In adults, cartilage has limited potential for natural repair and thus injuries typically result in the progression of osteoarthritis.1 The study of osteoarthritis is of particular interest in equine medicine because joint injury and joint disease are major causes of lameness2 and consequent wastage of athletic horses.3 Histologic lesions associated with osteoarthritis can include fissure formation in the articular cartilage surface, chondrocyte cell death, focal cell loss, chondrocyte cluster formation, and loss of ECM such as proteoglycans and collagen fibrils.2,4,5
In vitro models of cartilage injury can enable investigation of the responses of cartilage to traumatic injury in a highly controlled and reproducible manner. Additionally, well-developed in vitro models allow for high throughput and inexpensive preliminary testing of approaches to treat and prevent the progression of osteoarthritis.
Many in vitro models of cartilage injury exist and have been summarized in literature reviews.6,7 Investigators have used adult and immature cartilage explants from various animal species, including humans. Immature cartilage tissue subjected to injurious compression has been used to gain information regarding traumatic injury in immature human knees.8 Studies performed with adult articular cartilage subjected to injurious loads in vitro have relevance to early, middle, and later stages of osteoarthritis. These changes in cartilage in vitro have been evaluated and compared with results of animal studies9–14 in which impact loads have been directly applied to joint cartilages in vivo.
The purpose of the study reported here was to develop an in vitro model of cartilage injury in full-thickness cartilage specimens (containing the superficial, middle, and deep cartilage zones) extracted from the stifle joint of adult horses by use of a computer-controlled single-impact model of unconfined compression.15–18 Our objective was to determine the degree of strain sufficient to reproducibly induce pathological change in cartilage that would mimic early osteoarthritic changes in vivo.
Materials and Methods
Specimens—Cartilage specimens were extracted from the stifle joints of 6 equine cadavers within 16 hours after death. The horses had been between 3 and 10 years of age and had died for reasons unrelated to the musculoskeletal system and unrelated to this study.
A mosaicplasty osteochondral grafting systema was used to extract at least 15 osteochondral plugs (diameter, 4.5 mm) that included the intact superficial cartilage zone from the proximal region of the trochlear ridges of the left and right femurs from each horse. Specimens from each horse were pooled. Cartilage was removed from the subchondral bone at the calcified and noncalcified cartilage junction immediately after extraction from the joint, followed by incubation in growth medium (Dulbecco modified Eagle medium, 10% fetal bovine serum [vol/vol], ascorbic acid [20 μg/mL], 1% penicillin and streptomycin [vol/vol], 10mM HEPES, 0.1mM nonessential amino acids, 0.4mM proline, and amphotericin B [0.25 μg/mL]) for 48 hours prior to injury.
Mechanical injury induction—Just prior to injury induction, the thickness of each cartilage plug (plane perpendicular to the articular surface) was measured to 0.01-mm accuracy with digital calipers. For each horse, 3 cartilage specimens were assigned to each of 5 groups: 1 free-swell control and 4 injury groups at various degrees of strain.
Cartilage plugs assigned to injury groups were removed from medium and placed into a sterile polysulphone loading chamber consisting of a well aligned co-axially with an impermeable platen (diameter, 10 mm) in the absence of medium (Figure 1). A test systemb was used to apply an initial compressive tare load of 2 N. After creep equilibrium was attained, an injurious compression at a rate of 100% strain/s was applied until 50%, 60%, 70%, or 80% final cartilage strain was achieved (Figure 2). Compression was then released at the same rate, and plugs were immediately placed in culture medium for 28 days. All medium was changed at similar intervals (every 2 to 3 days). Control plugs were kept in medium throughout the study, which was changed at a similar rate.
Photograph of a polysulphone loading chamber used to evaluate the effects of injurious compression on cartilage specimens from the trochlear ridge of the femur from an adult horse. For injury induction, a cartilage specimen was placed into the well of the chamber directly under the platen.
Citation: American Journal of Veterinary Research 74, 1; 10.2460/ajvr.74.1.40
Representative stress-strain curve showing the results of a testing machine used to load and induce compressive injury in the cartilage specimens described in Figure 1.
Citation: American Journal of Veterinary Research 74, 1; 10.2460/ajvr.74.1.40
Twenty-eight days after cartilage injury, all cartilage plugs, including those in the control group, were examined grossly and microscopically and digital photographic images were obtained to document the integrity of the articular surface and presence of fissures. Specimens were sectioned across the diameter perpendicular to the fissures when fissures were evident or arbitrarily across when not. Half of each specimen was placed into neutral-buffered 10% formalin for fixation. The remaining half was embedded in cryoembedding medium,c snap-frozen in liquid nitrogen, and stored at −80°C for further use.
Histologic evaluation—Formalin-fixed specimens were processed, paraffin embedded, and sectioned (thickness, 5 μm) with a microtome, then transferred to slides. One slide (containing 2 tissue sections) was evaluated for each stain used. Four sections each from an injured and control specimen were stained with H&E to evaluate cellular changes or SOFG to identify changes in GAG content. Digital images of SOFG-stained sections and image management softwared were used to measure fissure size, defined as the area within the fissure, and fissure depth, determined as the depth of the fissure as a percentage of the total height of each specimen. The software was calibrated to the microscope and camera.
IHC evaluation—Immunohistochemical staining was performed to detect aggrecan and collagen (type I to assess repair and type II to assess unaffected matrix). Briefly, frozen embedded specimens were sectioned (thickness, 8 μm) with a cryostat and mounted onto adhesive slides.e Mouse antibodies raised against aggrecan at a 1:20 dilutionf or collagen types I and II by use of undiluted supernatantg were applied, followed by goat anti-mouse secondary antibody conjugated with horseradish peroxidase at a 1:500 dilution.h For negative control specimens, additional sections were probed with mouse serum at a concentration equal to that of the primary antibody.
Scoring of cartilage sections—All histologic and IHC sections were scored with the aid of a grading scale for detailing the severity of osteoarthritic characteristics in equine cartilage,19 with minor modifications. Necrotic nuclei were not present in the cartilage sections; instead, there were empty lacunae attributed to chondroptosis, which is a process of chondrocyte cell death that results in the presence of an empty lacuna.20 Therefore, as opposed to evaluating chondrocyte necrosis, chondrocyte cell death was evaluated and defined by the presence of lacunae lacking discernible nuclei. Focal cell loss was measured in a manner similar to the established grading scale but defined as the presence of distinct regions devoid of discernible nuclei, and chondrocyte cluster formation (lacunae containing chondrocytes with > 1 nucleus) was evaluated as defined.19 All specimens were separately graded for each osteoarthritis characteristic in the superficial, middle, and deep regions as well as at the region adjacent to the fissure (defined as the region 3 times the area of the largest fissure).
In brief, each region was assigned a grade for each osteoarthritis characteristic that corresponded to the severity by which the specimen deviated in histologic appearance from undamaged articular cartilage as follows: 0 = no change or abnormality, 1 = slight deviance, 2 = mild deviance, and 3 = moderate deviance (the grade for severe deviance [4] was not used). For each osteoarthritis characteristic, scores for each region (superficial, middle, deep, and fissures) were evaluated separately or, to gain an understanding of the section as a whole, were summed together for each slide to provide a cumulative score with a maximum score of 12. All slides were graded by the consensus of 2 evaluators (CML and DDF), who were unaware of group assignment.
Histologic and IHC-stained sections were used to evaluate changes in the ECM. The SOFG method is an indirect method for identifying the presence of GAG because GAG chains in healthy articular cartilage are negatively charged and the SOFG stain is cationic. A decrease in the degree of SOFG staining was interpreted as indicating a decrease in GAG content, and thus the SOFG-stained sections were used to identify any regions with less GAG content than in control specimens. To identify ECM molecules more specifically IHC-stained sections were used to assess any changes in aggrecan, collagen type II, and collagen type I content, compared with the content in control sections.
Statistical analysis—Statistical analysis was conducted to determine differences in IHC and histologic staining results accounting for both injury and region, with horse as a random variable in a generalized linear mixed model.i This method of analysis allowed for intra-group comparisons as well as the ability to compare static culture with the various injury groups to account for any effects static culture may have had. Predictive F values were used to identify statistical differences among injury and control groups or among cartilage regions, and specific comparisons (when indicated by a significant F test result) were made with a least squares means procedure. Values of P < 0.05 were considered significant for all analyses.
Results
Pathological changes in the cartilage matrix—The mean ± SD height of the full-thickness cartilage specimens obtained from the trochlear ridge of femurs from adult horses was summarized (Table 1). Peak stress was significantly (P < 0.001) affected by strain, ranging from 11.39 ± 5.55 MPa for cartilages specimens subjected to 50% strain to 22.29 ± 4.74 MPa for specimens subjected to 80% strain. Compression at 60% and 70% strain induced similar stresses of 14.68 ± 5.55 MPa and 16.02 ± 5.32 MPa, respectively. All specimens developed fissures when compressed to 70% strain. Most of the specimens developed fissures when compressed to 60% and 80% strain (14/18 and 7/9, respectively), and 9 of 18 developed fissures when compressed to 50% strain.
Mean ± SD values of characteristics of full-thickness cartilage specimens from the trochlear ridge of femurs from 6 adult horses that were subjected to various degrees of strain.
| Strain | |||||
|---|---|---|---|---|---|
| Variable | 50% (n = 18) | 60% (n = 18) | 70% (n = 13) | 80% (n = 9) | P value |
| Specimen height (mm) | 1.65 ± 0.55a | 1.73 ± 0.55a | 1.73 ± 0.50a | 1.72 ± 0.45a | 0.89 |
| Stress achieved (MPa) | 11.39 ± 5.55a | 14.68 ± 5.55b | 16.02 ± 5.32b | 22.29 ± 4.74c | < 0.001 |
| Fissure rate (%) | 50a | 78b | 100c | 78b | < 0.001 |
| Fissure size (μm2) | 309.11 ± 1,036.51a | 619.14 ± 756.96a,b | 1,031.16 ± 690.37b,c | 1,595.40 ± 846.84c | 0.01 |
| Fissure depth (μm) | 16.07 ± 14.12a | 23.93 ± 10.01a,b | 25.54 ± 9.07b | 29.19 ± 11.55b | 0.07 |
Values with different superscript letters differ significantly (P < 0.05) from each other.
Fissure size and depth were significantly smaller in specimens compressed to 50% than in those compressed to 70% strain (P = 0.026 and P = 0.030, respectively) and 80% strain (P = 0.002 and P = 0.014, respectively). All specimens compressed to 50% and 60% strain remained acceptable for use in the study; however, some specimens in the 70% and 80% strain groups were lost during compression because the maximum load reached during compression exceeded the capacity of the 500-N load cell.
Strain and pathological change—Compared with control specimens, GAG content was significantly (all P < 0.001) lower in specimens compressed to 60%, 70%, and 80% and type II collagen content was significantly (P = 0.003) lower in specimens compressed to 60% strain (Table 2). No effect of compression on type I collagen or aggrecan content was identified. Compression also had a significant impact on cellular change. Compression to ≥ 60% strain induced a significant increase in chondrocyte cell death, focal cell loss, and chondrocyte cluster formation, compared with values in control specimens and specimens compressed to 50% strain. When scores for all cellular changes were cumulatively evaluated, all compressed specimens had more severe total cellular pathological scores than did control specimens, with severity increasing with increasing degree of strain. In summary, specimens compressed to ≥ 60% strain had the greatest amount of macroscopic and microscopic cellular pathological change, compared with control specimens (Figure 3).
Representative microphotographs of uninjured (control) cartilage sections from the trochlear ridge of adult horses and sections injured by mechanical compression at 50%, 60%, 70%, or 80% strain. Images obtained at 10× resolution show tissue damage such as fissures in 60%, 70%, and 80% strain groups. Images obtained at 20× resolution show cellular pathological changes such as chondrocyte cluster formation in the 60% and 70% strain specimens and cell death in the superficial cartilage region of 60%, 70%, and 80% specimens. Images on the left are the regions marked by the square in the 20× resolution images, which have been enlarged for visual clarity. H&E stain; bar = 100 μm.
Citation: American Journal of Veterinary Research 74, 1; 10.2460/ajvr.74.1.40
Mean ± SD cumulative lesion severity scores for superficial, middle, and deep cartilage layers and fissures in full-thickness cartilage specimens from the horses in Table 1 that were or were not (control) subjected to mechanical compression at various degrees of strain.
| Strain | ||||||
|---|---|---|---|---|---|---|
| Injury type | Control (n = 18) | 50% (n = 18) | 60% (n = 18) | 70% (n = 13) | 80% (n = 9) | P value |
| GAG | 8.5 ± 1.95a | 9.3 ± 1.95a | 10.9 ± 1.91b | 11 ± 1.77b | 11 ± 1.47b | < 0.001 |
| Collagen type II | 1.0 ± 4.03a | 2.9 ± 5.19a,b | 4.6 ± 3.82b | 3.3 ± 1.77a,b | 2.7 ± 3.9a,b | 0.05 |
| Collagen type I | 0 ± 0.59a,b | 0 ± 0.59a | 0 ± 0.55a | 0.2 ± 0.54a,b | 0.4 ± 0.54b | 0.24 |
| Aggrecan | 3.3 ± 3.9a | 3.6 ± 3.77a | 5.2 ± 3.69a | 4.8 ± 3.43a | 4.8 ± 4.38a | 0.53 |
| Cell death | 3.3 ± 2.42a | 4.7 ± 2.42a | 7.3 ± 2.37b | 7.8 ± 2.27b | 8.5 ± 2.43b | < 0.001 |
| Focal cell loss | 0.79 ± 2.46a | 1.9 ± 2.46a | 4.4 ± 2.42b | 5.6 ± 2.27b | 8.6 ± 2.43a | < 0.001 |
| Cluster formation | 0.6 ± 2.20a | 8.6 ± 2.20a,b | 3.9 ± 2.12c | 4.4 ± 2.06c | 2.8 ± 2.22b,c | < 0.001 |
| Total severity | 4.7 ± 5.51a | 8.6 ± 5.51b | 15.6 ± 5.51c | 17.8 ± 5.05c,d | 20 ± 5.4d | < 0.001 |
A score of 0 corresponds to unaffected, healthy cartilage, whereas a score of 12 corresponds to moderate severity of pathological change.
Within a row, values with different superscript letters differ significantly (P < 0.05) from each other.
Regional pathological change—Chondrocyte cluster formation was significantly affected by region and injury (Table 3). For all injury groups, there was a significant increase in cluster formation in the deep region, compared with in all other regions. Injury induced through 60% strain was the only injury group with a significant difference in all 3 regions of cartilage. Independent of injury, severity of changes characteristic of osteoarthritis differed significantly by region when all specimens (control and 50%, 60%, 70%, and 80% strain) were analyzed as 1 group for evaluation of regional (superficial, middle, and deep cartilage) differences. Glycosaminoglycan and aggrecan content were most severely decreased in the superficial region, compared with content in other regions (P < 0.001 for both comparisons; Figure 4). Chondrocyte cell death was most severe in the superficial region (P = 0.002), whereas focal cell loss was most pronounced in the superficial and deep regions alike (P < 0.001). Chondrocyte cluster formation was also most severe in the deep region (P < 0.001). When evaluated cumulatively, total cellular pathological change was significantly (P < 0.001) more severe in the superficial and deep regions than in the middle region.
Mean ± SD cumulative lesion severity scores for superficial (n = 76), middle (76), and deep (76) layers of cartilage specimens from 6 horses, regardless of whether the specimens underwent mechanical compression. A score of 0 corresponds to unaffected, healthy cartilage, and a score of 3 corresponds to moderate pathological change. Total severity represents the combination of scores for cell death, focal cell loss, and chondrocyte cluster formation. a–cWithin a variable, values with different letters are significantly (P < 0.05) different.
Citation: American Journal of Veterinary Research 74, 1; 10.2460/ajvr.74.1.40
Mean ± SD cumulative lesion severity scores for superficial (n = 43), middle (43), and deep (43) layers and for fissures (43) of cartilage specimens that developed fissures after mechanical compression. See Figure 4 for remainder of key.
Citation: American Journal of Veterinary Research 74, 1; 10.2460/ajvr.74.1.40
Mean ± SD severity scores for chondrocyte cluster formation by injury or strain in the cartilage specimens described in Table 2.
| Strain | |||||
|---|---|---|---|---|---|
| Collagen region | Control (n = 18) | 50% (n = 18) | 60% (n = 18) | 70% (n = 13) | 80% (n = 9) |
| Superficial | 0.2 ± 0.76a | 0.5 ± 0.76a,b,c | 0.8 ± 0.76b,c | 0.4 ± 0.72a,b,c | 0.1 ± 3.26a |
| Middle | 0.1 ± 0.76a | 0.4 ± 0.76a,b,c | 0.8 ± 0.76b,c | 1.0 ± 0.72b | 0.9 ± 3.26b,c |
| Deep | 0.3 ± 0.76a,c | 0.9 ± 0.76b | 1.5 ± 0.76d,e | 2.0 ± 0.72d | 1.0 ± 3.26b,e |
A score of 0 corresponds to unaffected, healthy cartilage, whereas a score of 3 corresponds to moderate cluster formation.
Within a row, individual injury scores with different superscript letters differ significantly (P < 0.05) from each other.
In addition, specimens with fissures were analyzed separately to determine whether the severity of osteoarthritis characteristics in the region around the fissure differed significantly from the severity in other regions. In all comparisons, the fissure region had the same severity of osteoarthritis characteristics as did the superficial region, except for chondrocyte cluster formation, which was more severe in the middle region than in the superficial zone (Figure 5).
Discussion
Injury to adult articular cartilage commonly leads to the degeneration of articular cartilage and the progression of osteoarthritis. Indeed, mechanical injury can induce matrix damage,11,13,21–25 chondrocyte death,13,15,16,23–33 and chondrocyte cluster formation,34 all of which are hallmark characteristics of osteoarthritis.
The goal of the present study was to induce and detect histologic changes in equine cartilage that emulate the natural changes found in osteoarthritis, with interest in determining regional histologic characteristics. Full-thickness cartilage explants harvested from the trochlear ridges of the distal aspect of the femur of adult horses were injured by an unconfined compression to 50%, 60%, 70%, or 80% strain and evaluated after the explants had been incubated for 28 days in culture medium for histologic changes characteristic of early osteoarthritis. The data acquired demonstrated that histologic change can be produced that is consistent with previous cartilage injury models typical of early osteoarthritis in vivo.4 For adult full-thickness cartilage from the trochlear ridge, it was necessary to compress explants by at least 60% strain, resulting in peak stresses > 14.68 ± 5.56 MPa, to induce significant pathological changes, which included GAG loss, chondrocyte cell death, focal cell loss, and chondrocyte cluster formation.
The mechanical properties of cartilage are largely determined by the composition of the collagen-proteoglycan matrix,35,36 which in mature articular cartilage differs by cartilage region. The superficial zone has a relatively low proteoglycan content, and the collagen fibers are oriented parallel to the articular surface.37–39 The middle zone has the highest proteoglycan content and collagen fibers that are randomly oriented.40–42 The deep zone is also rich in proteoglycan content, but the collagen fibers are oriented perpendicular to the tidemark and subchondral bone.41,43
During unconfined compression, the collagen network is mainly responsible for controlling instantaneous deformation, whereas proteoglycans contribute to stiffness through regulation of osmotic pressure.44 As one would expect given the differential architecture, compressive load induces region-specific bending or crimping patterns of collagen fibers,45–47 which are associated with region-specific chondrocyte deformation.48 Therefore, we examined each region of cartilage independently and cumulatively to determine the effects of each degree of strain on ECM and cellular pathological change.
Chondrocyte cluster formation was the only histologic characteristic affected by compression magnitude and region. However, when pathological change was evaluated independent of compression magnitude, significant regional differences in severity became evident. Similar to what others have found,37 for most variables, the pathological change was most severe in the superficial region, compared with in other regions. This finding was not surprising given that the superficial region is more compliant under rapid loading, with a lower compressive modulus and greater permeability than other regions of the tissue.49 Indeed, when load is applied to cartilage of intact joints, most of the collagen fiber deformation occurs in the superficial zone, and with in vivo investigations, pathological change is also more concentrated in the region closest to the impact.14
Cartilage injury and posttraumatic osteoarthritis result in a loss of proteoglycans and type II collagen from the ECM by direct damage23,50 and cell-mediated degradation.51,52 As expected and as was found in another cartilage impact study,53 we observed a significant decrease in GAG content, predominantly in the superficial zone, when specimens were injured at ≥ 60% strain. Immunohistochemical staining for the proteoglycan aggrecan did not reveal a significant decrease of the substance in injured versus control specimens.
A cationic dye, SOFG is commonly used to detect the presence of negatively charged GAGs, whereas IHC is a more specific approach to detect the protein of interest, which in our situation was aggrecan. The disagreement between results for the SOFG stain and IHC technique may have been attributable to the difference in specificity and sensitivity of each staining method and indicates the importance of using both techniques when evaluating regional changes in GAG content.
Although there was no significant difference in type II collagen content among injury groups, compression at 60% strain caused a significant decrease, compared with type II collagen content in control specimens. In a previous in vivo study14 of cartilage injury by single-impact compression, degradation of type II collagen was also detected; however, this degradation was concentrated around the impact site and extended through the full depth of the tissue. In that study,14 collagen content was evaluated at the earliest 84 days after injury. It is possible that had the in vitro model used in the present study been maintained longer than 28 days, we may have observed significant regional changes in type II collagen content.
Chondrocytes play the important role of producing and maintaining the ECM, including matrix repair after injury. Chondrocyte death independent of any other insult to the cartilage will eventually result in matrix loss.54 Single-impact trauma has been repeatedly shown to promote chondrocyte death in both in vivo31 and in vitro.23,25 Reductions in bovine chondrocyte viability are predominately in the superficial region of the cartilage, and as impact stress increases, cell viability decreases.25 In the present study, cell death was significantly higher in the superficial cartilage region than in other regions and in explants compressed to ≥ 60% strain than in those subjected to less compression. Consistent with these data, compression to ≥ 60% strain induced distinct regions of cell loss.
Chondrocyte cluster formation is a major characteristic of osteoarthritis.55,56 Clusters express markers of hypertrophy such as type X collagen,57 alkaline phosphatase,58 and osteocalcin59 as well as the matrix molecule fibronectin60; however, the role of chondrocyte clusters in the progression of osteoarthritis remains largely unknown. With the model used in the present study, chondrocyte cluster formation was significantly affected by degree of strain and region. Cluster formation was significantly higher in the deep zone of all compressed specimens, compared with in the same region in control specimens, and in the middle zone of specimens compressed to ≥ 60% strain. However, only specimens compressed to 60% strain had significant increases in cluster formation in the superficial zone, compared with formation in control specimens. It is possible that compression to 70% or 80% strain generated localized stresses in the superficial region that prevented chondrocyte cluster formation or it could simply be that these high strains caused cell death, thereby preventing cluster formation. Interestingly, significant increases in chondrocyte cluster formation were observed in injured cartilage, even when there was little or no change in ECM composition. That finding suggested that although chondrocyte clusters are typically found in diseased or damage cartilage at a time when there is a loss of ECM macromolecules, cluster formation may form in response to a factor that is independent of ECM degradation. The ability to reproducibly promote cluster formation with the in vitro model used in our study will allow for highly controlled investigations into the role these clusters play in the progression of osteoarthritis.
The objective of the present study was to develop an in vitro model of cartilage injury that made use of full-thickness equine cartilage and computer-controlled uniaxial unconfined compression. Certain limitations to this type of work should be taken into consideration but are difficult to avoid. First, the data were generated from fresh cadaveric tissue sources and, consequently, biological variation could be expected in the material properties of the cartilage and the response of the cartilage to injury between horses and within the joint of each horses. To examine interanimal variation, we included tissue from 6 horses and used a mixed-model approach to interpret the results to control for horse as a random variable.
To minimize the effects of variability in cartilage material properties by location within each horse, specimen collection was limited to a proximal region of the trochlear ridges that represents approximately 15% of the articular cartilage surface area of the joint. This small region was specifically chosen because the entire region is in contact with the patella during the usual loading process and because the thickness of the cartilage in this region is reportedly rather constant.61 To further control for interhorse variation, all specimens were pooled and 3 specimens were randomly assigned to each treatment group. Indeed, the thickness of the cartilage used did not differ significantly among groups. Specimen selection from this small region prevented the quantitative evaluation of ECM molecules by biochemical analysis, which would have made our model more complete and reliable.
Like all in vitro experiments, the model developed in the present study lacked certain in vivo conditions, including the presence of additional tissue types, cytokines, and biomechanical loading. We believe our experiments are the first step toward developing an in vitro model for further evaluation in more defined conditions, including quantitative biochemical analysis of the response of cartilage to injury.
ABBREVIATIONS
| ECM | Extracellular matrix |
| GAG | Glycosaminoglycan |
| IHC | Immunohistochemistry |
| SOFG | Safranin O fast green |
Smith and Nephew, Andover, Mass.
Mini Bionix II MTS, MTS, Eden Prairie, Minn.
OCT, Sakura, Dublin, Ohio.
Rincon, Optronics, Goleta, Calif.
CryoJane Tape-Transfer system, Instrumedics Inc, St Louis, Mo.
Novus Biologics, Littleton, Colo.
Hybridoma Bank, Iowa City, Iowa.
Jackson Immunoresearch, West Grove, Pa.
PROC GLIMMIX, SAS, version 9.2, SAS Institute Inc, Cary, NC.
References
- 1.↑
Hunziker EB. Articular cartilage repair: basic science and clinical progress. A review of the current status and prospects. Osteoarthritis Cartilage 2002; 10: 432–463.
- 2.↑
McIlwraith CW. General pathobiology of the joint and response to injury. In: Mcilwraith CW, Trotter, GW, eds. Joint disease in the horse. Philadelphia: WB Saunders Co, 1996; 40–70.
- 3.↑
Rossdale PD, Hopes R, Digby NJ, et al. Epidemiological study of wastage among racehorses 1982 and 1983. Vet Rec 1985; 116: 66–69.
- 4.↑
Johnston SA. Osteoarthritis. Joint anatomy, physiology, and pathobiology. Vet Clin North Am Small Anim Pract 1997; 27: 699–723.
- 5.
McIlwraith CW, Vachon A. Review of pathogenesis and treatment of degenerative joint disease. Equine Vet J Suppl 1988;(6):3–11.
- 6.
Scott CC, Athanasiou KA. Mechanical impact and articular cartilage. Crit Rev Biomed Eng 2006; 34: 347–378.
- 7.
Chai DH, Stevens AL, Gordzinsky AJ. Biomechanical aspects: joint injury and osteoarthritis. In: Bronner F, Farach-Carson MC, eds. Bone and osteoarthritis. London: Springer, 2008; 165–179.
- 8.↑
Oeppen RS, Connolly SA, Bencardino JT, et al. Acute injury of the articular cartilage and subchondral bone: a common but unrecognized lesion in the immature knee. AJR Am J Roentgenol 2004; 182: 111–117.
- 9.
Radin EL, Ehrlich MG, Chernack R, et al. Effect of repetitive impulsive loading on the knee joints of rabbits. Clin Orthop Relat Res 1978; 9: 288–293.
- 10.
Newberry WN, Garcia JJ, Mackenzie CD, et al. Analysis of acute mechanical insult in an animal model of post-traumatic osteoarthrosis. J Biomech Eng 1998; 120: 704–709.
- 11.
Thompson RC Jr, Oegema TR Jr, Lewis JL, et al. Osteoarthrotic changes after acute transarticular load. An animal model. J Bone Joint Surg Am 1991; 73: 990–1001.
- 12.
Pickvance EA, Oegema TR Jr, Thompson RC Jr. Immunolocalization of selected cytokines and proteases in canine articular cartilage after transarticular loading. J Orthop Res 1993; 11: 313–323.
- 13.
Milentijevic D, Rubel IF, Liew AS, et al. An in vivo rabbit model for cartilage trauma: a preliminary study of the influence of impact stress magnitude on chondrocyte death and matrix damage. J Orthop Trauma 2005; 19: 466–473.
- 14.↑
Bolam CJ, Hurtig MB, Cruz A, et al. Characterization of experimentally induced post-traumatic osteoarthritis in the medial femorotibial joint of horses. Am J Vet Res 2006; 67: 433–447.
- 15.
Kurz B, Jin M, Patwari P, et al. Biosynthetic response and mechanical properties of articular cartilage after injurious compression. J Orthop Res 2001; 19: 1140–1146.
- 16.
D'Lima DD, Hashimoto S, Chen PC, et al. Human chondrocyte apoptosis in response to mechanical injury. Osteoarthritis Cartilage 2001; 9: 712–719.
- 17.
Lee JH, Fitzgerald JB, Dimicco MA, et al. Mechanical injury of cartilage explants causes specific time-dependent changes in chondrocyte gene expression. Arthritis Rheum 2005; 52: 2386–2395.
- 18.
Kisiday JD, Vanderploeg EJ, McIlwraith CW, et al. Mechanical injury of explants from the articulating surface of the inner meniscus. Arch Biochem Biophys 2010; 494: 138–144.
- 19.↑
McIlwraith CW, Frisbie DD, Kawcak CE, et al. The OARSI histopathology initiative—recommendations for histological assessments of osteoarthritis in the horse. Osteoarthritis Cartilage 2010; 18(suppl 3):S93–S105.
- 20.↑
Roach HI, Aigner T, Kouri JB. Chondroptosis: a variant of apoptotic cell death in chondrocytes? Apoptosis 2004; 9: 265–277.
- 21.
Radin EL, Martin RB, Burr DB, et al. Effects of mechanical loading on the tissues of the rabbit knee. J Orthop Res 1984; 2: 221–234.
- 22.
Dekel S, Weissman SL. Joint changes after overuse and peak overloading of rabbit knees in vivo. Acta Orthop Scand 1978; 49: 519–528.
- 23.
Jeffrey JE, Gregory DW, Aspden RM. Matrix damage and chondrocyte viability following a single impact load on articular cartilage. Arch Biochem Biophys 1995; 322: 87–96.
- 24.
Huser CA, Davies ME. Validation of an in vitro single-impact load model of the initiation of osteoarthritis-like changes in articular cartilage. J Orthop Res 2006; 24: 725–732.
- 25.↑
Quinn TM, Allen RG, Schalet BJ, et al. Matrix and cell injury due to sub-impact loading of adult bovine articular cartilage explants: effects of strain rate and peak stress. J Orthop Res 2001; 19: 242–249.
- 26.
Natoli RM, Scott CC, Athanasiou KA. Temporal effects of impact on articular cartilage cell death, gene expression, matrix biochemistry, and biomechanics. Ann Biomed Eng 2008; 36: 780–792.
- 27.
Chen CT, Burton-Wurster N, Borden C, et al. Chondrocyte necrosis and apoptosis in impact damaged articular cartilage. J Orthop Res 2001; 19: 703–711.
- 28.
Lotz M, Hashimoto S, Kuhn K. Mechanisms of chondrocyte apoptosis. Osteoarthritis Cartilage 1999; 7: 389–391.
- 29.
Tew SR, Kwan AP, Hann A, et al. The reactions of articular cartilage to experimental wounding: role of apoptosis. Arthritis Rheum 2000; 43: 215–225.
- 30.
Repo RU, Finlay JB. Survival of articular cartilage after controlled impact. J Bone Joint Surg Am 1977; 59: 1068–1076.
- 31.↑
Borrelli J Jr, Tinsley K, Ricci WM, et al. Induction of chondrocyte apoptosis following impact load. J Orthop Trauma 2003; 17: 635–641.
- 32.
Quinn TM, Grodzinsky AJ, Hunziker EB, et al. Effects of injurious compression on matrix turnover around individual cells in calf articular cartilage explants. J Orthop Res 1998; 16: 490–499.
- 33.
Lewis JL, Deloria LB, Oyen-Tiesma M, et al. Cell death after cartilage impact occurs around matrix cracks. J Orthop Res 2003; 21: 881–887.
- 34.↑
Borrelli J Jr, Silva MJ, Zaegel MA, et al. Single high-energy impact load causes posttraumatic OA in young rabbits via a decrease in cellular metabolism. J Orthop Res 2009; 27: 347–352.
- 35.
Setton LA, Elliott DM, Mow VC. Altered mechanics of cartilage with osteoarthritis: human osteoarthritis and an experimental model of joint degeneration. Osteoarthritis Cartilage 1999; 7: 2–14.
- 36.
Guilak F, Ratcliffe A, Lane N, et al. Mechanical and biochemical changes in the superficial zone of articular cartilage in canine experimental osteoarthritis. J Orthop Res 1994; 12: 474–484.
- 37.↑
Clark JM. Variation of collagen fiber alignment in a joint surface: a scanning electron microscope study of the tibial plateau in dog, rabbit, and man. J Orthop Res 1991; 9: 246–257.
- 38.
Bullough P, Goodfellow J. The significance of the fine structure of articular cartilage. J Bone Joint Surg Br 1968; 50: 852–857.
- 39.
Mow VC, Lai WM, Eisenfeld J, et al. Some surface characteristics of articular cartilage. II. On the stability of articular surface and a possible biomechanical factor in etiology of chondrode-generation. J Biomech 1974; 7: 457–468.
- 40.
Aspden RM, Hukins DW. Collagen organization in articular cartilage, determined by x-ray diffraction, and its relationship to tissue function. Proc R Soc Lond B Biol Sci 1981; 212: 299–304.
- 41.
Redler I, Mow VC, Zimny ML, et al. The ultrastructure and biomechanical significance of the tidemark of articular cartilage. Clin Orthop Relat Res 1975; 112: 357–362.
- 42.
Muir H, Bullough P, Maroudas A. The distribution of collagen in human articular cartilage with some of its physiological implications. J Bone Joint Surg Br 1970; 52: 554–563.
- 43.
Clarke IC. Articular cartilage: a review and scanning electron microscope study. 1. The interterritorial fibrillar architecture. J Bone Joint Surg Br 1971; 53: 732–750.
- 44.↑
Mizrahi J, Maroudas A, Lanir Y, et al. The “instantaneous” deformation of cartilage: effects of collagen fiber orientation and osmotic stress. Biorheology 1986; 23: 311–330.
- 45.
Kobayashi S, Yonekubo S, Kurogouchi Y. Cryoscanning electron microscopy of loaded articular cartilage with special reference to the surface amorphous layer. J Anat 1996; 188: 311–322.
- 46.
Notzli H, Clark J. Deformation of loaded articular cartilage prepared for scanning electron microscopy with rapid freezing and freeze-substitution fixation. J Orthop Res 1997; 15: 76–86.
- 47.
Glaser C, Putz R. Functional anatomy of articular cartilage under compressive loading: quantitative aspects of global, local and zonal reactions of the collagenous network with respect to the surface integrity. Osteoarthritis Cartilage 2002; 10: 83–99.
- 48.↑
Kaab MJ, Richards RG, Ito K, et al. Deformation of chondrocytes in articular cartilage under compressive load: a morphological study. Cells Tissues Organs 2003; 175: 133–139.
- 49.↑
Chen AC, Bae WC, Schinagl RM, et al. Depth- and strain-dependent mechanical and electromechanical properties of full-thickness bovine articular cartilage in confined compression. J Biomech 2001; 34: 1–12.
- 50.
Jeffrey JE, Thomson LA, Aspden RM. Matrix loss and synthesis following a single impact load on articular cartilage in vitro. Biochim Biophys Acta 1997; 1334: 223–232.
- 51.
Glasson SS, Askew R, Sheppard B, et al. Deletion of active ADAMTS5 prevents cartilage degradation in a murine model of osteoarthritis. Nature 2005; 434: 644–648.
- 52.
Stanton H, Rogerson FM, East CJ, et al. ADAMTS5 is the major aggrecanase in mouse cartilage in vivo and in vitro. Nature 2005; 434: 648–652.
- 53.↑
Patwari P, Cheng DM, Cole AA, et al. Analysis of the relationship between peak stress and proteoglycan loss following injurious compression of human post-mortem knee and ankle cartilage. Biomech Model Mechanobiol 2007; 6: 83–89.
- 54.↑
Simon WH, Richardson S, Herman W, et al. Long-term effects of chondrocyte death on rabbit articular cartilage in vivo. J Bone Joint Surg Am 1976; 58: 517–526.
- 55.
Mankin HJ, Dorfman H, Lippiello L, et al. Biochemical and metabolic abnormalities in articular cartilage from osteo-arthritic human hips. II. Correlation of morphology with biochemical and metabolic data. J Bone Joint Surg Am 1971; 53: 523–537.
- 56.
Poole CA. Articular cartilage chondrons: form, function and failure. J Anat 1997; 191: 1–13.
- 57.↑
von der Mark K, Kirsch T, Nerlich A, et al. Type X collagen synthesis in human osteoarthritic cartilage. Indication of chondrocyte hypertrophy. Arthritis Rheum 1992; 35: 806–811.
- 58.↑
Rees JA, Ali SY. Ultrastructural localisation of alkaline phosphatase activity in osteoarthritic human articular cartilage. Ann Rheum Dis 1988; 47: 747–753.
- 59.↑
Pullig O, Weseloh G, Ronneberger D, et al. Chondrocyte differentiation in human osteoarthritis: expression of osteocalcin in normal and osteoarthritic cartilage and bone. Calcif Tissue Int 2000; 67: 230–240.
- 60.↑
Rees JA, Ali SY, Brown RA. Ultrastructural localisation of fibronectin in human osteoarthritic articular cartilage. Ann Rheum Dis 1987; 46: 816–822.
- 61.↑
Frisbie DD, Cross MW, McIlwraith CW. A comparative study of articular cartilage thickness in the stifle of animal species used in human pre-clinical studies, compared with articular cartilage thickness in the human knee. Vet Comp Orthop Traumatol 2006; 19: 142–146.
