage 1 (mean ± SD of 19,400,000 ± 12,800,000 of PMSCs/g of periosteum obtained in a mean ± SD of 13 ± 1.64 days).

Conclusions and Clinical Relevance—Results indicated that canine muscle and periosteum may be sources of MSCs. Periosteum was a superior tissue source for MSC yield and may be useful in allogenic applications.

Despite the advances in veterinary surgery and medicine, treatment of many canine orthopedic and neurologic conditions often does not result in the desired clinical outcome or patient's return to normal function. Mesenchymal stem cells have shown tremendous promise in experimental and clinical studies of veterinary and human diseases.17 Little has been published regarding MSCs in dogs; however, the therapeutic applications that have been reported have contributed to our basic understanding of canine MSCs.

Dogs affected with bilateral coxofemoral osteoarthritis had considerable improvement in lameness and fewer signs of pain on range of motion after receiving intra-articular injections of autologous AMSCs.8 This was in contrast to a placebo group that failed to improve, as determined on the basis of veterinary and owner assessments.

In a canine disk degeneration experimental study,9 dogs that received injection of BMSCs into the degenerated disk 4 weeks after nucleotomy had a significantly higher disk height index radiographically, stronger disk signal intensity on T2-weighted MRI images, significantly higher proteoglycan content, and greater Fas ligand expression, compared with the nucleotomy-only group. Macroscopically, the treatment group looked similar to the control group and lacked the narrowed disk space and connective tissue invasion seen in the nucleotomy group. The results support the notion that the injected MSCs decelerated the effects of disk degeneration.

Bruder et al10 evaluated the effect of cultured autologous BMSCs on the healing of a critical-sized defect in canine femurs. Substantial new bone formation was present in the group treated with a carrier loaded with MSCs; atrophic nonunion was present in all untreated femurs at 16 weeks. Because MSCs are involved in all 4 strategies used for bone regeneration (osteogenesis, osteoinduction, osteoconduction, and osteopromotion), their role in bone tissue engineering is an area of clinical interest.7

There are many properties of MSCs that make their use an attractive option for clinical applications. They can be effectively isolated and expanded with high efficiency11 Mesenchymal stem cells can be cryopreserved and still maintain their viability and be induced to differentiate along multiple lineages.7 Mesenchymal stem cells are considered immunoprivileged. This is attributed to their lack of expression of several surface antigens important for T- and B-cell recognition and also their capability to suppress lymphocytes.6 This property may lend support to the use of allogenic MSCs, which may be banked for future use as needed.12 Mesenchymal stem cells also have the property of homing, meaning that they can migrate to sites of injury when given systemically.11 The homing mechanism is attributed to the multiple growth factor, cytokine, and CD44 receptors present on MSCs and their inherent ability to attach to injured endothelium.13 Mesenchymal stem cells were once thought to provide a therapeutic effect only by differentiating into a specific cell lineage, like an osteoblast, but it is likely that paracrine effects have a major role in these observations.13 The cells are reported to secrete a multitude of biologically active molecules, including interleukins 6, 7, 8, 11, 12, 14, and 15; basic fibroblast growth factor; vascular endothelial growth factor; and platelet-derived growth factor.14 Cell-based treatments exploiting these properties have been applied in various fields, including neurology, cardiology, myology, nephrology, and orthopedics.11,15,16

Despite the positive results of clinical and experimental MSC studies in animals, there are many unknown factors associated with their use. For example, in the literature on canine MSCs, the ideal cell transplantation number, cell yield per gram of donor tissue, and ideal tissue source are not definitively known. Conventionally, high numbers of MSCs have been used in veterinary and human cell-based treatments.1,17,18 Wagner et al19 reported that in human medicine, 1 to 5 million cells/kg are administered IV or directly into the tissue. To obtain cell numbers of this magnitude, culture expansion would be required. Superior tissue sources have been identified in a variety of species and depend on the intended use of the MSCs.20–25 Rat BMSCs and PMSCs had superior osteogenic potential in comparison with AMSCs on the basis of significantly greater in vitro mineralized matrix formation and in vivo new bone formation.22 Canine AMSCs had reduced osteogenic capability in contrast to BMSCs under hypoxic conditions.23 Mesenchymal stem cells obtained from human synovial tissue had superior chondrogenic and osteogenic potential in comparison with BMSCs, AMSCs, MMSCs, and PMSCs.20

It is clear that further basic knowledge surrounding canine MSCs is required and is justified on the basis of their potential therapeutic applications in veterinary medicine. Identifying potential donor tissue sources of canine MSCs, characterizing their phenotype, and determining the postexpansion cell yield per gram of donor tissue would contribute to our understanding of canine MSCs. The first objective of the study reported here was to isolate and characterize canine MMSCs and PMSCs. The second objective was to compare the proliferation potential of MSCs from these 2 potential donor tissue sources with 2 conventional canine sources: bone marrow and adipose tissue. We hypothesized that canine muscle and periosteum would be sources of multipotent MSCs. On the basis of the proliferation potential of equine MSCs observed in our laboratory, we hypothesized that canine MMSCs and PMSCs would have a proliferation potential equivalent, if not superior, to that of BMSCs and AMSCs.

Materials and Methods

Bone marrow, adipose tissue, muscle, and periosteum were collected from 7 randomly selected dogs that were euthanized at local animal shelters as part of a population control program. All experimental protocols were reviewed and approved by the Institutional Animal Care Committee. The dogs were young adult mixed-breed dogs weighing from 20 to 35 kg. Four dogs were sexually intact males, 1 was a neutered male, and 2 were females of unknown ovariohysterectomy status. Tissues were collected immediately after euthanasia in an aseptic manner.

Isolation and culture of MSCs from bone marrow—Bone marrow was harvested from the proximal aspect of both humeri in 6 dogs and from 1 humerus in another dog by use of a 15-gauge Illinois bone marrow biopsy needlea with 2,500 U of heparinb in a 12-mL collecting syringe. The maximal amount of bone marrow that could be harvested by use of this technique (5 to 27.5 mL) was collected and suspended in αMEM.c Marrow samples were kept on ice for immediate transport to the on-site laboratory. The samples were divided in half and placed in 50-mL centrifuge tubesd and centrifuged at 1,500 × g for 10 minutes. The buffy coat was collected and placed in a flaske with 10 mL of standard medium, which was composed of αMEM supplemented with 10% FBS,f l-glutamineg (2mM), 10,000 U of penicillin and 10 mg of streptomycin/mL,h and 250 μg of amphotericin Bi/mL. Cell cultures were maintained in a humidified 5% carbon dioxide and 95% air atmosphere incubator at 37°C. Unattached cells were removed after 48 hours by washing with sterile PBS solution.j The medium was renewed 3 times/wk. The cells were cultured until they reached > 75% confluency or were present in culture for 11 days. At that time, the cells were detached, counted, and subcultured or cryopreserved for further studies by use of cryopreservation medium (90% FBS and 10% dimethyl sulfoxidek).

Collection and cryopreservation of adipose tissue, bone, muscle, and periosteum—Adipose tissue was collected from the dorsum of the sacrococcygeal region. A 3-cm skin incision was made over the tail base. The underlying subcutaneous tissue was excised and collected from each dog and placed in αMEM solution and kept on ice. The muscle and periosteum were collected from the hind limb; the tissues were collected from both hind limbs in a select number of dogs. A standard craniomedial approach was made to the stifle joint and extended distally over the tibia. The cranial tibial muscle (15.30 to 42.26 g) was excised, cut into smaller pieces, and placed in chilled αMEM solution on ice. The periosteum was incised on the medial aspect of the proximal portion of the tibia and elevated by use of a periosteal elevator. Approximately 0.49 to 1.60 g of periosteum was collected from each dog and placed in chilled αMEM solution. Cortical bone was collected from the proximal portion of the tibia of 1 dog by use of Lempert rongeurs and placed in αMEM solution similar to the other tissues. The bone was used as a control for the RT-PCR experiments evaluating osteogenic differentiation. Tissues were processed within 24 hours; tissues that were not processed immediately were kept on ice and refrigerated at 4°C. Cold, sterile PBS solution was placed in Petri dishes to keep a moist environment for the tissues as they were cut into 1-cm segments. Tissue segments were placed into 2-mL cryovialsl and submerged in freezing medium composed of 92.5% PBS solution and 7.5% dimethyl sulfoxide. The cryovials remained at room temperature (20°C) for 30 minutes to allow for freezing medium to penetrate the tissue. The samples were then placed in closed-cell extruded polystyrene foam containers and put in a −80°C freezer for a minimum of 24 hours. All the samples were placed into a liquid nitrogen tank within 72 hours after processing.

Isolation and culture of MSCs from adipose tissue, muscle, and periosteum—Cells were isolated from tissues by use of an enzyme digestion technique. Cryopreserved adipose and muscle tissues were warmed in a water bath at 37°C for approximately 5 minutes until the liquid was defrosted. Tissue handling was performed with sterile technique in a biosafety cabinet. Each tissue was removed from the cryovials and placed in a 50-mL centrifuge tube containing 25 mL of sterile PBS solution. The tissue was rinsed with PBS solution, weighed, and minced. Minced tissue was placed in centrifuge tubes containing 10 mL of collagenase type Im (2,000 U/mL), vortexed, and placed in a 37°C incubator. The tubes were vortexed every 20 minutes for 60 minutes. Once the tissue was digested, 10 mL of standard medium was added to the mixture to inhibit further enzyme digestion. The cell suspension was filtered through a 100-μm filtern followed by a 70-μm filtern and centrifuged at 377 × g for 10 minutes. The supernatant was removed, and the cell pellet was resuspended in standard medium. Viable cell numbers, determined on the basis of 0.4% Trypan blueo dye exclusion, were counted with a hemocytometer. Adipose tissue cells were plated at a mean ± SD cell density of 2.2 × 104 cells/cm2 ± 0.84 × 104 cells/cm2, and muscle cells were plated at a mean ± SD cell density of 3.6 × 104 cells/cm2 ± 1.2 × 104 cells/cm2 with standard medium. The cell cultures were maintained in a humidified 5% carbon dioxide and 95% air atmosphere incubator at 37°C.

Isolation of cells from the periosteum was similar to that of adipose tissue and muscle except that the minced tissue was pretreated with type I collagenase (2,000 U/mL) for 10 minutes. The partially digested tissue was rinsed and treated with type I collagenase (2,000 U/mL) for an additional 160 minutes. Viable periosteal cells were plated at a mean ± SD cell density of 3.2 × 104 cells/cm2 ± 2.2 × 104 cells/cm2 with standard medium.

The medium was renewed 3 times/wk. The cells were grown to 75% to 100% confluency, at which time they were detached, counted, and subcultured or cryopreserved for further studies.

Isolation of bone cells for use in RT-PCR assay experiments evaluating osteogenic differentiation—Bone cells were isolated and cultured from cryopreserved cortical bone in a similar manner to adipose tissue and muscle. However, the tissue was enzyme digested for 5 hours with agitation every 30 minutes. Bone cells were maintained in standard culture medium until confluent for use as a control in the evaluation of osteogenic transcription factors.

Evaluation of cell surface markers and genes associated with pluripotency—Cells derived from bone marrow, adipose tissue, muscle, and periosteum from 3 dogs were expanded in standard medium to 80% to 100% confluency in passage 1 for use in evaluation of cell surface CD protein markers with fluorescent-labeled specific antibodies. These cells were also separately evaluated for the expression of pluripotency gene markers by use of PCR assay.

Immunofluorescence of CD cell surface markers—Isolated canine MSCs were evaluated via immunofluorescence microscopy for MSC-specific markers CD90 and CD44 and hematopoietic stem cell markers CD45, CD34, and CD146.26,27 Cells were seeded at a cell density of 1,500 cells/cm2 in a 24-well platep for 48 hours. Cells were fixed with 4% paraformaldehydeq for 15 minutes at room temperature. Nonspecific binding was blocked with 1% bovine serum albuminr for 1 hour at room temperature followed by the addition of diluted primary antibodies in a dark environment. Cells were incubated with CD34s (1:1,000), CD44t (1:1,000), CD45s (1:1,000), CD90u (1:1,000), and CD 146v (2:1,000) overnight at 4°C. Controls included wells that did not have added antibody. All primary antibodies were labeled with fluorescein isothiocyanate isomer 1 except the CD90 antibody. A secondary goat anti-mouse IgMw antibody, tagged with fluorescein isothiocyanate isomer 1, was used to detect the CD90 antibody (1:1,000). After the addition of the secondary antibody, cells were incubated for 1 hour at room temperature. The cells were washed and counterstained with Hoechst 33258x (pentahydrate bis-benzamide dye solution; 1 μL in 10 mL of distilled water) for 5 minutes at room temperature in a dark environment. Photomicrographs were obtained with a fluorescent microscope.

RT-PCR assay for genes associated with pluripotency—Cells were seeded at a cell density of 4,200 cells/cm2 in a 6-well platep and cultured in standard medium until confluent. Total RNA was extracted from cells by use of a total RNA kity following manufacturer instructions. The RNA was treated with DNase to remove contaminating DNA. The cDNAs were synthesized from 1 μg of total RNA by use of a cDNA synthesis kit.z Primers derived from the coding regions of SOX2, NANOG, OCT4, the genes associated with pluripotency, and β2 microglobulin (a housekeeping gene) were summarized (Table 1).28 Twenty-five microliters of PCR reactions was prepared with 1.6 μg of cDNA, 0. 1μM of each primer, 12.5 μL of a 2× mix for real-time PCR applications,aa and sterile deionized water.

Table 1—

Primers used in an RT-PCR assay for mRNA expression of pluripotency- and osteogenic-related transcription factors associated with various genes in canine MSCs.

GenePrimer sequence (5′-3′)Amplicon size (bp)Annealing temperature (°C)
OCT4Forward: GAGTGAGAGGCAACCTGGAG27460
Reverse: GTGAAGTGAGGGCTCCCATA
NANOGForward: GAATAACCCGAATTGGAGCAG14160
Reverse: AGCGATTCCTCTTCACAGTTG
SOX2Forward: AGTCTCCAAGCGACGAAAAA14258
Reverse: GCAAGAAGCCTCTCCTTGAA
ALPForward: TCAACAGACCCTGAAATACGC20257
Reverse: TCTTGGAGAGGGCCACGTAAG
OSTEOPONTINForward: CATATGATGGCCGAGGTGATAG11460
Reverse: CAAGTGATGTGAAGTCCTCCTC
OSTERIXForward: ACGACACTGGGCAAAGCAG28560
Reverse: CATGTCCAGGGAGGTGTAGAC
RUNX2Forward: GTCTCCTTCCAGAATGCTTCC10062
Reverse: GGAACTGAGGATGAGGAGAC
β2 MICROGLOBULINForward: TCTACATTGGGCACTGTGTCAC13660
Reverse: AAGAGTTCAGGTCTGACCAAG

The PCR assay was run by use of a thermal cycler.bb Cycling conditions were as follows: 95°C for 5 minutes and 45 cycles at 95°C for 20 seconds, optimal annealing temperature (Table 1) for 20 seconds, and 72°C for 30 seconds. No template controls (nuclease-free water instead of cDNA) were used as a negative control. The PCR products were stained with DNA gel stain,cc separated on 1% agarose geldd by use of electrophoresis, and inspected under UV light. Digital images were captured via an imaging systemee with image acquisitionff and gel documentationgg software.

Differentiation of MSCs into adipogenic, chondrogenic, and osteogenic cell lineages—Cells in passage 1, derived from bone marrow, adipose tissue, muscle, and periosteum from 3 dogs, were used for adipogenic, chondrogenic, and osteogenic differentiation assays. All medium changes were performed 3 times/wk.

Adipogenesis—Cells were plated at a seeding density of 40,000 cells/cm2 in 12-well platesp and supplemented with an adipogenic medium consisting of isobutyl-methylxanthinehh (0.5 mmol/L), rosiglitazoneii (5 μmol/L), dexamethasonejj (1 μmol/L), biotinkk (33 μmol/L), insulinll (1 μmol/L), pantothenatemm (17 μmol/L), 10,000 U of penicillin and 10 mg of streptomycin/mL, amphotericin B (250 μg/mL), and l-glutamine (2mM) in D-minimal essential mediumnn with 3% FBS and 5% rabbit serumoo for 21 days. Equal numbers of cells were plated as controls and cultured in standard medium containing 5% FBS instead of 10%. Oil red O staining was used in histochemical and morphological evaluation.

Chondrogenesis—For chondrogenic differentiation, a pellet culture technique was used.29 Five hundred thousand cells were placed in 15-mL polypropylene tubes,pp in duplicate, and centrifuged at 377 × g for 5 minutes to achieve micromass pellets. The pellets were cultured for 28 days in chondrogenic medium similar to that of Csaki et al30 (D-minimal essential medium supplemented with dexamethasone [10−-7M], liquid media supplementqq [culture supplement containing bovine insulin, transferrin, selenous acid, linoleic acid, and bovine serum albumin], ascorbic acid 2-phosphaterr [50 μg/mL], l-glutamine [2mM], 10,000 U of penicillin and 10 mg of streptomycin/mL, and amphotericin B [250 μg/mL]) with and without hTGF-β1ss (10 ng/mL) in 2 dogs. The same pellet culture technique was used in the third dog except the pellets were only maintained in culture for 21 days. Pellets were not cultured in a standard medium as a control equivalent because of the experience our laboratory has had with cell loss during the culture period. Vidal et al29 reported a similar experience and hypothesized that cells cultured by use of a micromass pellet technique in standard media lacked a compact structure, resulting in cell loss with medium changes. For microscopic evaluation, the pellets were embedded in paraffin, cut into 5-μm-thick sections, and stained with H&E, Alcian blue, and Safranin O.

Osteogenesis—Cells were plated at a seeding density of 10,000 cells/cm2 in 12-well plates and supplemented with osteogenic medium (αMEM supplemented with 5% FBS, dexamethasone [10−-8M], β-glycerophosphatett [10mM], ascorbic acid-2 phosphate [50 μg/mL], l-glutamine [2mM], 10,000 U of penicillin and 10 mg of streptomycin/mL, and amphotericin B [250 μg/mL]) for 8 weeks. Equal numbers of cells were plated as controls and cultured in standard medium containing 5% FBS instead of 10% FBS. Von Kossa and Alizarin red staining were used for histochemical and morphological evaluation of osteogenic differentiation.

The ability to differentiate BMSCs, AMSCs, MMSCs, and PMSCs down the osteogenic lineage was also evaluated via gene expression of osteoblast markers ALP, RUNX2, OSTERIX, and OSTEOPONTIN by use of RT-PCR assay.26,28,31 Cells obtained from each of the donor tissues of 3 dogs were plated at a seeding density of 52,000 cells/cm2 in 6-well plates and cultured in an osteogenic medium for 4, 6, and 8 weeks. Equal numbers of cells were plated in the nontreatment group and cultured in standard medium containing 5% FBS instead of 10% for 4 weeks. Canine cortical bone cells were cultured in 5% standard medium for 8 weeks. Ribonucleic acid samples were obtained at 4, 6, and 8 weeks from the treatment group and the canine cortical bone cells and at 4 weeks from the nontreatment group; the cells were treated in a similar manner to the RNA extracted for evaluation of genes associated with pluripotency. The primer sequences, amplicon sizes, and annealing temperatures were summarized (Table 1). Cycling conditions were as follows: 95°C for 5 minutes and 40 cycles at 94°C for 20 seconds, optimal annealing temperature for 20 seconds, and 72°C for 30 seconds. The PCR products were run on 1% agarose gel as described.

Proliferation potential of canine MSCs—Passage 1 MSCs derived from bone marrow, adipose tissue, muscle, and periosteum from 7 dogs were cultured at a seeding density of 3,100 cells/cm2 in 6-well dishes containing standard medium. The cells were plated in triplicate for 24, 48, 72, and 96 hours. At each time point, the cells were washed with PBS solution and trypsinized with 0.05% trypsinuu for 30 minutes. The reaction was stopped with the addition of standard medium. Viable cells, as determined with use of 0.04% Trypan blue, were counted by use of a hemocytometer. The culture medium was changed every 2 days.

Postexpansion MSC yield per gram of donor tissue—Data were collected from the 7 dogs to determine the mean MSC yield obtained per gram of tissue of bone marrow, adipose tissue, muscle, and periosteum after the cells were grown to 80% to 100% confluency in passage 1. The volume of bone marrow and the weight of the 3 other tissue sources were recorded during the isolation and culture procedures. However, to statistically compare all 4 donor tissue sources, the volume of bone marrow required conversion to a unit of weight. We justified a 1:1 (wt/vol) conversion factor in a 3-step process. First, a thorough search of the literature did not identify the mass density of canine bone marrow. However, it is reported that human bone marrow has a mass density of 0.98 to 1.03 g/mL.32 Our laboratory also confirmed that equine bone marrow has a mass density of 1 g/mL and that canine blood has a mass density of 1 g/mL. Although not ideal, assuming that canine bone marrow has a similar composition to canine blood and that canine bone marrow may have a similar mass density to human and equine bone marrow, we considered the conversion factor to be reasonable. Thus, quantity of bone marrow was reported in grams.

Statistical analysis—Significant differences among the mean of the natural logarithm from each triplicate MSC count among the 4 tissue-derived MSCs were determined with a linear mixed model33 (ie, a 3-level random intercept model). The tissue and time effects were considered to be fixed effects. The contribution of each dog to the log-transformed MSC count was considered a random effect. The dependence of tissue on dog was considered a random effect. Analyses were performed with statistical software.vv Significant (P < 0.05) differences among the square root of MSC yield per gram of tissue were determined with a general linear model. A Bonferroni procedure was used to adjust for multiple tissue comparisons.34 Statistical and process management softwareww was used to perform this statistical analysis. Significance was set at P < 0.05.

Results

Characterization of canine MSCs derived from bone marrow, adipose tissue, muscle, and periosteum—Plastic-adherent cells with the typical fibroblastic phenotype35 were isolated and expanded from all 4 donor tissues from all 7 dogs. Most colonies derived from 3 of the 4 tissue sources (bone marrow, adipose tissue, and periosteum) became 80% to 100% confluent within 6 to 8 days of initial seeding of flasks (passage zero). However, MMSCs consistently took longer and achieved only 45% to 75% confluency during this time frame. In passage 1, the colonies from all tissues reached 80% to 100% confluency within 11 to 20 days (mean, 16 days).

Cell surface CD markers—Cells isolated from all 4 donor tissues and cultured under standard conditions in passage 1 strongly expressed the cell surface antigen CD90 and weakly expressed CD44, as determined on the basis of positive staining with immunofluorescence. Bone marrow–derived MSCs from 2 of 3 dogs had weak positive expression of CD45; the other 3 tissue-derived MSCs were negatively labeled for this cell surface marker. None of the isolated cells stained positively for CD34 and CD146 (Figure 1).

Figure 1—
Figure 1—

Photomicrographs of immunofluorescence labeling of canine PMSCs. Notice strong staining for stem cell surface–specific marker CD90 and lack of staining for hematopoietic stem cell surface markers CD34 and CD45 and for CD146. Immunofluorescent stain; bar = 100 μm.

Citation: American Journal of Veterinary Research 73, 8; 10.2460/ajvr.73.8.1305

Expression of pluripotency-associated transcription factors—Bone marrow, adipose tissue, muscle, and periosteum-derived MSCs positively expressed the pluripotency-associated transcription factors SOX2, OCT4, and NANOG, as determined via RT-PCR analysis and gel electrophoresis (Figure 2).

Figure 2—
Figure 2—

Results of RT-PCR analysis and gel electrophoresis for mRNA expression of pluripotency-associated transcription factors SOX2, OCT4, and NANOG and the housekeeping gene β2 MICROGLOBULIN in each of 4 canine tissue–derived MSCs (BMSCs, AMSCs, MMSCs, and PMSCs). The 4 bands above each transcription factor label correspond (from left to right) with MSCs derived from bone marrow, adipose tissue, muscle, and periosteum, respectively. The DNA ladders are labeled L, and each band corresponds to 100 bp, with the lowest band representing 100 bp.

Citation: American Journal of Veterinary Research 73, 8; 10.2460/ajvr.73.8.1305

MSC trilineage differentiation—Bone marrow–derived MSCs, AMSCs, MMSCs, and PMSCs were successfully differentiated down the adipogenic and osteogenic cell lineages. Chondrogenic differentiation of MSCs derived from these 4 canine donor tissue sources was not detected.

The cells cultured in an adipogenic differentiation medium for 21 days had positive oil red O staining of lipid droplets (Figure 3). The number of lipid droplets and intensity of staining were most impressive with MMSCs. Cells cultured in standard medium did not develop lipid droplets or have positive staining with oil red O.

Figure 3—
Figure 3—

Photomicrographs of canine BMSCs, AMSCs, MMSCs, and PMSCs cultured in adipogenic or standard medium for 21 days. Notice staining of lipids in MSCs cultured in adipogenic medium and lack of staining in MSCs cultured in standard medium. Oil red O stain; bar = 40 μm.

Citation: American Journal of Veterinary Research 73, 8; 10.2460/ajvr.73.8.1305

Bone marrow–derived MSCs, AMSCs, MMSCs, and PMSCs from 3 dogs were set up in a pellet culture system in an attempt to induce chondrogenic differentiation. The suspended cells formed a visible white opaque pellet within 2 days. Cellular morphology and staining characteristics of each of the recovered pellets were all similar regardless of tissue source, time in culture, or whether the culture was supplemented with hTGF-β1 (Figure 4). Routine H&E staining of each of the pellets revealed a large central zone composed of eosinophilic amorphous necrotic material admixed with sparse karyorrhectic debris (consistent with areas of tissue necrosis). The latter areas were surrounded by a thin peripheral layer (thickness, approx 1 to 10 cell layers) comprised of bland, uniform, streaming, spindloid mesenchymal cells. These cells had fusiform nuclei; dense, fine chromatin; unapparent to small nucleoli; and small amounts of poorly defined pinkish blue cytoplasm. Spindloid cells were separated by small amounts of pale, eosinophilic matrix that appeared blue when stained with Alcian blue but that did not stain appreciably with Safranin O. There was no evidence of chondrocytes, lacunae, or a chondroid matrix to support chondrogenic differentiation in any of the examined BMSC, AMSC, MMSC, and PMSC samples.

Figure 4—
Figure 4—

Photomicrographs of canine MMSCs cultured in chondrogenic medium with (+) and without (−) hTGF-β1 for 28 days. Notice that there is no appreciable difference in the cellular morphology or staining characteristics between the 2 culture conditions and that there is no evidence of chondrogenic differentiation. A large central area of tissue necrosis with a mesenchymal population in the periphery of each photomicrograph is visible. H&E, Alcian blue, and Safranin O stains; bar = 100 μm.

Citation: American Journal of Veterinary Research 73, 8; 10.2460/ajvr.73.8.1305

Osteogenic differentiation was detected in all 4 donor tissue–derived MSCs in 3 dogs after induction with an osteogenic medium for 8 weeks (Figures 5 and 6). The morphology of the MSCs progressed from a fibroblast-like appearance to a polygonal shape followed by the formation of nodular aggregates. These nodular aggregates stained with Von Kossa and Alizarin red, revealing the presence of a phosphate mineral composition and the presence of calcium salts, respectively. The stain uptake of the nodules by use of both techniques helped confirm the presence of mineralization and differentiation of the MSCs down the osteogenic cell lineage. Mesenchymal stem cells cultured in standard medium did not form nodules and did not stain with Von Kossa or Alizarin red stain.

Figure 5—
Figure 5—

Photomicrographs of canine BMSCs, AMSCs, MMSCs, and PMSCs cultured for 8 weeks in osteogenic or standard medium. Notice stained nodules in MSCs cultured in osteogenic medium and no staining in MSCs cultured in standard medium. Von Kossa stain; bar = 40 μm.

Citation: American Journal of Veterinary Research 73, 8; 10.2460/ajvr.73.8.1305

Figure 6—
Figure 6—

Photomicrographs of canine BMSCs, AMSCs, MMSCs, and PMSCs cultured for 8 weeks in osteogenic or standard medium. Notice stained nodules in MSCs cultured in osteogenic medium and no staining in MSCs cultured in standard medium. Alizarin red stain; bar = 40 μm.

Citation: American Journal of Veterinary Research 73, 8; 10.2460/ajvr.73.8.1305

Alkaline phosphatase, RUNX2, OSTERIX, and OSTEOPONTIN gene expression was evident in the osteoinduced BMSCs, AMSCs, MMSCs, and PMSCs at 4, 6, and 8 weeks on the basis of visualization of the PCR products via gel electrophoresis. Canine bone cells, which were used as a control, had similar expression of each of the transcription factors at 4, 6, and 8 weeks when cultured in standard medium. Basal expression was not present in the MSCs cultured in standard medium at 4 weeks (Figure 7). A nonspecific PCR product in the OSTEOPONTIN run was evident for BMSCs cultured in standard medium from only 1 dog; however, the amplicon size was not consistent with this transcription factor.

Figure 7—
Figure 7—

Results of RT-PCR analysis and gel electrophoresis for mRNA expression of osteogenic transcription factors ALP (b), RUNX2 (c), OSTERIX (d), and OSTEOPONTIN (e) in canine MSCs derived from bone marrow (B), adipose tissue (A), muscle (M), and periosteum (P) cultured in osteogenic medium for 4 weeks (OM4), 6 weeks (OM6), and 8 weeks (OM8). Notice that basal expression of these transcription factors was not seen in any of the 4 tissue-derived sources of MSCs cultured in standard medium, for 4 weeks (SM4). Bone marrow–derived MSCs cultured in SM4 had nonspecific gene expression that was inconsistent with OSTEOPONTIN. The housekeeping gene β2-microglobulin (a) was expressed in all tissue-derived MSCs regardless of culture conditions. The positive control tissue (c), cortical bone cultured in SM4, reveals mRNA expression of ALP, RUNX2, OSTERIX, and OSTEOPONTIN as expected. See Figure 2 for remainder of key.

Citation: American Journal of Veterinary Research 73, 8; 10.2460/ajvr.73.8.1305

Proliferation potential of canine MSCs—Mean ± SD log-transformed MSC counts for BMSCs, AMSCs, MMSCs, and PMSCs at each time point (24, 48, 72, and 96 hours) of the proliferation assay were summarized (Table 2). Mean of the log-transformed MSC counts was higher for MMSCs than for BMSCs, AMSCs, and PMSCs at each of the time points. The SD of the log-transformed MSC counts was higher for BMSCs than for AMSCs, MMSCs, and PMSCs at all time points. The SD of the log-transformed MSC counts was the lowest for AMSCs at 48, 72, and 96 hours. Mean of the log-transformed MSC counts for BMSCs, AMSCs, MMSCs, and PMSCs increased at each time point.

Table 2—

Mean ± SD values of log-transformed MSC counts obtained from canine BMSCs, AMSCs, MMSCs, and PMSCs at 24, 48, 72, and 96 hours of proliferation assay 1.

MSC24 hours48 hours72 hours96 hours
BMSC2.69 ± 0.633.58 ± 1.023.82 ± 0.924.69 ± 1.09
AMSC2.76 ± 0.503.15 ± 0.654.17 ± 0.354.76 ± 0.33
MMSC3.22 ± 0.453.60 ± 0.834.63 ± 0.625.30 ± 0.71
PMSC2.73 ± 0.593.22 ± 0.694.31 ± 0.644.90 ± 0.48

Time had a significant (P < 0.001) effect on the mean of the log-transformed MSC counts for BMSCs, AMSCs, MMSCs, and PMSCs. There was no significant (P = 0.36) difference among the 4 donor tissue–derived MSCs for the mean of the log-transformed MSC counts. The estimated variance at the dog level was 0.08, and the variance at the residual level was 0.47.

Postexpansion MSC yield per gram of tissue—Mean ± SD of the MSC yield per gram of bone marrow, adipose tissue, muscle, and periosteum and the days for these cells to reach 80% to 100% confluency in passage 1 were summarized (Table 3). Mean of the square root of the MSC yield per gram of tissue for each of the tissues was determined. The data indicated that periosteum had the highest MSC yield per gram of tissue and bone marrow had the lowest MSC yield. Bone marrow–derived MSC counts had the highest SD for the mean number of days to reach 80% to 100% confluency in passage 1, whereas MMSC counts had the lowest SD.

Table 3—

Mean ± SD values of MSC yield per gram of tissue from canine BMSCs, AMSCs, MMSCs, and PMSCs and mean ± SD number of days for those cells to reach 80% to 100% confluency in passage 1.

MSCMSC yieldNo. of days
BMSC1,449,788 ± 1,198,60216.17 ± 4.17
AMSC2,334,463 ± 1,253,12613.86 ± 2.04
MMSC3,367,969 ± 2,088,82515.00 ± 0.58
PMSC19,400,000 ± 12,800,000*13.2 ± 1.64

Significantly (P < 0.05) different from other values in this column.

Results revealed that the effect of time for the cells to reach 80% to 100% confluency in passage 1 was not significantly (P = 0.740) different among BMSCs, AMSCs, MMSCs, and PMSCs. There was also no significant (P = 0.443) difference for the dog effect. Results revealed a significant (P < 0.001) difference among the means of the square root of MSC yield per gram of bone marrow, adipose tissue, muscle, and periosteum. Periosteum-derived MSCs provided the highest mean of the square root of the MSC yield per gram of periosteum, compared with the means of the square root of the MSC yield per gram of bone marrow, adipose tissue, and muscle.

Discussion

Mesenchymal stem cells were successfully isolated from canine bone marrow, adipose tissue, muscle, and periosteum. This was determined on the basis of their ability to adhere to plastic,36 characteristic morphology,35 positive expression of cell surface markers CD90 and CD44, negative expression of CD34 and CD45,37 and expression of pluripotency-associated transcription factors SOX2, OCT4, and NANOG.28 The isolated MSCs had bilineage potential and were differentiated into osteoblasts and adipocytes. Periosteum was a superior tissue source in providing the greatest number of MSCs per gram of tissue when the cells were grown to 80% to 100% confluency in passage 1. To the authors' knowledge, the present report is the first to describe the isolation and characterization of canine MMSCs and PMSCs.

The International Society for Cellular Therapy has provided a list of criteria to define human MSCs. This includes their ability to adhere to plastic; be positive for cell surface markers CD73, CD90, and CD105; be negative for cell surface markers CD 11b, CD14, CD19, CD29α, CD34, CD45, and HLA-DE; and be able to differentiate into osteoblasts, adipocytes, and chondroblasts.37 Mesenchymal stem cells have been isolated from humans, baboons, rabbits, pigs, rats, mice,38 sheep,39 horses,40 cows,41 dogs,28 and cats.42 Tissue sources include bone marrow, periosteum, adipose tissue, synovium, muscle,6,20,21 peripheral blood, and the CNS.30 There is a belief that MSCs are associated with pericytes so that any vascularized tissue could be a potential source of MSCs.13

Mesenchymal stem cells from other species are defined on the basis of similar criteria, but the literature indicates that species differences exist.12,14 No single definitive marker defines an MSC in vitro or in vivo.37,39,43 Therefore, MSC identification is on the basis of the presence of a number of specific cell surface markers and the absence of others. In the present study, MSCs derived from all 4 donor tissue sources had expression of surface markers CD90 and CD44, Thy-1 glycoprotein and hyaluronate receptor–adhesion molecules, respectively.36 These results are in agreement with those of other studies that evaluated MSCs from canine bone marrow15,30 and adipose tissue26 as well as equine44,45 and human-derived MSCs.20 Most MSCs had negative results for the hematopoietic receptors CD34 and CD45. Bone marrow–derived MSCs from 2 dogs stained weakly for CD45 by use of immunofluorescence. This finding could potentially be attributable to nonspecific staining as was postulated by Braun et al,45 who found equine AMSCs to have a weak signal for CD45 by use of flow cytometry but no genetic expression of the surface marker by use of RT-PCR assay. The expression of CD45 could also be a reflection of hematopoietic contaminants that can be present in the early passages of MSC cultures.14

All 4 donor tissue–derived MSCs had negative results for CD146, a transmembrane glycoprotein that was conventionally considered a marker for endothelial cells. The marker is not detected on hematopoietic cells.14,46 However, pericytes express CD146 and are considered to be precursors of MSCs.46 Sorrentino et al27 isolated human BMSCs that were positive for CD146, in addition to the more commonly expressed surface cell antigens CD90 and CD105. Species differences exist and may account for this finding; murine MSC expression of CD34 is not clearly defined but is absent from human and rat MSCs.14,38 Javazon et al38 suggest that inconsistencies in cell surface marker expression may exist because of differences in isolation methods and culture conditions, species origin, and tissue origin. The cell surface marker CD146 has not been commonly evaluated in veterinary species, and further study would be required to make any firm conclusions on the basis of our data. Although immunofluorescence microscopy was a satisfactory technique in qualitative evaluation of cell surface marker expression and its use is reported by others,30,44 additional assessments by use of flow cytometry and RT-PCR assay could provide quantitative measurements as well and would be considered superior.

OCT4, SOX2, and NANOG are pluripotency-associated markers expressed in embryonic stem cells.47 Expression of these stem cell–related transcription factors has been reported in canine AMSCs28 and rhesus monkey BMSCs.48 The present study found expression of OCT4, SOX2, and NANOG in canine BMSCs, AMSCs, MMSCs, and PMSCs. The expression of these transcription factors may have a role in the molecular mechanisms governing MSC multipotency and proliferation potential.

Mesenchymal stem cells are further classified on the basis of their ability to differentiate down multiple lineages. Adipogenic, osteogenic, and chondrogenic differentiation are most commonly reported.28,30,44,45 However, some authors preferred to substitute myogenic26,43 or neurogenic49 differentiation for chondrogenic differentiation; others documented only bilineage differentiation.35 In the present study, adipogenic differentiation was confirmed by the appearance of positively stained lipid vacuoles within cells subjected to an adipogenic medium, as reported with canine BMSCs and AMSCs.26,30 The canine MMSCs appeared to have the greatest adipogenic potential on the basis of a subjectively greater number of positively stained lipid vacuoles, compared with the other tissue-derived MSCs. This is in contrast to the study by Yoshimura et al,21 who found that rat SMSCs and AMSCs had a greater adipogenic potential, compared with BMSCs, MMSCs, and PMSCs, on the basis of an objective assessment of oil red O staining–positive colony rate. Human SMSCs and AMSCs had a similar superiority to BMSCs, MMSCs, and PMSCs for adipogenic differentiation.20 However, the culture techniques differed among the 3 studies; canine MSCs in the present study underwent adipogenic induction for a period of 3 weeks, in comparison with 4 days for rat MSCs and 2 weeks for human MSCs.

All tissue-derived MSCs treated with an osteogenic medium differentiated into osteoblasts. This was supported by the morphological appearance of nodular aggregates that stained positively with Alizarin red and Von Kossa20,28,35,50 and the mRNA expression of osteoblast markers.26,28,51 Alkaline phosphatase, RUNX2, OSTERIX, and OSTEOPONTIN expression was found in all osteoinduced MSCs. Basal expression of these markers was not detected in MSCs treated with standard culture medium, confirming differentiation down the osteogenic lineage. Alkaline phosphatase and RUNX2 are early-stage transcription factors of osteogenesis, whereas OSTERIX is a late-stage transcription factor. OSTEOPONTIN, also known as bone sialoprotein 1, is expressed in other tissues but is considered an osteoblast-specific marker.28,51 Osteogenic differentiation and expression of osteogenic transcription factors have not been previously reported for canine MMSCs and PMSCs.

Our attempts at differentiating MSCs down the chondrogenic lineage were unsuccessful on the basis of morphological and histochemical assessments, despite use of a standard pellet culture system20,29,49,52–56 with an induction technique similar to that of Csaki et al.30 Others have had difficulty in obtaining cartilage tissue in a pellet culture and have reported similar central necrosis and undifferentiated cells.57 The uptake of Alcian blue stain may suggest that a proteoglycan matrix exists, but this was not supported by results of H&E and Safranin O staining or histologic appearance. Similar images to ours, which did not clearly reveal chondrocyte morphology, exist in the literature and are not convincing of chondrogenic differentiation, whereas other studies29,54,58,59 clearly reveal chondrogenic differentiation by use of histologic evaluation. The mRNA expression of chondrocyte markers collagen type II, aggrecan, and SOX9 would be useful to confirm differentiation down the chondrogenic lineage26 in future studies.

Technique differences could explain the chondrogenic differentiation results of the present study in comparison with successful pellet cultures. For instance, the present study used a higher initial cell plating number and a longer culture period than did Csaki et al.30 Even though successful differentiation accomplished by use of similar methodologies to that of the present study has been reported,21,45,49,59–61 each of these factors may have contributed to deficient nutrient diffusion and resultant cellular necrosis.29,57 Others speculate that isolated MSCs are incapable of chondrogenic differentiation because only a low percentage of an MSC population has the capacity to differentiate down ≥ 3 lineages.38,62 Many protocols for chondrogenic differentiation are not standardized, making comparison among studies difficult. Initial cell numbers, growth factors, time in culture, and plating techniques differ among studies.21,26,28–30,45,63 A 3-D culture system for chondrogenesis is superior to the monolayer technique.64 Micromass culture induces a larger amount of homogenized cartilage tissue with increased toluidine blue staining and expression of collagen type II, compared with pellet culture.57 Dexamethasone, fibroblast growth factor-2, TGF-β,56 and bone morphogenetic protein-253 also enhance MSC chondrogenesis in vitro. Although the present study used dexamethasone and hTGF-β in the culture media, successful differentiation may have occurred if the effects of fibroblast growth factor-2 or bone morphogenetic protein-2 had been exploited.

Pellets stained similarly regardless of whether hTGF-β1 was within the chondrogenic medium. This finding was in agreement with Neupane et al,28 who detected Alcian blue staining in treated and control micromass cultures of cells, but was not in agreement with the findings of another study30 that found that untreated pellets had reduced staining, compared with those induced with chondrogenic medium. Neupane et al28 concluded that MSC culture in a 3-D construct is sufficient to stimulate early chondrogenesis. However, addition of a growth factor from the TGF-β family is reported to be necessary for chondrogenic differentiation of MSCs. Improved chondrogenesis has been detected with addition of TGF-β2 and TGF-β3 in comparison with TGF-β1.11 In the present study, culture of canine BMSCs, AMSCs, MMSCs, and PMSCs in a 3-D construct, alone or with the addition of hTGF-β1, did not result in chondrogenic differentiation.

The effect of time on increasing cell numbers in culture was an expected finding in both the proliferation assay and the evaluation of postexpansion MSC yield per gram of tissue; it is a reflection of the inherent proliferative capacity of MSCs in vitro.20,29 In a comparison of the proliferative potential of MSCs in the proliferation assay, there was no significant difference in the proliferation rate between BMSCs, AMSCs, MMSCs, and PMSCs over the 4 days in culture in the present study. This was in agreement with results of another study21 that detected similar proliferative potential among rat AMSCs, MMSCs, and PMSCs for up to 10 days in culture when plated at 100 and 500 cells/cm2.

Canine periosteum provided the greatest postexpansion MSC yield per gram of tissue in the present study. Comparisons among MSCs obtained from multiple tissue sources are limited in the literature, but a report21 of human- and rat-derived MSCs exists; rat SMSCs had the greatest proliferative potential and the highest cell yield, compared with that of BMSCs, AMSCs, MMSCs, and PMSCs. Human AMSCs had the lowest proliferative capacity in comparison with BMSCs, MMSCs, PMSCs, and SMSCs.20 The author noted that even though human AMSCs had the lowest proliferative capacity, the final cell yield in passage 3 provided 109 cells and would suffice for current therapeutic and experimental requirements.

Mesenchymal stem cell yield per gram of tissue is infrequently reported. With a mean of 16 days in culture, the present study obtained a mean of 1.45 × 106 BMSCs/g of bone marrow, 2.33 × 106 AMSCs/g of adipose tissue, 3.37 × 106 MMSCs/g of muscle, and 19.40 × 106 PMSCs/g of periosteum. In another study,28 culture of canine AMSCs resulted in a yield of 0.53 × 106 AMSCs/g of tissue within 5 to 6 days. The differences in MSC yield are likely a factor of time in culture and passage number.20 Other factors reported to influence cell yield include culture media65 and cell doubling times.20

An ideal tissue source for use in regenerative treatments could be defined as a tissue that is in abundance, can be harvested with minimal invasiveness and morbidity to the patient, is economically advantageous, and provides a high number of effective MSCs within a short time. In the present study, tissues were collected in a manner that could easily be applied to clinical cases, again keeping in mind the definition of an ideal tissue source. Clinically, obtaining muscle and periosteum from dogs would be more invasive in comparison with obtaining adipose tissue and bone marrow, but if a niche demanded their availability, postmortem collection and storage in a bank for allogenic purposes may be a possibility. The benefits of allogenic MSCs are still controversial,66,67 but studies exist of the positive effects in models of canine myocardial infarction68 and spinal cord injury.67 Allogenic MSCs are an appealing option because they have the potential to avoid host immune rejection,12 have immunosuppressive capabilities,69 and can be obtained readily in large numbers without waiting the necessary time required for culture expansion of autogenous MSCs.

In the present study, canine skeletal muscle and periosteum were successfully used as sources of MSCs. Periosteum was a superior tissue source in providing the highest postexpansion MSC yield per gram of tissue within a clinically relevant time period. Isolation, characterization, and measurement of proliferative capacity of MSCs were performed from canine bone marrow, adipose tissue, muscle, and periosteum. The present study contributes to the basic understanding of canine MSCs, which ideally should be sought prior to their clinical application in veterinary regenerative treatments and tissue engineering. Their use may be enhanced by better understanding of the tissue sources available, the ideal tissue source, cell transplantation number, mode of administration, mechanisms of action, and long-term safety and efficacy. It is also important to be aware that information gathered from other species is meaningful but may not be applicable to all species.

ABBREVIATIONS

ALP

Alkaline phosphatase

αMEM

α-Minimal essential medium

AMSC

Adipose–derived mesenchymal stem cell

BMSC

Bone marrow-derived mesenchymal stem cell

FBS

Fetal bovine serum

hTGF

Human transforming growth factor

MMSC

Muscle-derived mesenchymal stem cell

MSC

Mesenchymal stem cell

PMSC

Periosteum-derived mesenchymal stem cell

RT-PCR

Reverse transcriptase PCR

SMSC

Synovium-derived mesenchymal stem cell

TGF

Transforming growth factor

a.

Illinois bone marrow biopsy needle, CareFusion, San Diego, Calif.

b.

Heparin, Leo Pharma Inc, Thornhill, ON, Canada.

c.

αMEM, Invitrogen, Toronto, ON, Canada.

d.

50-mL centrifuge tubes, BD Falcon, Franklin Lakes, NJ.

e.

T75 flask, Corning Inc, Corning, NY.

f.

FBS, PAA Laboratories Inc, Etobicoke, ON, Canada.

g.

l-glutamine, Invitrogen, Toronto, ON, Canada.

h.

Penicillin/Streptomycin, Sigma, Oakville, ON, Canada.

i.

Amphotericin B, Invitrogen, Toronto, ON, Canada.

j.

PBS, Invitrogen, Toronto, ON, Canada.

k.

Dimethyl sulfoxide, Sigma, Oakville, ON, Canada.

l.

Cryovials, Corning Inc, Corning, NY.

m.

Collagenase Type I, Invitrogen, Toronto, ON, Canada.

n.

70- and 100-μm filters, BD Falcon, Franklin Lakes, NJ.

o.

Trypan blue, Sigma, Oakville, ON, Canada.

p.

6-, 12-, 24-well plate, Corning Inc, Corning, NY.

q.

Paraformaldehyde, Fisher Scientific, Nepean, ON, Canada.

r.

Bovine Serum Albumin, Fisher Scientific, Nepean, ON, Canada.

s.

CD34, CD45, AbD Serotec, Raleigh, NC.

t.

CD44, Abcam Inc, Cambridge, Mass.

u.

CD90, Accurate Chemical and Scientific Corp, Westbury, NY.

v.

CD 146, Millipore, Billerica, Mass.

w.

Goat anti-mouse IgM antibody, Cedarlane Laboratories Ltd, Burlington, ON, Canada.

x.

Hoechst 33258, Invitrogen, Toronto, ON, Canada.

y.

Aurum Total RNA mini kit, Bio-Rad Laboratories, Hercules, Calif.

z.

iScript cDNA synthesis kit, Bio-Rad Laboratories, Hercules, Calif.

aa.

iQ SYBR Green Supermix, Bio-Rad Laboratories, Hercules, Calif.

bb.

Rotorgene-6 RG 3000, Corbett Research, Montreal, QC, Canada.

cc.

Syber Safe DNA gel stain, Invitrogen, Toronto, ON, Canada.

dd.

1% agarose gel, Biorad Laboratories Inc, Hercules, Calif.

ee.

BioSpectrum AC Image System, UVP LLC, Upland, Calif.

ff.

VisionWorks LS, UVP LLC, Upland, Calif.

gg.

GelDoc, UVP LLC, Upland, Calif.

hh.

Isobutyl-methylxanthine, Sigma, Oakville, ON, Canada.

ii.

Rosiglitazone, Toronto Research Chemicals, Toronto, ON, Canada.

jj.

Dexamethasone, Sigma, Oakville, ON, Canada.

kk.

Biotin, Sigma, Oakville, ON, Canada.

ll.

Insulin, Sigma, Oakville, ON, Canada.

mm.

Pantothenate, Sigma, Oakville, ON, Canada.

nn.

D-minimal essential medium, Sigma, Oakville, ON, Canada.

oo.

Rabbit serum, Invitrogen, Toronto, ON, Canada.

pp.

15-mL polypropylene tube, BD Falcon, Franklin Lakes, NJ.

qq.

ITS+1, Sigma, Oakville, ON, Canada.

rr.

Ascorbic acid 2-phosphate, Sigma, Oakville, ON, Canada.

ss.

hTGF-β1, Millipore, Billerica, Mass.

tt.

β-Glycerophosphate, Sigma, Oakville, ON, Canada.

uu.

Trypsin, Invitrogen, Toronto, ON, Canada.

vv.

STATA, version 10.0, StataCorp, College Station, Tex.

ww.

Minitab, version 16, Minitab Inc, State College, Pa.

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Contributor Notes

Supported by the Atlantic Veterinary College Internal Research Grant Fund and the Atlantic Canada Opportunities Agency.

The Atlantic Centre for Comparative Biomedical Research supplied laboratory equipment.

The authors thank Dr. Andrea Bourque for support with histologic samples and Dr. David Sims for technical assistance.

Address correspondence to Dr. Kisiel (agatha.kisiel@gmail.com).