• View in gallery

    Photographs of the typical appearance of monolayer tissue-like constructs formed in 75-cm2culture flasks by canine synoviocytes after 16 days of growth factor exposure (A) and after enzymatic release (B). The tissue was formed by synoviocytes harvested from a canine stifle joint with naturally occurring cranial cruciate ligament degeneration; the dog had been euthanatized for reasons unrelated to the study.

  • View in gallery

    Photographs illustrating the spontaneous formation of micromasses in a monolayer culture of osteoarthritic-joint synoviocytes harvested from a canine stifle joint that had undergone cranial cruciate ligament transection (Pond-Nuki method) 3 months earlier; the dog had been euthanatized for reasons unrelated to the study. A–-Image of a micromass in a 75-cm2 culture flask. B–-Image of 3 micromasses collected from culture medium after 16 days of sequential growth factor exposure.

  • View in gallery

    Photomicrographs of monolayer tissue constructs generated by non-osteoarthritic (normal)-joint synoviocytes (column A) from one dog (2 joints) and osteoarthritic-joint synoviocytes (column B) obtained from another dog (2 joints) that had undergone culture with sequential exposure to growth factors for 16 days and were stained with H&E stain (row 1), Masson trichrome stain (row 2), or toluidine blue (row 3). In row 2, blue fibrillar staining indicates the presence of collagen (thin arrows). In row 3, notice the lack of deep purple or indigo staining that indicates the presence of GAGs in the ECM. In all panels, bar = 50 μm.

  • View in gallery

    Photomicrographs of monolayer tissue constructs generated by normal-joint synoviocytes from one dog (2 joints [A and B]) and osteoarthritic-joint synoviocytes (C and D) obtained from another dog (2 joints) that had undergone culture with sequential exposure to growth factors for 16 days and were stained immunohistochemically for collagen I (A and C) or without the primary antibody (negative control samples; B and D). Notice the strong immunoreactivity to collagen I in the normal- and osteoarthritic-joint synoviocytes; the cells have a rounded phenotype (thick arrows) and fibrillar ECM (thin arrows). In panels A and C, immunohistochemical stain for collagen I, red chromagen, and Mayer hematoxylin stain; bar = 50 μm. In panels B and D, negative control immunohistochemical stain, red chromagen, and Mayer hematoxylin stain; bar = 50 μm.

  • View in gallery

    Photomicrographs of monolayer tissue constructs generated by normal-joint synoviocytes (A and B) from one dog (2 joints) and osteoarthritic-joint synoviocytes (C and D) obtained from another dog (2 joints) that had undergone culture with sequential exposure to growth factors for 16 days and were stained immunohistochemically for collagen II (A and C) or without the primary antibody (negative control samples; B and D). Notice the mild intracellular and pericellular immunoreactivity to collagen II in the normal- and osteoarthritic-joint synoviocytes; the cells have a rounded phenotype. In panels A and C, immunohistochemical stain for collagen II, red chromagen, and Mayer hematoxylin stain; bar = 50 μm. In panels B and D, negative control immunohistochemical stain, red chromagen, and Mayer hematoxylin stain; bar = 50 μm.

  • 1.

    Walmsley JR, Phillips TJ, Townsend HG. Meniscal tears in horses: an evaluation of clinical signs and arthroscopic treatment of 80 cases. Equine Vet J 2003; 35:402406.

    • Search Google Scholar
    • Export Citation
  • 2.

    Peroni JF, Stick JA. Evaluation of a cranial arthroscopic approach to the stifle joint for the treatment of femorotibial joint disease in horses: 23 cases (1998–1999). J Am Vet Med Assoc 2002; 220:10461052.

    • Search Google Scholar
    • Export Citation
  • 3.

    Jackson J, Vasseur PB, Griffey S, et al. Pathologic changes in grossly normal menisci in dogs with rupture of the cranial cruciate ligament. J Am Vet Med Assoc 2001; 218:12811284.

    • Search Google Scholar
    • Export Citation
  • 4.

    Ralphs SC, Whitney WO. Arthroscopic evaluation of menisci in dogs with cranial cruciate ligament injuries: 100 cases (1999–2000). J Am Vet Med Assoc 2002; 221:16011604.

    • Search Google Scholar
    • Export Citation
  • 5.

    Johnson KA, Francis DJ, Manley PA, et al. Comparison of the effects of caudal pole hemi-meniscectomy and complete medial meniscectomy in the canine stifle joint. Am J Vet Res 2004; 65:10531060.

    • Search Google Scholar
    • Export Citation
  • 6.

    Burks RT, Metcalf MH, Metcalf RW. Fifteen-year follow-up of arthroscopic partial meniscectomy. Arthroscopy 1997; 13:673679.

  • 7.

    Arnoczky SP, Warren RF. The microvasculature of the meniscus and its response to injury. An experimental study in the dog. Am J Sports Med 1983; 11:131141.

    • Search Google Scholar
    • Export Citation
  • 8.

    Kobayashi K, Fujimoto E, Deie M, et al. Regional differences in the healing potential of the meniscus—an organ culture model to eliminate the influence of microvasculature and the synovium. Knee 2004; 11:271278.

    • Search Google Scholar
    • Export Citation
  • 9.

    Arnoczky SP, Warren RF, Spivak JM. Meniscal repair using an exogenous fibrin clot. An experimental study in dogs. J Bone Joint Surg Am 1988; 70:12091217.

    • Search Google Scholar
    • Export Citation
  • 10.

    Okuda K, Ochi M, Shu N, et al. Meniscal rasping for repair of meniscal tear in the avascular zone. Arthroscopy 1999; 15:281286.

  • 11.

    Peretti GM, Gill TJ, Xu JW, et al. Cell-based therapy for meniscal repair: a large animal study. Am J Sports Med 2004; 32:146158.

  • 12.

    Klompmaker J, Veth RP, Jansen HW, et al. Meniscal repair by fibrocartilage in the dog: characterization of the repair tissue and the role of vascularity. Biomaterials 1996; 17:16851691.

    • Search Google Scholar
    • Export Citation
  • 13.

    Klompmaker J, Veth RP, Jansen HW, et al. Meniscal replacement using a porous polymer prosthesis: a preliminary study in the dog. Biomaterials 1996; 17:11691175.

    • Search Google Scholar
    • Export Citation
  • 14.

    de Groot JH, de Vrijer R, Pennings AJ, et al. Use of porous polyurethanes for meniscal reconstruction and meniscal prostheses. Biomaterials 1996; 17:163173.

    • Search Google Scholar
    • Export Citation
  • 15.

    Cook JL, Tomlinson JL, Kreeger JM, et al. Induction of meniscal regeneration in dogs using a novel biomaterial. Am J Sports Med 1999; 27:658665.

    • Search Google Scholar
    • Export Citation
  • 16.

    Stone KR, Steadman JR, Rodkey WG, et al. Regeneration of meniscal cartilage with use of a collagen scaffold. Analysis of preliminary data. J Bone Joint Surg Am 1997; 79:17701777.

    • Search Google Scholar
    • Export Citation
  • 17.

    Cox JS, Nye CE, Schaefer WW, et al. The degenerative effects of partial and total resection of the medial meniscus in dogs' knees. Clin Orthop Relat Res1975;178183.

    • Search Google Scholar
    • Export Citation
  • 18.

    Nishimura K, Solchaga LA, Caplan AI, et al. Chondroprogenitor cells of synovial tissue. Arthritis Rheum 1999; 42:26312637.

  • 19.

    Pei M, He F, Kish VL, et al. Engineering of functional cartilage tissue using stem cells from synovial lining: a preliminary study. Clin Orthop Relat Res 2008; 466:18801889.

    • Search Google Scholar
    • Export Citation
  • 20.

    Shintani N, Hunziker EB. Chondrogenic differentiation of bovine synovium: bone morphogenetic proteins 2 and 7 and transforming growth factor beta1 induce the formation of different types of cartilaginous tissue. Arthritis Rheum 2007; 56:18691879.

    • Search Google Scholar
    • Export Citation
  • 21.

    Yoshimura H, Muneta T, Nimura A, et al. Comparison of rat mesenchymal stem cells derived from bone marrow, synovium, periosteum, adipose tissue, and muscle. Cell Tissue Res 2007; 327:449462.

    • Search Google Scholar
    • Export Citation
  • 22.

    Park Y, Sugimoto M, Watrin A, et al. BMP-2 induces the expression of chondrocyte-specific genes in bovine synovium-derived progenitor cells cultured in three-dimensional alginate hydrogel. Osteoarthritis Cartilage 2005; 13:527536.

    • Search Google Scholar
    • Export Citation
  • 23.

    Arnoczky SP, Warren RF, Kaplan N. Meniscal remodeling following partial meniscectomy—an experimental study in the dog. Arthroscopy 1985; 1:247252.

    • Search Google Scholar
    • Export Citation
  • 24.

    Lindhorst E, Vail TP, Guilak F, et al. Longitudinal characterization of synovial fluid biomarkers in the canine meniscectomy model of osteoarthritis. J Orthop Res 2000; 18:269280.

    • Search Google Scholar
    • Export Citation
  • 25.

    Smith GN, Mickler EA, Albrecht ME, et al. Severity of medial meniscus damage in the canine knee after anterior cruciate ligament transection. Osteoarthritis Cartilage 2002; 10:321326.

    • Search Google Scholar
    • Export Citation
  • 26.

    van Tienen TG, Heijkants RG, de Groot JH, et al. Presence and mechanism of knee articular cartilage degeneration after meniscal reconstruction in dogs. Osteoarthritis Cartilage 2003; 11:7884.

    • Search Google Scholar
    • Export Citation
  • 27.

    Wyland DJ, Guilak F, Elliott DM, et al. Chondropathy after meniscal tear or partial meniscectomy in a canine model. J Orthop Res 2002; 20:9961002.

    • Search Google Scholar
    • Export Citation
  • 28.

    Ochi M, Ishida O, Daisaku H, et al. Immune response to fresh meniscal allografts in mice. J Surg Res 1995; 58:478484.

  • 29.

    Rodeo SA, Seneviratne A, Suzuki K, et al. Histological analysis of human meniscal allografts. A preliminary report. J Bone Joint Surg Am 2000; 82:10711082.

    • Search Google Scholar
    • Export Citation
  • 30.

    Pessina A, Bonomi A, Baglio C, et al. Microbiological risk assessment in stem cell manipulation. Crit Rev Microbiol 2008; 34:112.

  • 31.

    Graf KW Jr, Sekiya JK, Wojtys EM. Long-term results after combined medial meniscal allograft transplantation and anterior cruciate ligament reconstruction: minimum 8.5-year follow-up study. Arthroscopy 2004; 20:129140.

    • Search Google Scholar
    • Export Citation
  • 32.

    Jeffreys TE. Synovial chondromatosis. J Bone Joint Surg Br 1967; 49:530534.

  • 33.

    Blom AB, van Lent PL, Holthuysen AE, et al. Synovial lining macrophages mediate osteophyte formation during experimental osteoarthritis. Osteoarthritis Cartilage 2004; 12:627635.

    • Search Google Scholar
    • Export Citation
  • 34.

    van Lent PL, Blom AB, van der Kraan P, et al. Crucial role of synovial lining macrophages in the promotion of transforming growth factor β-mediated osteophyte formation. Arthritis Rheum 2004; 50:103111.

    • Search Google Scholar
    • Export Citation
  • 35.

    Giurea A, Ruger BM, Hollemann D, et al. STRO-1+ mesenchymal precursor cells located in synovial surface projections of patients with osteoarthritis. Osteoarthritis Cartilage 2006; 14:938943.

    • Search Google Scholar
    • Export Citation
  • 36.

    Mussener A, Funa K, Kleinau S, et al. Dynamic expression of transforming growth factor-betas (TGF-β) and their type I and type II receptors in the synovial tissue of arthritic rats. Clin Exp Immunol 1997; 107:112119.

    • Search Google Scholar
    • Export Citation
  • 37.

    Acosta CA, Izal I, Ripalda P, et al. Gene expression and proliferation analysis in young, aged, and osteoarthritic sheep chondrocytes effect of growth factor treatment. J Orthop Res 2006; 24:20872094.

    • Search Google Scholar
    • Export Citation
  • 38.

    Murphy JM, Dixon K, Beck S, et al. Reduced chondrogenic and adipogenic activity of mesenchymal stem cells from patients with advanced osteoarthritis. Arthritis Rheum 2002; 46:704713.

    • Search Google Scholar
    • Export Citation
  • 39.

    Outerbridge RE. The etiology of chondromalacia patellae. J Bone Joint Surg Br 1961; 43:752757.

  • 40.

    Pei M, He F, Vunjak-Novakovic G. Synovium-derived stem cell-based chondrogenesis. Differentiation 2008; 76:10441056.

  • 41.

    Farndale RW, Buttle DJ, Barrett AJ. Improved quantitation and discrimination of sulphated glycosaminoglycans by use of dimethylmethylene blue. Biochim Biophys Acta 1986; 883:173177.

    • Search Google Scholar
    • Export Citation
  • 42.

    Reddy GK, Enwemeka CS. A simplified method for the analysis of hydroxyproline in biological tissues. Clin Biochem 1996; 29:225229.

  • 43.

    Kuboki Y, Mechanic GL. The distribution of δ,δ'-dihydroxylysinonorleucine in bovine tendon and dentin. Connect Tissue Res 1974; 2:223230.

    • Search Google Scholar
    • Export Citation
  • 44.

    Reno C, Marchuk L, Sciore P, et al. Rapid isolation of total RNA from small samples of hypocellular, dense connective tissues. Biotechniques 1997; 22:10821086.

    • Search Google Scholar
    • Export Citation
  • 45.

    Pfaffl MW, Horgan GW, Dempfle L. Relative expression software tool (REST) for group-wise comparison and statistical analysis of relative expression results in real-time PCR. Nucleic Acids Res 2002; 30:e36e46.

    • Search Google Scholar
    • Export Citation
  • 46.

    Melrose J, Smith S, Cake M, et al. Comparative spatial and temporal localisation of perlecan, aggrecan and type I, II and IV collagen in the ovine meniscus: an ageing study. Histochem Cell Biol 2005; 124:225235.

    • Search Google Scholar
    • Export Citation
  • 47.

    Kambic HE, McDevitt CA. Spatial organization of types I and II collagen in the canine meniscus. J Orthop Res 2005; 23:142149.

  • 48.

    Stephan JS, McLaughlin RM Jr, Griffith G. Water content and glycosaminoglycan disaccharide concentration of the canine meniscus. Am J Vet Res 1998; 59:213216.

    • Search Google Scholar
    • Export Citation
  • 49.

    Sureshbabu A, Okajima H, Yamanaka D, et al. IGFBP-5 induces epithelial and fibroblast responses consistent with the fibrotic response. Biochem Soc Trans 2009; 37:882885.

    • Search Google Scholar
    • Export Citation
  • 50.

    Leask A, Abraham DJ. TGF-β signaling and the fibrotic response. FASEB J 2004; 18:816827.

  • 51.

    Molloy T, Wang Y, Murrell G. The roles of growth factors in tendon and ligament healing. Sports Med 2003; 33:381394.

  • 52.

    Wilkinson LS, Pitsillides AA, Worrall JG, et al. Light microscopic characterization of the fibroblast-like synovial intimal cell (synoviocyte). Arthritis Rheum 1992; 35:11791184.

    • Search Google Scholar
    • Export Citation
  • 53.

    Xu H, Edwards J, Banerji S, et al. Distribution of lymphatic vessels in normal and arthritic human synovial tissues. Ann Rheum Dis 2003; 62:12271229.

    • Search Google Scholar
    • Export Citation
  • 54.

    Pei M, Seidel J, Vunjak-Novakovic G, et al. Growth factors for sequential cellular de- and re-differentiation in tissue engineering. Biochem Biophys Res Commun 2002; 294:149154.

    • Search Google Scholar
    • Export Citation
  • 55.

    Pei M, Luo J, Chen Q. Enhancing and maintaining chondrogenesis of synovial fibroblasts by cartilage extracellular matrix protein matrilins. Osteoarthritis Cartilage 2008; 16:11101117.

    • Search Google Scholar
    • Export Citation
  • 56.

    Hoben GM, Hu JC, James RA, et al. Self-assembly of fibrochon-drocytes and chondrocytes for tissue engineering of the knee meniscus. Tissue Eng 2007; 13:939946.

    • Search Google Scholar
    • Export Citation
  • 57.

    Pazzano D, Mercier K, Moran J, et al. Comparison of chondrogenesis in static and perfused bioreactor culture. Biotechnol Prog 2000; 16:893896.

    • Search Google Scholar
    • Export Citation
  • 58.

    Smith RL, Donlon BS, Gupta MK, et al. Effects of fluid induced shear on articular chondrocyte morphology and metabolism in vitro. J Orthop Res 1995; 13:824831.

    • Search Google Scholar
    • Export Citation
  • 59.

    Davisson T, Sah RL, Ratcliffe A. Perfusion increases cell content and matrix synthesis in chondrocyte three-dimensional cultures. Tissue Eng 2002; 8:807816.

    • Search Google Scholar
    • Export Citation
  • 60.

    Fiorito S, Magrini L, Adrey J, et al. Inflammatory status and cartilage regenerative potential of synovial fibroblasts from patients with osteoarthritis and chondropathy. Rheumatology (Oxford) 2005; 44:164171.

    • Search Google Scholar
    • Export Citation
  • 61.

    Klocke NW, Snyder PW, Widmer WR, et al. Detection of synovial macrophages in the joint capsule of dogs with naturally occurring rupture of the cranial cruciate ligament. Am J Vet Res 2005; 66:493499.

    • Search Google Scholar
    • Export Citation
  • 62.

    Krey PR, Scheinberg MA, Cohen AS. Fine structural analysis of rabbit synovial cells. II. Fine structure and rosette-forming cells of explant and monolayer cultures. Arthritis Rheum 1976; 19:581592.

    • Search Google Scholar
    • Export Citation
  • 63.

    Sutton S, Clutterbuck A, Harris P, et al. The contribution of the synovium, synovial derived inflammatory cytokines and neuropeptides to the pathogenesis of osteoarthritis. Vet J 2009; 179:1024.

    • Search Google Scholar
    • Export Citation
  • 64.

    Bondeson J, Wainwright SD, Lauder S, et al. The role of synovial macrophages and macrophage-produced cytokines in driving aggrecanases, matrix metalloproteinases, and other destructive and inflammatory responses in osteoarthritis. Arthritis Res Ther 2006; 8:R187R199.

    • Search Google Scholar
    • Export Citation
  • 65.

    Lefebvre V, Huang W, Harley VR, et al. SOX9 is a potent activator of the chondrocyte-specific enhancer of the pro alpha1(II) collagen gene. Mol Cell Biol 1997; 17:23362346.

    • Search Google Scholar
    • Export Citation
  • 66.

    Bi W, Deng JM, Zhang Z, et al. Sox9 is required for cartilage formation. Nat Genet 1999; 22:8589.

  • 67.

    Appleton CT, James CG, Beier F. Regulator of G-protein signaling (RGS) proteins differentially control chondrocyte differentiation. J Cell Physiol 2006; 207:735745.

    • Search Google Scholar
    • Export Citation
  • 68.

    Enomoto-Iwamoto M, Kitagaki J, Koyama E, et al. The Wnt antagonist Frzb-1 regulates chondrocyte maturation and long bone development during limb skeletogenesis. Dev Biol 2002; 251:142156.

    • Search Google Scholar
    • Export Citation
  • 69.

    Doom M, de Bruin T, de Rooster H, et al. Immunopathological mechanisms in dogs with rupture of the cranial cruciate ligament. Vet Immunol Immunopathol 2008; 125:143161.

    • Search Google Scholar
    • Export Citation
  • 70.

    Benito MJ, Veale DJ, FitzGerald O, et al. Synovial tissue inflammation in early and late osteoarthritis. Ann Rheum Dis 2005; 64:12631267.

    • Search Google Scholar
    • Export Citation
  • 71.

    Smith MD, Triantafillou S, Parker A, et al. Synovial membrane inflammation and cytokine production in patients with early osteoarthritis. J Rheumatol 1997; 24:365371.

    • Search Google Scholar
    • Export Citation

Advertisement

Evaluation of in vitro growth factor treatments on fibrochondrogenesis by synovial membrane cells from osteoarthritic and nonosteoarthritic joints of dogs

Jennifer J. Warnock DVM, PhD1, Derek B. Fox DVM, PhD2, Aaron M. Stoker PhD3, and James L. Cook DVM, PhD4
View More View Less
  • 1 Comparative Orthopaedic Laboratory, College of Veterinary Medicine, University of Missouri, Columbia, MO 65211.
  • | 2 Comparative Orthopaedic Laboratory, College of Veterinary Medicine, University of Missouri, Columbia, MO 65211.
  • | 3 Comparative Orthopaedic Laboratory, College of Veterinary Medicine, University of Missouri, Columbia, MO 65211.
  • | 4 Comparative Orthopaedic Laboratory, College of Veterinary Medicine, University of Missouri, Columbia, MO 65211.

Abstract

Objective—To determine the in vitro effects of selected growth factors on fibrochondrogenesis by synovial membrane cells from nonosteoarthritic (normal) and osteoarthritic joints of dogs.

Animals—5 dogs with secondary osteoarthritis of shoulder or stifle joints and 6 dogs with normal joints.

Procedures—Synovial membrane cells were harvested from normal and osteoarthritic joints and cultured in monolayer with or without (control) basic fibroblast growth factor, transforming growth factor-β1, and insulin-like growth factor-1. In the cultured cells, fibrochondrogenesis was measured by use of a real-time reverse transcriptase PCR assay to determine relative expressions of collagen I, collagen II, and aggrecan genes and of 3 genes involved in embryonic chondrogenesis: Sry-type homeobox protein-9 (SOX-9), frizzled-motif associated with bone development (Frzb), and regulator of G-protein signaling-10 (RGS-10). Tissue collagen content was measured via a hydroxyproline assay, and sulfated glycosaminoglycan content was measured via a 1,9-dimethylmethylene blue assay. Cellularity was determined via a double-stranded DNA assay. Immunohistochemical analysis for collagens I and II was also performed.

Results—In vitro collagen synthesis was enhanced by growth factor stimulation. Although osteoarthritic-joint synoviocytes could undergo a fibrocartilage-like phenotypic shift, their production of collagenous extracellular matrix was less than that of normal-joint synoviocytes. Gene expressions of SOX-9 and RGS-10 were highest in the osteoarthritic-joint cells; Frzb expression was highest in growth factor treated cells.

Conclusions and Clinical Relevance—Autogenous synovium may be a viable cell source for meniscal tissue engineering. Gene expressions of SOX-9 and RGS-10 may be potential future targets for in vitro enhancement of chondrogenesis.

Abstract

Objective—To determine the in vitro effects of selected growth factors on fibrochondrogenesis by synovial membrane cells from nonosteoarthritic (normal) and osteoarthritic joints of dogs.

Animals—5 dogs with secondary osteoarthritis of shoulder or stifle joints and 6 dogs with normal joints.

Procedures—Synovial membrane cells were harvested from normal and osteoarthritic joints and cultured in monolayer with or without (control) basic fibroblast growth factor, transforming growth factor-β1, and insulin-like growth factor-1. In the cultured cells, fibrochondrogenesis was measured by use of a real-time reverse transcriptase PCR assay to determine relative expressions of collagen I, collagen II, and aggrecan genes and of 3 genes involved in embryonic chondrogenesis: Sry-type homeobox protein-9 (SOX-9), frizzled-motif associated with bone development (Frzb), and regulator of G-protein signaling-10 (RGS-10). Tissue collagen content was measured via a hydroxyproline assay, and sulfated glycosaminoglycan content was measured via a 1,9-dimethylmethylene blue assay. Cellularity was determined via a double-stranded DNA assay. Immunohistochemical analysis for collagens I and II was also performed.

Results—In vitro collagen synthesis was enhanced by growth factor stimulation. Although osteoarthritic-joint synoviocytes could undergo a fibrocartilage-like phenotypic shift, their production of collagenous extracellular matrix was less than that of normal-joint synoviocytes. Gene expressions of SOX-9 and RGS-10 were highest in the osteoarthritic-joint cells; Frzb expression was highest in growth factor treated cells.

Conclusions and Clinical Relevance—Autogenous synovium may be a viable cell source for meniscal tissue engineering. Gene expressions of SOX-9 and RGS-10 may be potential future targets for in vitro enhancement of chondrogenesis.

Meniscal injuries are a common cause of stifle joint arthritis and pain in dogs and people and have been identified as a cause of lameness and decreased performance in horses.1–6 Tears within the axial avascular portion of the meniscus possess a limited ability to heal spontaneously,7,8 and attempts to successfully encourage avascular meniscal healing9–14 or regeneration12–16 have had variable outcomes. Complete restoration of the avascular damaged meniscus, both in form and function, has not been accomplished in any species,11 to our knowledge. To ameliorate painful joint locking and popping, most avascular meniscal injuries in dogs and humans are treated with partial meniscectomy. However, partial meniscectomy itself results in articular cartilage damage, and progression of osteoarthritis usually ensues.7,17

Currently, tissue engineering is being investigated for the treatment of avascular meniscal injury or total meniscal loss. A major procedural challenge in tissue engineering is determining the ideal source of cells for in vitro formation of replacement tissue. Synovium has been investigated as a cell source for articular cartilage tissue engineering; synoviocytes readily undergo in vitro hyaline chondrogenesis.18–21 Many of these investigations have used bovine, lapine, or murine synovium from clinically normal joints of young animals.18,20,22 In contrast, meniscus-deficient joints have clinical signs of osteoarthritis with pathologically affected synovium.17,23–27 Use of normal-joint synoviocytes as a cell source for meniscal tissue engineering would require surgical violation of another unaltered joint within the patient's body or allogenic donation. Primary joint exploration for surgical treatment for ligamentous instability or a torn meniscus would provide an excellent opportunity for harvest of autologous synovium. Should a second surgery be required for progressive meniscal disease or postliminary meniscal tears, the patient's own cells would be available to create replacement meniscal tissue. Advantages of the use of autogenous cells for tissue engineering include lack of immunoreactivity28,29 and decreased risk of infectious disease transmission.30,31 Thus, use of autologous synovium from affected joints may be a rational cell source choice for meniscal fibrocartilage engineering.

Because of the synovium's ability to undergo chondrogenesis in secondary osteochondromatosis32 and its involvement in periarticular osteophyte formation,33,34 osteoarthritic-joint synovium could be primed for chondrogenic differentiation and may have advantages over normal-joint synovial membrane cells for fibrocartilage engineering. Compared with findings in healthy organ donors, twice as many potentially chondrogenic mesenchymal precursor cells have been observed on the tips of synovial villi in patients with osteoarthritis.35 Although both normal- and osteoarthritic-joint type B synoviocytes produce TGF-β136(a chondrogenic growth factor), only osteoarthritic-joint type B synoviocytes express TGF-β1 receptors.36 Conversely, the proliferation of inflammatory mediators and destructive enzymes from the synovial membrane in an osteoarthritic joint could negatively impact the in vitro chondrogenic potential of synoviocytes. Other cell types in degenerative joints have decreased ECM synthesis capabilities; for example, osteoarthriticjoint chondrocytes from sheep have reduced proliferation and poor ECM formation in response to chondrogenic growth factor treatment, compared with normal-joint chondrocytes in monolayer culture.37 Mesenchymal stem cells aspirated from bone marrow adjacent to severely arthritic joints in humans have decreased in vitro chondrogenic activity, compared with similar cells from bone marrow of the iliac crest in individuals without arthritis.38 The true in vitro chondrogenic potential of canine osteoarthritic-joint synovium is unknown; this crucial variable must be delineated if autologous synovium is to be used for meniscal fibrocartilage tissue engineering in that species.

The primary objective of the study reported here was to compare the fibrochondrogenic potential of synoviocytes from osteoarthritic and nonosteoarthritic (normal) joints in dogs by examining fibrocartilage-like ECM synthesis in the cells in monolayer culture with and without exposure to chondrogenic growth factors. A second objective was to determine whether 3 genes involved in embryonic chondrogenesis were induced and expressed in adult tissue in response to growth factors. We hypothesized that the fibrocartilage ECM production and gene expression in canine osteoarthriticjoint synoviocytes cultured in monolayer and treated with chondrogenic growth factors would not differ from findings in similarly treated normal-joint synoviocytes, and that expressions of the SOX-9, Frzb-1, and RGS-10 genes (known to be involved in fetal chondrogenesis) in synoviocytes from normal and osteoarthritic joints would not differ.

Materials and Methods

Synoviocyte culture–-Dogs were included in the study if they underwent an antemortem orthopedic examination and were not receiving any medications. Dogs were euthanatized for reasons unrelated to the study via an IV overdose injection of pentobarbital sodium, according to guidelines of the University of Missouri Animal Care and Use Committee. Within 1 to 2 hours of euthanasia, joints of interest were surgically approached via a lateral arthrotomy. The intra-articular structures and joint surfaces were observed and palpated with a hook-tipped probe. Synovium (including the intima and subintima) from the entire joint was aseptically harvested. Synovium was collected from both stifle joints of 6 large-breed dogs that were subjectively determined to be free of orthopedic disease on the basis of results of antemortem orthopedic examination and postmortem visual and tactile examination of the joint interior. Synovium was also harvested from 5 large-breed dogs that had signs of pain, decreased range of motion, crepitus, and joint effusion localized to the stifle joint (naturally occurring cranial cruciate ligament degeneration and medial meniscal tear in 2 dogs [3 joints], cranial cruciate ligament transection [Pond-Nuki method; 3 months' duration] in 1 dog [1 joint]) and the shoulder joint (humeral head osteochondrosis dissecans in 1 dog [2 joints] and chronic shoulder joint instability in 1 dog [2 joints]). On visual and tactile examination of the joints of the 5 dogs with osteoarthritic joints, moderate to severe periarticular osteophytosis, grossly visible synovial villous tufts, and grade 3 or 4 Outerbridge cartilage lesions39 were evident. With the exception of the dog that was treated via the Pond-Nuki method, the duration of osteoarthritis in all other joints was estimated to be very chronic and longer than 3 months. The synovial tissue from each joint was placed in DMEM supplemented with 10% fetal bovine serum, 0.008% HEPES buffer, 0.008% nonessential amino acids, 0.002% penicillin (100 U/mL) and streptomycin (100 μg/mL), amphotericin B (25 μg/mL), 0.015% L-ascorbate (0.15 mg/mL), pyruvate (0.22 mg/mL), and 0.01% L-glutamine in preparation for monolayer culture.

The synovium was sectioned into 2.0 × 2.0-mm pieces with a Bard-Parker scalpel and No. 10 bladea under sterile conditions. The tissue fragments were digested with sterile type 1A clostridial collagenase solutionb(7.5 mg/mL) in RPMI 1640 solution. The mixture was agitated and maintained at 37°C with 5% CO2and 95% humidity for 6 hours until the tissue was completely digested. Cells were recovered through centrifugation; the supernatant was decanted and the cellular pellet was resuspended in 5 mL of supplemented DMEM. The cell solution was transferred to a 75-cm2tissue culture flask containing 10 mL of supplemented DMEM. The flasks were incubated at 37°C with 5% CO2and 95% humidity; a sterile medium change was performed every 3 days. Synoviocytes were monitored for growth by use of an inverted microscope until observance of 95% cellular confluence in the tissue culture flask. At 95% confluence, twenty 75-cm2flasks/dog were seeded with normal- or osteoarthritic-joint synoviocytes (second passage cells) at a concentration of 350,000 cells/flask. Cells were grown to 99% confluence and then randomly allocated into 4 groups: a control group of normal-joint synoviocytes treated with serum-free medium (5 flasks/dog), a control group of osteoarthritic-joint synoviocytes treated with serum-free medium (5 flasks/dog), a treatment group of normal-joint synoviocytes treated with serum-free medium and growth factors (5 flasks/dog), and a treatment group of osteoarthritic-joint synoviocytes treated with serum-free medium and growth factors (5 flasks/dog). The serum-free medium contained 0.008% HEPES buffer, 0.008% nonessential amino acids, 0.002% penicillin (100 U/mL) and streptomycin (100 μg/mL), amphotericin B (25 μg/mL), 0.015% L-ascorbate (0.15 mg/mL), pyruvate (0.22 mg/mL), and 0.01% L-glutamine. Growth factor treatment consisted of human recombinant (bFGFc[50 ng/mL] for 3 days, followed by TGF-β1d[10 ng/mL] with IGF-1e[500 ng/mL] as described by Pei et al19,40) for 13 days. All flasks contained 14 mL of medium. Total duration of culture for all groups was 16 days; a sterile medium change was performed every 3 days. On the 16th day, the flasks were washed with serum-free medium, and the cells were enzymatically released from the culture flask by use of a cell dissociation reagent.f Cell viability was assessed by use of a trypan blue exclusion assay.

Histologic analysis–-Tissue-like constructs harvested from each flask were cut in 3 portions, and 1 portion was fixed in neutral-buffered 10% formalin and sectioned. For histologic analysis, sections were stained with H&E, Masson trichrome, toluidine blue, or immunohistochemical stains for collagen I and II. Samples were processed within 24 hours of tissue collection.

For immunohistochemical analysis, tissues were cut in 4-μm-thick sections, which were placed on positively charged slides, microwaved, and kept on a slide warmer at 43°C overnight (approx 12 hours). The sections on the slides were then hydrated, placed in 0.4% pepsin, and heated in an incubator at 37°C for 20 minutes. The slides were rinsed in tap water and placed in Tris buffer for at least 5 minutes. Subsequent staining was done on an automated slide staining system.g Slides were treated with 3% H2O2for 15 minutes and with a protein block for 5 minutes. For detection of collagen I, slides were incubated in goat anti-collagen type I antibodiesh(1:100 concentration) for 30 minutes; negative control slides were treated with goat IgG (1:1,000 concentration) instead of the primary antibody. Secondary and tertiary reagents in a labeled streptavidin biotin reagent systemi were applied for 20 minutes each. For detection of collagen II, slides were incubated with rabbit anti-bovine collagen type II AB746P antibodyi(1:100 concentration) for 30 minutes. Negative control slides were treated with rabbit IgG (1:1,000 concentration) instead of the primary antibody. Secondary reagent in a labeled polymer immunohistochemical staining systemi was applied for 30 minutes. Following antibody treatment for collagen I or collagen II, slides were incubated with a red chromagenj for 10 minutes. Slides were counterstained in Mayer hematoxylin staink for 1 minute and then dehydrated and coverslipped.

Histologic specimens were examined by use of a standard light microscopel at ×10 magnification. Images of each section (3 from the scaffold periphery and 3 from the scaffold center) at 2, 6, and 10 o'clock positions were digitally captured by use of a digital cameram and saved electronically.

DNA quantification–-A third of the harvested tissue was lyophilized, and its dry weight was measured. Samples were incubated in 1.0 mL of papain solution (2mM dithiothreitol and 300 μg of papain/mL) at 60°C in a water bath for 12 hours.

A double-stranded DNA quantification assayn was used for double-stranded DNA quantification. Double-stranded DNA extracted from bovine thymus was mixed with Tris-EDTA buffero to create standard DNA concentrations of 1,000, 100, 10, and 1 ng/mL The standards and 100 μL of each papain-digested sample (used in the aforementioned assays) were added to a black 96-well plate; 100 μL of a fluorescent nucleic acid stain for double-stranded DNAn(2 μg/mL) was added to each well, and the plate was incubated for 5 minutes. Sample fluorescence was read by use of a spectrophotometric plate readerp(excitation, 485 nm; emission, 528 nm). Absorbances were converted to concentrations (in ng/mL) and total double-stranded DNA yield (in ng) with computer software.q

Biochemical ECM analysis–-By use of the aforementioned papain digest, each group was analyzed for GAG and collagen contents. The GAG content of each scaffold was assessed by use of a dimethylmethylene blue sulfated GAG assay.41 The collagen content of each scaffold was assessed by use of the Erlich hydroxyproline assay, as described by Reddy et al.42 Median percentage GAG or hydroxyproline content was standardized to tissue cellularity ([μg of GAG or hydroxyproline/μg of double-stranded DNA] × 100%) as a chondrogenic index19 to identify fibrochondrogenic activity of each tested cell type; the median percentage GAG or hydroxyproline content was also standardized to tissue dry weight to allow comparison of the experimental tissue data with normal meniscal ECM content. Hydroxyproline is approximately 13% of collagen content, and hydroxyproline was converted to collagen content by use of an equation16,43 as follows:

article image

Relative gene expression–-A third of the harvested tissue was analyzed via RT-PCR assay to determine the relative expressions of collagen I, collagen II, aggrecan, SOX-9, RGS-10, and Frzb genes (expressed relative to GAPDH gene expression). Primer sequences were designed by use of primer development softwarer(Appendix). Samples were stored in an RNA stabilization reagents at −20°C.

Samples were transferred to 0.5-mL screw-cap tubes filled with 1.0-mm-diameter zirconia beads and a total RNA isolation reagentt and homogenized by use of a minibead beater for 2 minutes. The RNA was extracted from each homogenate by use of the TRIspin method.44 Five microliters of each sample's RNA was diluted with 95 μL of RNase-free water, and the absorbances were read at 260 and 280 nm with a spectrophotometric plate reader.p Absorbances were converted to concentrations (in μg/μL) by data reduction software.q Reverse transcription was performed with equal amounts of sample RNA, which was converted to cDNA by use of random hexamer primers and RT enzymeu in an RT system.v The PCR assay was then performed with primers that corresponded to cDNA sequences for collagen I, collagen II, aggrecan, SOX-9, RGS-10, Frzb-1, and the housekeeping gene GAPDH in a thermal cyclerw and with a SYBR green PCR kitx following the manufacturer's guidelines. The PCR profile for all tests consisted of an initial incubation at 94°C for 15 minutes, followed by 55 cycles of 5 seconds at 94°C (melting), 10 seconds at 57°C (annealing), and 20 seconds at 72°C (extension). After the PCR profile was completed, a melt curve analysis was done to ensure specific amplification for each sample. The SYBR green fluorescence was monitored during the extension step of the PCR profile, and takeoff values and amplification efficiencies were determined with the thermal cycler software. Relative expressions of the target genes evaluated were expressed as a ratio to the expression of the GAPDH gene. Differences in gene expression among groups were determined by use of a relative expression statistical tool.45

Statistical analysis–-Nonparametric data including relative gene expressions and DNA, collagen, and GAG contents were compared between control and growthfactor–-treated synoviocytes and between normal-joint synoviocytes and osteoarthritic-joint synoviocytes by use of a Kruskal-Wallis 1-way ANOVA on ranks followed by Mann-Whitney rank sum tests. Significance was set at a value of P>0.05. Calculations were completed by use of a statistical software program.y

Results

Gross observations–-Normal- and osteoarthritic-joint cells in the growth-factor–treated groups formed fibrous tissue-like constructs, which remained in a cohesive sheet after enzymatic release (Figure 1). The fibrous appearance was apparent in growth-factor–treated normal-joint synoviocytes 3 or 4 days earlier than it was in similarly treated osteoarthritic-joint synoviocytes. The corners of all growth-factor–treated samples began to contract slightly during the last 2 or 3 days of culture. Growth-factor–treated osteoarthritic-joint synoviocytes from a dog that had undergone cranial cruciate ligament transection 3 months prior to the study spontaneously formed micromasses 6 days after addition of growth factors (Figure 2). The control normal- and osteoarthritic-joint synoviocytes were noncohesive cells and did not produce visible ECM, thereby precluding histologic analysis.

Figure 1—
Figure 1—

Photographs of the typical appearance of monolayer tissue-like constructs formed in 75-cm2culture flasks by canine synoviocytes after 16 days of growth factor exposure (A) and after enzymatic release (B). The tissue was formed by synoviocytes harvested from a canine stifle joint with naturally occurring cranial cruciate ligament degeneration; the dog had been euthanatized for reasons unrelated to the study.

Citation: American Journal of Veterinary Research 72, 4; 10.2460/ajvr.72.4.500

Figure 2—
Figure 2—

Photographs illustrating the spontaneous formation of micromasses in a monolayer culture of osteoarthritic-joint synoviocytes harvested from a canine stifle joint that had undergone cranial cruciate ligament transection (Pond-Nuki method) 3 months earlier; the dog had been euthanatized for reasons unrelated to the study. A–-Image of a micromass in a 75-cm2 culture flask. B–-Image of 3 micromasses collected from culture medium after 16 days of sequential growth factor exposure.

Citation: American Journal of Veterinary Research 72, 4; 10.2460/ajvr.72.4.500

Histologic analysis–-Histologic analysis of H&E-stained sections of tissue-like constructs obtained following growth factor treatment revealed regular layers of eosinophilic fibrous tissue, the cells of which had a rounded phenotype and were present in clusters or singly (Figure 3). Some single cells appeared in lacunae-like spaces. In the center of the micromasses, cellular disintegration, karyolysis, and amorphous debris consistent with cell death were present. In both the growth-factor–treated normal- and osteoarthritic-joint constructs, staining for collagen with Masson trichrome stain yielded positive results, but staining for GAGs with toluidine blue yielded negative results.

Figure 3—
Figure 3—

Photomicrographs of monolayer tissue constructs generated by non-osteoarthritic (normal)-joint synoviocytes (column A) from one dog (2 joints) and osteoarthritic-joint synoviocytes (column B) obtained from another dog (2 joints) that had undergone culture with sequential exposure to growth factors for 16 days and were stained with H&E stain (row 1), Masson trichrome stain (row 2), or toluidine blue (row 3). In row 2, blue fibrillar staining indicates the presence of collagen (thin arrows). In row 3, notice the lack of deep purple or indigo staining that indicates the presence of GAGs in the ECM. In all panels, bar = 50 μm.

Citation: American Journal of Veterinary Research 72, 4; 10.2460/ajvr.72.4.500

Cellularity and cell viability–-Tissue cellularity, as measured by DNA content standardized to dry weight, did not differ significantly (P= 0.19) among groups. Cellular viability was estimated at 95% to 99% in all groups except specimens that spontaneously formed micromasses. An exact measure of cell viability could not be obtained because of cell clumping and cell retention in the tissue-like constructs of the growth-factor–treated normal- and osteoarthritic-joint synoviocyte groups.

Biochemical ECM analysis–-Among synoviocyte groups, there was no significant (P= 0.873) difference in the relative expression of the collagen I gene (Table 1). Collagen II expression was evaluated only in the control osteoarthritic-joint synoviocytes and the growth-factor–treated normal- and osteoarthritic-joint synoviocytes because a pellet of nucleic acids could not be isolated from the control normal-joint synoviocytes. No significant (P= 0.239) difference in median relative expression of the collagen II gene was detected among groups. Median values of percentage hydroxyproline produced (standardized to double-stranded DNA content) by the growth-factor–treated normal-joint synoviocytes and growth-factor–treated osteoarthritic-joint synoviocytes were each significantly (P>0.001) higher than the value in each of the control groups. The median percentage hydroxyproline produced by the treated normal-joint synoviocytes was significantly (P= 0.03) higher, compared with the value for the treated osteoarthritic-joint synoviocytes.

Table 1—

Median (range) relative gene expressions (No. of target gene transcript copies/No. of GAPDH copies) of collagen I, collagen II, aggrecan, SOX-9, RGS-10, and Frzb and percentage tissue content of hydroxyproline, collagen, and GAGs in tissue constructs formed by canine synoviocytes from nonosteoarthritic (normal) and osteoarthritic joints* that were or were not (control cells) sequentially exposed to growth factors for 16 days.

 Control groupGrowth-factor–treated group
VariableNormal-joint synoviocytesOsteoarthritic-joint synoviocytesNormal-joint synoviocytesOsteoarthritic-joint synoviocytes
Relative collagen I gene expression65,100.019,100.021,600.045,700.0
 (8,539.800–121,767.000)(4,405.000–326,926.000)(10,120.549–103,911.802)(1,195.361–174,153.507)
Relative collagen II gene expressionNA0.1450.2100.540
  (0.009–0.220)(0.009–0.413)(0.088–1.057)
Relative aggrecan gene expression0.0120.014a0.290a0.570
 (0.020–0.023)(>0.001–0.060)(0.090–1.080)(0.009–5.740)
Relative SOX-9 gene expression0.0960.0720.1590.540
 (0.000–0.193)l(>0.001–0.190)m(0.137–0.468)n(0.396–1.939)l,m,n
Relative RGS-10 gene expression000.003o0.007o
   (>0.001–0.025)(0.005–0.012)
Relative Frzb-1 gene expression0.207p,q0.247p,q0.725p0.705q
 (0.097–0.283)(0.176–0.288)(0.681–0.769)(0.594–0.797)
Percentage tissue hydroxyproline content standardized to tissue DNA content§0.001d,e0.001d,e2,791.000d,f801.000e,f
 (0.001–08.040)(1,006.697–5,943.502)(272.656–2,431.620) 
Percentage tissue collagen content standardized to dry tissue weight0.060i,j0.080h,k19.500g,i,k9.000g,h,j
 (0.013–5.070)(0.013–5.551)(4.771–32.400)(3.520–14.664)
Percentage GAG content standardized to tissue DNA content882.300b,c753.2001,778.000b1,480.600c
 (672.600–952.300)(328.300–2,102.000)(1,147.400–2,283.000)(915.000–2,283.000)
Percentage GAG content standardized to dry tissue weight1.0000.7000.8000.900
 (0.685–1.771)(0.251–1.780)(0.702–1.204)(0.834–1.238)

Synoviocytes were harvested from normal stifle joints of 6 dogs (12 joints) and osteoarthritic stifle or shoulder joints of 5 dogs (8 joints) after euthanasia (performed for reasons unrelated to the study).

Growth factor exposure involved treatment with bFGF (50 ng/mL) for 3 days, followed by TGF-β1 (10 ng/mL), with IGF-1 (500 ng/mL) for 13 days; control cells were cultured under similar conditions without treatment with growth factors.

Gene expressions of collagen I, collagen II, aggrecan, SOX-9, RGS-10, and Frzb are expressed as the number of gene copies relative to the number of gene copies of GAPDH.

Percentage hydroxyproline content was standardized to tissue cellularity ([μg of hydroxyproline/μg of double-stranded DNA] × 100%).

Percentage collagen content was standardized to tissue dry weight; hydroxyproline is approximately 13% of collagen content, and hydroxyproline content was converted to collagen content by use of an equation16,42 as follows: Collagen content (μg) = Hydroxyproline content (μg) × dilution factor × 13.

Percentage GAG content was standardized to tissue cellularity ([μg of hydroxyproline/μg of double-stranded DNA] × 100%) and also standardized to tissue dry weight.

NA = Not applicable.

For a given variable, group values with the same superscript letter differ significantly as follows:

P= 0.004

P= 0.016

P= 0.05

P= 0.001

P= 0.001

P= 0.03)

P= 0.03

P= 0.006

P= 0.009

P= 0.014

P= 0.003

P= 0.05

P= 0.001

P= 0.005

P= 0.01

P= 0.01

P= 0.05.

Percentage collagen content, as measured by the hydroxyproline assay (values standardized to tissue dry weight), was highest in the growth-factor–treated normal-joint synoviocytes (Table 1). Immunohistochemical staining for collagen I was strong in all growth-factor–treated normal- and osteoarthritic-joint synoviocytes (Figure 4); mild immunoreactivity for collagen II was seen intracellularly and pericellularly in all normal- and osteoarthritic-joint synoviocyte samples (Figure 5).

Figure 4—
Figure 4—

Photomicrographs of monolayer tissue constructs generated by normal-joint synoviocytes from one dog (2 joints [A and B]) and osteoarthritic-joint synoviocytes (C and D) obtained from another dog (2 joints) that had undergone culture with sequential exposure to growth factors for 16 days and were stained immunohistochemically for collagen I (A and C) or without the primary antibody (negative control samples; B and D). Notice the strong immunoreactivity to collagen I in the normal- and osteoarthritic-joint synoviocytes; the cells have a rounded phenotype (thick arrows) and fibrillar ECM (thin arrows). In panels A and C, immunohistochemical stain for collagen I, red chromagen, and Mayer hematoxylin stain; bar = 50 μm. In panels B and D, negative control immunohistochemical stain, red chromagen, and Mayer hematoxylin stain; bar = 50 μm.

Citation: American Journal of Veterinary Research 72, 4; 10.2460/ajvr.72.4.500

Figure 5—
Figure 5—

Photomicrographs of monolayer tissue constructs generated by normal-joint synoviocytes (A and B) from one dog (2 joints) and osteoarthritic-joint synoviocytes (C and D) obtained from another dog (2 joints) that had undergone culture with sequential exposure to growth factors for 16 days and were stained immunohistochemically for collagen II (A and C) or without the primary antibody (negative control samples; B and D). Notice the mild intracellular and pericellular immunoreactivity to collagen II in the normal- and osteoarthritic-joint synoviocytes; the cells have a rounded phenotype. In panels A and C, immunohistochemical stain for collagen II, red chromagen, and Mayer hematoxylin stain; bar = 50 μm. In panels B and D, negative control immunohistochemical stain, red chromagen, and Mayer hematoxylin stain; bar = 50 μm.

Citation: American Journal of Veterinary Research 72, 4; 10.2460/ajvr.72.4.500

Relative aggrecan gene expression was significantly higher (P= 0.004) in growth-factor–treated normal-joint synoviocytes, compared with findings in control osteoarthritic-joint synoviocytes (Table 1). The median relative aggrecan expressions in normal- and osteoarthritic-joint synoviocytes that were growth factor treated were not significantly (P= 1.0) different. Relative gene expression was not significantly different between control osteoarthritic-joint synoviocytes and growth-factor–treated osteoarthritic-joint synoviocytes (P= 0.073), between control normal-joint synoviocytes and growth-factor–treated normal-joint synoviocytes (P= 0.223), or between control normal-joint synoviocytes and growth-factor–treated osteoarthritic-joint synoviocytes (P= 0.22). Median percentage GAG content (standardized to tissue cellularity) was greatest for growth-factor–treated normal-joint synoviocytes; values for the growth-factor–treated normal- and osteoarthritic-joint synoviocytes were each greater than that for the control normal-joint synoviocytes (P= 0.016 and P= 0.05, respectively). The GAG content did not differ between the growth-factor–treated normal-joint synoviocytes and growth-factor–treated osteoarthritic-joint synoviocytes (P= 0.339), between growth-factor–treated osteoarthritic-joint synoviocytes and control osteoarthritic-joint synoviocytes (P= 0.196), or between growth-factor–treated normal-joint synoviocytes and control osteoarthritic-joint synoviocytes (P= 0.064); however, the post hoc power of these tests was low (0.05, 0.04, and 0.48, respectively). On a dry-weight basis, median percentage GAG content in the 4 groups did not differ significantly from each other (for the 6 comparisons, P= 0.30 to 1.0).

Genes of embryonic chondrogenesis–-The growth-factor–treated osteoarthritic-joint synoviocytes had the greatest expression of the SOX-9 gene, which differed significantly from the expression in control normal-joint synoviocytes (P= 0.05), control osteoarthritic-joint synoviocytes (P= 0.001), or growth-factor–treated normal-joint synoviocytes (P= 0.005; Table 1). No difference in SOX-9 gene expression was detected between the 2 control groups (P= 0.864) or between control normal-joint synoviocytes and growth-factor–treated normal-joint synoviocytes (P= 0.332). The relative gene expression of RGS-10 in the growth-factor–treated osteoarthritic-joint synoviocytes was significantly (P= 0.01) greater than the expression in the growth-factor–treated normal-joint synoviocytes. The RGS-10 gene was not expressed by either control group. Relative expression of Frzb-1 in the growth-factor–treated normal joint synoviocytes was significantly greater than both controls (P= 0.01). Relative expression of Frzb-1 in the growth-factor–treated osteoarthritic joint synoviocytes was also significantly greater than both controls (P= 0.05). No significant differences were found between growth-factor–treated synoviocytes from normal and osteoarthritic joints (P= 0.226) nor between controls from normal and osteoarthritic joints (P= 0.20).

Discussion

In the present in vitro study, growth factors enhanced cellular collagen and GAG synthesis in normal-joint synoviocytes and enhanced cellular collagen synthesis in osteoarthritic-joint synoviocytes obtained from dogs. Canine synoviocytes from normal and osteoarthritic joints can express collagen I, collagen II, and the aggrecan genes in monolayer culture, which may be useful for meniscal tissue engineering. Collagen II and aggrecan are the principal ECM constituents of the axial, avascular portion of the stifle joint meniscus, whereas collagen type I is the major structural protein of the abaxial vascular meniscus.46–48 In osteoarthritic-joint synoviocytes, spontaneous signaling for cartilage formation was evident in the present study; osteoarthritic-joint synoviocytes that were not treated with growth factors expressed aggrecan and collagen II genes. This may be attributable to self-expression of TGF-β1 and its receptor35 in those cells. Expression of the collagen I gene seemed to be independent of the cells' pathological state and growth factor administration in our study. However, actual transcribed collagen I was strongly influenced by growth factor administration in both normal- and osteoarthritic-joint synoviocytes, and only the growth-factor–treated cell groups produced sufficient collagen I to form a tissue-like construct for histologic analysis. This finding is consistent with the fact that bFGF, TGF-β1, and IGF-1 are potent inducers of type I collagen formation.49–51 In addition, the synoviocyte cultures in our study likely contained a mixture of intimal mesenchymal progenitor cells, type B synoviocytes, subintimal fibroblasts, vascular fibroblasts, and lymphatic fibroblasts, all of which constitutively produce type I collagen.52,53

It is interesting to note that, regardless of the serum-free environment, no differences in cell numbers were found between the control and growth factor treatment groups in the present study. Standardization of the total number of cells seeded into each culture flask may have accounted for this finding. Growth factor treatment was commenced when 99% cellular confluence was achieved, and cell contact inhibition may have prevented further cellular proliferation in these nonneoplastic synoviocytes. Use of human recombinant growth factors on canine cells could have also contributed to decreased bioactive efficacy. A growth factor protocol pioneered by Pei et al19,54 was originally chosen for our study because it resulted in superior hyaline synovial chondrogenesis, compared with results of treatments with other growth factor combinations and dosages. Recent research55 has revealed that treatment with a combination of TGF-β1, IGF-1, and bFGF followed by exposure to TGF-β1 and IGF-1 in serum-free medium is a better means of increasing growth of synoviocytes, compared with the protocol used in the present study.

It is known that human chondrocytes and meniscal fibrochondrocytes also form self-assembled constructs in plastic vessels,56,57 similar to the synovial tissue-like constructs generated in monolayer culture in the present study. The human chondrocyte constructs contained 4.1% collagen and 23% GAGs on a dry weight basis57; fibrochondrocyte constructs contained 1% collagen and 2% GAGs on a dry weight basis.56 In our study, the growth-factor–treated synovial tissue constructs contained 9.0% to 19.5% collagen and 0.8% to 0.9% GAGs. The ECM content of synoviocyte-derived tissue is lower than the GAG and collagen contents of a normal meniscus (2% to 3% and 60% to 75%, respectively47,48). There are numerous possible reasons why the synoviocytes in the present study did not achieve matrix production equivalent to that of a normal meniscus. The synoviocytes were grown for a short period without scaffolding or biomechanical stimuli to guide ECM formation. Reported duration of cell culture that results in gross hyaline cartilaginous tissue formation ranges from 28 to 32 days,57–59 whereas the duration of cell culture in the present study was 16 days. A shorter duration of culture is financially desirable, decreases the requirement for time and personnel, and decreases the risks for contamination and culture complications. Thus, a culture period of 16 days was investigated in our study to determine whether viable tissue constructs with measureable ECM could be generated in a comparatively shorter period. Biomechanical stimulation, in the form of mechanical stimulation of cultured cells in bioreactors, has positive mechanotransductive effects on cell differentiation, cell viability, and ECM production.57–59 The ideal sequential growth factor–biomechanical protocol has not yet been determined for in vitro meniscogenesis, to our knowledge.

In our study, growth-factor–treated canine osteoarthritic-joint synoviocytes produced less collagen than canine normal-joint synoviocytes that were treated with growth factors. A similar finding was noted in a study by Fiorito et al60 in which in vitro hyaline chondrogenesis of pelleted human synoviocytes was investigated; human osteoarthritic-joint synoviocytes also have markedly reduced hyaline chondrogenic activity, compared with normal-joint synoviocytes. Although a mechanism for this decreased collagen formation was not determined in the present study, possible causes may be related to synovial inflammatory cell infiltrates. Synovial macrophages are present in increased numbers in the synovium of canine osteoarthritic joints and contribute to the inflammatory environment61; those cells may impact in vitro ECM formation. It is commonly assumed that synovial macrophages in first-pass monolayer culture are eliminated during the first several medium changes, as reported by Krey et al.62 However, recent work by Pei et al19 revealed that synoviocytes derived from conventional monolayer passage techniques, as used in our study, are contaminated with macrophages. Macrophage-contaminated synovial cell cultures have decreased in vitro hyaline chondrogenesis with sparse production of collagen II and 50% less GAGs, compared with findings in uncontaminated cultures.19 A plethora of inflammatory mediators (including tumor necrosis factor α, interleukins, prostaglandins, and matrix metalloproteinases60,63,64) and their receptors are secreted by osteoarthritic-joint (type B) synoviocytes and synovial macrophages, which could contribute to the reduced collagen formation by osteoarthritic-joint synoviocytes in the present study. Further investigation will be required to reveal the exact mechanism by which growth-factor–treated osteoarthritic-joint synoviocytes produce comparatively less collagen than growth-factor–treated normal-joint synoviocytes.

The mechanism for in vitro fibrochondrogenesis has not been fully elucidated, and identification of the molecular signals in this process may provide targets for optimization of in vitro fibrochondrogenesis. Srytype homeobox protein-9 is a high-mobility-group domain transcription factor that is expressed in all primordial cartilages and induces collagen II and aggrecan synthesis.65,66 Targeting SOX-9 expression could enhance axial meniscal ECM production.47,48 In the present study, SOX-9 gene expression was highest in osteoarthritic-joint synoviocytes that were stimulated with growth factors. This may have been mediated by the TGF-β1 in our growth factor mixture as well as endogenous receptor priming of osteoarthritic-joint synoviocytes. Regulator of G-protein signaling-10 increases the rate and number of GAG and collagen IIa transcripts produced,67 through induction of SOX-9.67 In the present study, RGS-10 gene expression mirrored SOX-9 gene expression. Despite increased expressions of both RGS-10 and SOX-9 in growth-factor–treated osteoarthritic-joint synoviocytes, collagen II synthesis was not subjectively greater in those cells than it was in growth-factor–treated normal-joint synoviocytes. This finding suggested that the RGS-10 and SOX-9 genes may have different roles in this particular in vitro environment or that posttranscriptional and posttranslational confounding signals were present.

In the fetus, Frzb causes epiphyseal chondrocytes to maintain a stable immature phenotype.68 In the present study, gene expression of Frzb-1 was of particular interest because it is unknown whether fibrocartilage synthesized in vitro can maintain its altered phenotype without growth factor treatment. The action of Frzb-1 or specific upregulation of its expression could be useful in maintaining a chondrocytic phenotype in synoviocytes. This gene was expressed in all the adult synoviocytes in our study and was highest in the growth-factor–treated cells. This gene may be constitutively expressed in canine synovium, or it is possible that the dedifferentiation of synoviocytes in monolayer culture induced Frzb-1 expression. Given their ontogenetic chondrogenic role, it is not surprising that the growth factors used in the present study would increase expression of the Frzb-1 gene, which supports an immature chondrocytic phenotype. The precise role of the Frzb-1 gene in the formation of our synoviocyte tissue-like constructs remains unclear at this time, as the tissues did not resemble developing cartilage, but did possess cartilage-like rounded cell phenotypes and occasional cells within lacunae.

The present study had several weaknesses. The methods used to characterize joints as normal may not have detected subclinical or early Outerbridge grade 1 or 2 osteoarthritis lesions. Collection and histologic analysis of synovial biopsy specimens were not performed at the time of tissue harvest, which would have helped define the preexisting inflammatory status of the source tissues. Also, the outcome measures for collagen I and collagen II were not quantified. Additionally, comparative synthetic capabilities of canine chondrocytes and meniscal fibrochondrocytes were not studied, although it is unlikely that healthy autologous meniscofibrochondrocytes would be used in tissue engineering because of patient morbidity. Although we attempted to select donor osteoarthritic joints that had gross pathological changes of similar severity, the primary causes of osteoarthritis in the study joints were variable, including naturally occurring cranial cruciate degenerative disease with meniscal tears, experimental cranial cruciate ligament transection (Pond-Nuki model) with meniscal tears, and naturally occurring chronic shoulder joint instability. The autoimmune processes that are potentially associated with cranial cruciate ligament degeneration in dogs69 could support an enhanced inflammatory status of harvested synovial tissue. In addition, the duration of osteoarthritis was variable among dogs, with a range of 12 weeks to possibly years. Osteoarthritis is a temporally heterogeneous disease that has different cytokine, growth factor, and enzymatic profiles in the acute and chronic stages.70,71 In addition, adult human synovial stem cells have variable in vitro chondrogenic activity, despite normalization of joint location, primary disease, and patient age.35 Thus, the different causes of osteoarthritis, affected joints, duration of disease, and possibly individual variation in synoviocyte chondrogenic potential reduced the power of our study.

The results of the present study indicated that normal-joint synoviocytes may be a better cell source for meniscal tissue engineering purposes in dogs because synoviocytes from osteoarthritic joints had a comparatively diminished ability to produce collagenous ECM in the same monolayer culture environment. Synoviocytes from osteoarthritic joints had the highest expressions of SOX-9 and RGS-10 genes, thereby revealing a genomic potential for chondrogenesis. With growth factor stimulation, canine synoviocytes from normal and osteoarthritic joints can produce a tissue containing GAGs, collagen I, and small amounts of collagen II. These data provide justification for continued pursuit of synoviocyte-based tissue engineering strategies for fibrocartilage repair and regeneration.

ABBREVIATIONS

bFGF

Basic fibroblast growth factor

DMEM

Dulbecco modified Eagle medium ECM Extracellular matrix

Frzb

Frizzled-motif associated with bone development

GAG

Glycosaminoglycan

GAPDH

Glyceraldehyde-3-phosphate dehydrogenase

IGF-1

Insulin-like growth factor-1

RGS-10

Regulator of G-protein signaling

RT

Real-time reverse transcriptase

SOX-9

Sry-type homeobox protein-9

TGF-β1

Transforming growth factor-β1

a.

BD, Franklin Lakes, NJ.

b.

Type 1A clostridial collagenase, Sigma, St Louis, Mo.

c.

Human recombinant bFGF, BD Biosciences, Bedford, Mass.

d.

Human recombinant TGF-β, BD Biosciences, Bedford, Mass.

e.

Human recombinant IGF-1, BD Biosciences, Bedford, Mass.

f.

TrypLE Express, Innovative Cell Technologies, San Diego, Calif.

g.

Autostainer, Dakocytomation, Carpenteria, Calif.

h.

Anti-collagen type I antibodies, Chemicon, Temecula, Calif.

i.

Rabbit Envision LSAB+ system, Dakocytomation, Carpenteria, Calif.

j.

Nova Red, Chemicon, Temecular, Calif.

k.

Newcomer's Supply, Appleton, Wis.

l.

Carl Zeiss, Thornwood, NY.

m.

Olympus DP-70 digital camera, Olympus, Melville, NY.

n.

Quant-iT PicoGreen kit, Invitrogen, Paisley, Scotland.

o.

TE buffer, Invitrogen, Paisley, Scotland.

p.

Synergy HT–KC-4 spectrophotometric plate reader, BioTec, Winooski, Vt.

q.

FT-4 software, BioTec, Winooski, Vt.

r.

Primer EXE, The Whitehead Institute for Biomedical Research, Massachusetts Institute of Technology, Cambridge, Mass.

s.

RNALater, Ambion, Austin, Tex.

t.

Trizol reagent, Invitrogen, Carlsbad, Calif.

u.

StrataScript RT, Stratagene, La Jolla, Calif.

v.

GeneAmp PCR System 9700, Applied Biosystems Inc, Foster City, Calif.

w.

Light Cycler 480, Roche, Palo Alto, Calif.

x.

Quantitect SYBR green PCR kit, Qiagen, Valencia, Calif.

y.

SigmaStat, version 3.5, Jandel Scientific, San Rafael, Calif.

Appendix

Forward and reverse primers used in RT-PCR assays for detection of collagen I, collagen II, aggrecan, SOX-9, RGS-10, and Frzb-1 gene expressions in synoviocytes harvested from osteoarthritic and nonosteoarthritic (normal) joints of dogs.

Target genePrimerNucleotide sequence (5′→3′)
GAPDHForwardGTGACTTCAACAGTGACACC
 ReverseCCTTGGAGGCCATGTAGACC
Collagen IForwardTGCACGAGTCACACTGGAGC
 ReverseATGCCGAATTCCTGGTCTGG
Collagen IIForwardGGCCTGTCTGCTTCTTGTAA
 ReverseATCAGGTCAGGTCAGCCATT
AggrecanForwardATCGAAGGGGACTTCCGCTG
 ReverseATCACCACACAGTCCTCTCCG
Sox-9ForwardGAGAGCGAGGAGGACAAGTT
 ReverseGCTTGACGTGCGGCTTGTTC
RGS-10ForwardGAACCGAGGAAGAGGAAGAA
 ReverseCCTGCATGGTCCTGAGAGTG
Frzb-1ForwardAGCGTGCCAGATTACTGTTG
 ReverseGTGGAATCACTGTGGCTAGA

References

  • 1.

    Walmsley JR, Phillips TJ, Townsend HG. Meniscal tears in horses: an evaluation of clinical signs and arthroscopic treatment of 80 cases. Equine Vet J 2003; 35:402406.

    • Search Google Scholar
    • Export Citation
  • 2.

    Peroni JF, Stick JA. Evaluation of a cranial arthroscopic approach to the stifle joint for the treatment of femorotibial joint disease in horses: 23 cases (1998–1999). J Am Vet Med Assoc 2002; 220:10461052.

    • Search Google Scholar
    • Export Citation
  • 3.

    Jackson J, Vasseur PB, Griffey S, et al. Pathologic changes in grossly normal menisci in dogs with rupture of the cranial cruciate ligament. J Am Vet Med Assoc 2001; 218:12811284.

    • Search Google Scholar
    • Export Citation
  • 4.

    Ralphs SC, Whitney WO. Arthroscopic evaluation of menisci in dogs with cranial cruciate ligament injuries: 100 cases (1999–2000). J Am Vet Med Assoc 2002; 221:16011604.

    • Search Google Scholar
    • Export Citation
  • 5.

    Johnson KA, Francis DJ, Manley PA, et al. Comparison of the effects of caudal pole hemi-meniscectomy and complete medial meniscectomy in the canine stifle joint. Am J Vet Res 2004; 65:10531060.

    • Search Google Scholar
    • Export Citation
  • 6.

    Burks RT, Metcalf MH, Metcalf RW. Fifteen-year follow-up of arthroscopic partial meniscectomy. Arthroscopy 1997; 13:673679.

  • 7.

    Arnoczky SP, Warren RF. The microvasculature of the meniscus and its response to injury. An experimental study in the dog. Am J Sports Med 1983; 11:131141.

    • Search Google Scholar
    • Export Citation
  • 8.

    Kobayashi K, Fujimoto E, Deie M, et al. Regional differences in the healing potential of the meniscus—an organ culture model to eliminate the influence of microvasculature and the synovium. Knee 2004; 11:271278.

    • Search Google Scholar
    • Export Citation
  • 9.

    Arnoczky SP, Warren RF, Spivak JM. Meniscal repair using an exogenous fibrin clot. An experimental study in dogs. J Bone Joint Surg Am 1988; 70:12091217.

    • Search Google Scholar
    • Export Citation
  • 10.

    Okuda K, Ochi M, Shu N, et al. Meniscal rasping for repair of meniscal tear in the avascular zone. Arthroscopy 1999; 15:281286.

  • 11.

    Peretti GM, Gill TJ, Xu JW, et al. Cell-based therapy for meniscal repair: a large animal study. Am J Sports Med 2004; 32:146158.

  • 12.

    Klompmaker J, Veth RP, Jansen HW, et al. Meniscal repair by fibrocartilage in the dog: characterization of the repair tissue and the role of vascularity. Biomaterials 1996; 17:16851691.

    • Search Google Scholar
    • Export Citation
  • 13.

    Klompmaker J, Veth RP, Jansen HW, et al. Meniscal replacement using a porous polymer prosthesis: a preliminary study in the dog. Biomaterials 1996; 17:11691175.

    • Search Google Scholar
    • Export Citation
  • 14.

    de Groot JH, de Vrijer R, Pennings AJ, et al. Use of porous polyurethanes for meniscal reconstruction and meniscal prostheses. Biomaterials 1996; 17:163173.

    • Search Google Scholar
    • Export Citation
  • 15.

    Cook JL, Tomlinson JL, Kreeger JM, et al. Induction of meniscal regeneration in dogs using a novel biomaterial. Am J Sports Med 1999; 27:658665.

    • Search Google Scholar
    • Export Citation
  • 16.

    Stone KR, Steadman JR, Rodkey WG, et al. Regeneration of meniscal cartilage with use of a collagen scaffold. Analysis of preliminary data. J Bone Joint Surg Am 1997; 79:17701777.

    • Search Google Scholar
    • Export Citation
  • 17.

    Cox JS, Nye CE, Schaefer WW, et al. The degenerative effects of partial and total resection of the medial meniscus in dogs' knees. Clin Orthop Relat Res1975;178183.

    • Search Google Scholar
    • Export Citation
  • 18.

    Nishimura K, Solchaga LA, Caplan AI, et al. Chondroprogenitor cells of synovial tissue. Arthritis Rheum 1999; 42:26312637.

  • 19.

    Pei M, He F, Kish VL, et al. Engineering of functional cartilage tissue using stem cells from synovial lining: a preliminary study. Clin Orthop Relat Res 2008; 466:18801889.

    • Search Google Scholar
    • Export Citation
  • 20.

    Shintani N, Hunziker EB. Chondrogenic differentiation of bovine synovium: bone morphogenetic proteins 2 and 7 and transforming growth factor beta1 induce the formation of different types of cartilaginous tissue. Arthritis Rheum 2007; 56:18691879.

    • Search Google Scholar
    • Export Citation
  • 21.

    Yoshimura H, Muneta T, Nimura A, et al. Comparison of rat mesenchymal stem cells derived from bone marrow, synovium, periosteum, adipose tissue, and muscle. Cell Tissue Res 2007; 327:449462.

    • Search Google Scholar
    • Export Citation
  • 22.

    Park Y, Sugimoto M, Watrin A, et al. BMP-2 induces the expression of chondrocyte-specific genes in bovine synovium-derived progenitor cells cultured in three-dimensional alginate hydrogel. Osteoarthritis Cartilage 2005; 13:527536.

    • Search Google Scholar
    • Export Citation
  • 23.

    Arnoczky SP, Warren RF, Kaplan N. Meniscal remodeling following partial meniscectomy—an experimental study in the dog. Arthroscopy 1985; 1:247252.

    • Search Google Scholar
    • Export Citation
  • 24.

    Lindhorst E, Vail TP, Guilak F, et al. Longitudinal characterization of synovial fluid biomarkers in the canine meniscectomy model of osteoarthritis. J Orthop Res 2000; 18:269280.

    • Search Google Scholar
    • Export Citation
  • 25.

    Smith GN, Mickler EA, Albrecht ME, et al. Severity of medial meniscus damage in the canine knee after anterior cruciate ligament transection. Osteoarthritis Cartilage 2002; 10:321326.

    • Search Google Scholar
    • Export Citation
  • 26.

    van Tienen TG, Heijkants RG, de Groot JH, et al. Presence and mechanism of knee articular cartilage degeneration after meniscal reconstruction in dogs. Osteoarthritis Cartilage 2003; 11:7884.

    • Search Google Scholar
    • Export Citation
  • 27.

    Wyland DJ, Guilak F, Elliott DM, et al. Chondropathy after meniscal tear or partial meniscectomy in a canine model. J Orthop Res 2002; 20:9961002.

    • Search Google Scholar
    • Export Citation
  • 28.

    Ochi M, Ishida O, Daisaku H, et al. Immune response to fresh meniscal allografts in mice. J Surg Res 1995; 58:478484.

  • 29.

    Rodeo SA, Seneviratne A, Suzuki K, et al. Histological analysis of human meniscal allografts. A preliminary report. J Bone Joint Surg Am 2000; 82:10711082.

    • Search Google Scholar
    • Export Citation
  • 30.

    Pessina A, Bonomi A, Baglio C, et al. Microbiological risk assessment in stem cell manipulation. Crit Rev Microbiol 2008; 34:112.

  • 31.

    Graf KW Jr, Sekiya JK, Wojtys EM. Long-term results after combined medial meniscal allograft transplantation and anterior cruciate ligament reconstruction: minimum 8.5-year follow-up study. Arthroscopy 2004; 20:129140.

    • Search Google Scholar
    • Export Citation
  • 32.

    Jeffreys TE. Synovial chondromatosis. J Bone Joint Surg Br 1967; 49:530534.

  • 33.

    Blom AB, van Lent PL, Holthuysen AE, et al. Synovial lining macrophages mediate osteophyte formation during experimental osteoarthritis. Osteoarthritis Cartilage 2004; 12:627635.

    • Search Google Scholar
    • Export Citation
  • 34.

    van Lent PL, Blom AB, van der Kraan P, et al. Crucial role of synovial lining macrophages in the promotion of transforming growth factor β-mediated osteophyte formation. Arthritis Rheum 2004; 50:103111.

    • Search Google Scholar
    • Export Citation
  • 35.

    Giurea A, Ruger BM, Hollemann D, et al. STRO-1+ mesenchymal precursor cells located in synovial surface projections of patients with osteoarthritis. Osteoarthritis Cartilage 2006; 14:938943.

    • Search Google Scholar
    • Export Citation
  • 36.

    Mussener A, Funa K, Kleinau S, et al. Dynamic expression of transforming growth factor-betas (TGF-β) and their type I and type II receptors in the synovial tissue of arthritic rats. Clin Exp Immunol 1997; 107:112119.

    • Search Google Scholar
    • Export Citation
  • 37.

    Acosta CA, Izal I, Ripalda P, et al. Gene expression and proliferation analysis in young, aged, and osteoarthritic sheep chondrocytes effect of growth factor treatment. J Orthop Res 2006; 24:20872094.

    • Search Google Scholar
    • Export Citation
  • 38.

    Murphy JM, Dixon K, Beck S, et al. Reduced chondrogenic and adipogenic activity of mesenchymal stem cells from patients with advanced osteoarthritis. Arthritis Rheum 2002; 46:704713.

    • Search Google Scholar
    • Export Citation
  • 39.

    Outerbridge RE. The etiology of chondromalacia patellae. J Bone Joint Surg Br 1961; 43:752757.

  • 40.

    Pei M, He F, Vunjak-Novakovic G. Synovium-derived stem cell-based chondrogenesis. Differentiation 2008; 76:10441056.

  • 41.

    Farndale RW, Buttle DJ, Barrett AJ. Improved quantitation and discrimination of sulphated glycosaminoglycans by use of dimethylmethylene blue. Biochim Biophys Acta 1986; 883:173177.

    • Search Google Scholar
    • Export Citation
  • 42.

    Reddy GK, Enwemeka CS. A simplified method for the analysis of hydroxyproline in biological tissues. Clin Biochem 1996; 29:225229.

  • 43.

    Kuboki Y, Mechanic GL. The distribution of δ,δ'-dihydroxylysinonorleucine in bovine tendon and dentin. Connect Tissue Res 1974; 2:223230.

    • Search Google Scholar
    • Export Citation
  • 44.

    Reno C, Marchuk L, Sciore P, et al. Rapid isolation of total RNA from small samples of hypocellular, dense connective tissues. Biotechniques 1997; 22:10821086.

    • Search Google Scholar
    • Export Citation
  • 45.

    Pfaffl MW, Horgan GW, Dempfle L. Relative expression software tool (REST) for group-wise comparison and statistical analysis of relative expression results in real-time PCR. Nucleic Acids Res 2002; 30:e36e46.

    • Search Google Scholar
    • Export Citation
  • 46.

    Melrose J, Smith S, Cake M, et al. Comparative spatial and temporal localisation of perlecan, aggrecan and type I, II and IV collagen in the ovine meniscus: an ageing study. Histochem Cell Biol 2005; 124:225235.

    • Search Google Scholar
    • Export Citation
  • 47.

    Kambic HE, McDevitt CA. Spatial organization of types I and II collagen in the canine meniscus. J Orthop Res 2005; 23:142149.

  • 48.

    Stephan JS, McLaughlin RM Jr, Griffith G. Water content and glycosaminoglycan disaccharide concentration of the canine meniscus. Am J Vet Res 1998; 59:213216.

    • Search Google Scholar
    • Export Citation
  • 49.

    Sureshbabu A, Okajima H, Yamanaka D, et al. IGFBP-5 induces epithelial and fibroblast responses consistent with the fibrotic response. Biochem Soc Trans 2009; 37:882885.

    • Search Google Scholar
    • Export Citation
  • 50.

    Leask A, Abraham DJ. TGF-β signaling and the fibrotic response. FASEB J 2004; 18:816827.

  • 51.

    Molloy T, Wang Y, Murrell G. The roles of growth factors in tendon and ligament healing. Sports Med 2003; 33:381394.

  • 52.

    Wilkinson LS, Pitsillides AA, Worrall JG, et al. Light microscopic characterization of the fibroblast-like synovial intimal cell (synoviocyte). Arthritis Rheum 1992; 35:11791184.

    • Search Google Scholar
    • Export Citation
  • 53.

    Xu H, Edwards J, Banerji S, et al. Distribution of lymphatic vessels in normal and arthritic human synovial tissues. Ann Rheum Dis 2003; 62:12271229.

    • Search Google Scholar
    • Export Citation
  • 54.

    Pei M, Seidel J, Vunjak-Novakovic G, et al. Growth factors for sequential cellular de- and re-differentiation in tissue engineering. Biochem Biophys Res Commun 2002; 294:149154.

    • Search Google Scholar
    • Export Citation
  • 55.

    Pei M, Luo J, Chen Q. Enhancing and maintaining chondrogenesis of synovial fibroblasts by cartilage extracellular matrix protein matrilins. Osteoarthritis Cartilage 2008; 16:11101117.

    • Search Google Scholar
    • Export Citation
  • 56.

    Hoben GM, Hu JC, James RA, et al. Self-assembly of fibrochon-drocytes and chondrocytes for tissue engineering of the knee meniscus. Tissue Eng 2007; 13:939946.

    • Search Google Scholar
    • Export Citation
  • 57.

    Pazzano D, Mercier K, Moran J, et al. Comparison of chondrogenesis in static and perfused bioreactor culture. Biotechnol Prog 2000; 16:893896.

    • Search Google Scholar
    • Export Citation
  • 58.

    Smith RL, Donlon BS, Gupta MK, et al. Effects of fluid induced shear on articular chondrocyte morphology and metabolism in vitro. J Orthop Res 1995; 13:824831.

    • Search Google Scholar
    • Export Citation
  • 59.

    Davisson T, Sah RL, Ratcliffe A. Perfusion increases cell content and matrix synthesis in chondrocyte three-dimensional cultures. Tissue Eng 2002; 8:807816.

    • Search Google Scholar
    • Export Citation
  • 60.

    Fiorito S, Magrini L, Adrey J, et al. Inflammatory status and cartilage regenerative potential of synovial fibroblasts from patients with osteoarthritis and chondropathy. Rheumatology (Oxford) 2005; 44:164171.

    • Search Google Scholar
    • Export Citation
  • 61.

    Klocke NW, Snyder PW, Widmer WR, et al. Detection of synovial macrophages in the joint capsule of dogs with naturally occurring rupture of the cranial cruciate ligament. Am J Vet Res 2005; 66:493499.

    • Search Google Scholar
    • Export Citation
  • 62.

    Krey PR, Scheinberg MA, Cohen AS. Fine structural analysis of rabbit synovial cells. II. Fine structure and rosette-forming cells of explant and monolayer cultures. Arthritis Rheum 1976; 19:581592.

    • Search Google Scholar
    • Export Citation
  • 63.

    Sutton S, Clutterbuck A, Harris P, et al. The contribution of the synovium, synovial derived inflammatory cytokines and neuropeptides to the pathogenesis of osteoarthritis. Vet J 2009; 179:1024.

    • Search Google Scholar
    • Export Citation
  • 64.

    Bondeson J, Wainwright SD, Lauder S, et al. The role of synovial macrophages and macrophage-produced cytokines in driving aggrecanases, matrix metalloproteinases, and other destructive and inflammatory responses in osteoarthritis. Arthritis Res Ther 2006; 8:R187R199.

    • Search Google Scholar
    • Export Citation
  • 65.

    Lefebvre V, Huang W, Harley VR, et al. SOX9 is a potent activator of the chondrocyte-specific enhancer of the pro alpha1(II) collagen gene. Mol Cell Biol 1997; 17:23362346.

    • Search Google Scholar
    • Export Citation
  • 66.

    Bi W, Deng JM, Zhang Z, et al. Sox9 is required for cartilage formation. Nat Genet 1999; 22:8589.

  • 67.

    Appleton CT, James CG, Beier F. Regulator of G-protein signaling (RGS) proteins differentially control chondrocyte differentiation. J Cell Physiol 2006; 207:735745.

    • Search Google Scholar
    • Export Citation
  • 68.

    Enomoto-Iwamoto M, Kitagaki J, Koyama E, et al. The Wnt antagonist Frzb-1 regulates chondrocyte maturation and long bone development during limb skeletogenesis. Dev Biol 2002; 251:142156.

    • Search Google Scholar
    • Export Citation
  • 69.

    Doom M, de Bruin T, de Rooster H, et al. Immunopathological mechanisms in dogs with rupture of the cranial cruciate ligament. Vet Immunol Immunopathol 2008; 125:143161.

    • Search Google Scholar
    • Export Citation
  • 70.

    Benito MJ, Veale DJ, FitzGerald O, et al. Synovial tissue inflammation in early and late osteoarthritis. Ann Rheum Dis 2005; 64:12631267.

    • Search Google Scholar
    • Export Citation
  • 71.

    Smith MD, Triantafillou S, Parker A, et al. Synovial membrane inflammation and cytokine production in patients with early osteoarthritis. J Rheumatol 1997; 24:365371.

    • Search Google Scholar
    • Export Citation

Contributor Notes

Address correspondence to Dr. Warnock (jennifer.warnock@oregonstate.edu).

Dr. Warnock's present address is Department of Clinical Sciences, College of Veterinary Medicine, Oregon State University, Corvallis, OR 97331.

This manuscript represents a portion of a dissertation submitted by the first author to the Department of Veterinary Pathobiology, College of Veterinary Medicine, University of Missouri, as partial fulfillment of the requirements for a Doctor of Philosophy degree.

Supported by the Veterinary Orthopedic Society Hohn-Johnson Research Grant and the Comparative Orthopaedic Laboratory.

Presented in part as an oral presentation at the Veterinary Orthopedic Society Conference, Big Sky, Mont, March 2008.