• We are sorry, but NCBI web applications do not support your browser and may not function properly. More information
Logo of nihpaAbout Author manuscriptsSubmit a manuscriptNIH Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
Arthritis Rheum. Author manuscript; available in PMC Mar 1, 2010.
Published in final edited form as:
PMCID: PMC2724839
NIHMSID: NIHMS87471

DEVELOPMENTAL AND OSTEOARTHRITIC CHANGES IN Col6a1 KNOCKOUT MICE: THE BIOMECHANICS OF COLLAGEN VI IN THE CARTILAGE PERICELLULAR MATRIX

Abstract

Chondrocytes are the sole cell type in articular cartilage and maintain the extracellular matrix through a homeostatic balance of anabolic and catabolic activities that are influenced by genetic factors, soluble mediators, and biophysical factors such as mechanical stress. Chondrocytes are encapsulated by a narrow tissue region termed the “pericellular matrix”, which in normal cartilage is defined by the exclusive presence of type VI collagen. Because the pericellular matrix completely surrounds each cell, it is hypothesized to serve as a filter or transducer for biochemical and/or biomechanical signals from the cartilage extracellular matrix. In this study, we used Col6a1−/− mice to investigate whether the lack of collagen VI may affect the development and biomechanical function of the pericellular matrix and alter the mechanical environment of the chondrocytes during joint loading. Col6a1−/− and Col6a1+/− mice possessed structurally intact pericellular matrices, but with significantly reduced mechanical properties as compared to wild-type controls. With age, Col6a1−/− showed accelerated development of osteoarthritic joint degeneration, as well as other musculoskeletal abnormalities such as delayed secondary ossification process and reduced bone mineral density. These findings suggest an important role for type VI collagen in regulating the physiology of the synovial joint, and provide indirect evidence that alterations in the mechanical environment of the chondrocytes, either due to loss of pericellular matrix properties or Col6α1−/− derived joint laxity, can lead to the progression of osteoarthritis.

Keywords: arthritis, collagen VI, proteoglycan, pericellular matrix, chondron, chondrocyte

Introduction

Articular cartilage is the tissue that lines the surfaces of diarthrodial joints and serves as the resilient, low-friction, load-bearing material for joint motion. A sparse population of cells – chondrocytes - maintains the extracellular matrix (ECM) of this tissue through a balance of anabolic and catabolic activities. The micromechanical environment of chondrocytes, in conjunction with biochemical (e.g., growth factors, cytokines) and genetic factors, plays an important role in cartilage homeostasis and, as a consequence, the health of the joint (13). Chondrocytes in articular cartilage are enclosed by a narrow region of tissue, the pericellular matrix (PCM), which together with the enclosed chondrocyte has been termed the chondron (47). The PCM is primarily characterized by the exclusive presence of type VI collagen in normal cartilage, but it also possesses a high concentration of proteoglycans, fibronectin, and types II and IX collagen (4, 8).

The functional role of the PCM in articular cartilage is still unknown, although the fact that it completely surrounds the cell suggests that it regulates the biomechanical, biophysical, and biochemical signals that the chondrocyte perceives (9). For example, interactions between cell surface receptors and the ECM significantly influence matrix metabolism, gene expression, and response to growth factors (1012). Furthermore, cytokines and growth factors that interact with the chondrocyte surface traverse the pericellular environment, where they may be retained and modified (13, 14). From a biomechanical standpoint, there has been considerable speculation that the PCM plays a critical biomechanical role in either in protecting the cells or serving as a “filter” or transducer of physical signals in the ECM (6, 9, 1517), potentially through an interaction of type VI collagen with integrins or other cell surface receptors (1821). Indirect evidence in support of these hypotheses is provided by experimental data showing that a newly formed PCM augments cellular metabolic response to biomechanical loading (22).

Type VI collagen serves as the defining boundary of the PCM in articular cartilage, but it is also found in the ECM of many connective tissues (23). It has a characteristic beaded filamentous structure of tetrameric units that consists of three different α-chains, α1(VI), α2(VI), and α3(VI). Collagen VI has high affinity with numerous ECM components (i.e., biglycan, decorin, hyaluronan, fibronectin, perlecan, and heparin) as well as with the cell membrane (2428). Thus, it has been hypothesized that collagen VI plays important roles in mediating cell-matrix interactions as well as intermolecular interactions in various tissues as well as cell cultures (2933). In articular cartilage, collagen type VI forms a network that anchors the chondrocyte to the PCM (3437) through its interaction with hyaluronan (28, 38), decorin (25), and fibronectin (39).

The goal of this study was to examine the hypothesis that lack of type VI collagen alters the biomechanical properties of the PCM and ECM of articular cartilage. Collagen VI deficient mice were generated by targeted gene disruption of the Col6a1 gene (40). Histological analysis and dual energy x-ray absorptiometry (DXA) were used to examine differences in skeletal development, bone mineral density, and progression of osteoarthritic joint degeneration in wild-type and collagen VI deficient mice. In addition, micromechanical testing was performed on the articular cartilage and on isolated chondrons using microindentation and micropipette aspiration techniques, respectively to determine the role of type VI collagen on the elastic properties of the ECM and PCM of articular cartilage.

Materials and Methods

Collagen VI knockout mice

Collagen VI knockout mice were generated on a CD1 genetic background by targeted gene disruption of the Col6a1 gene, which is responsible for the production of α1(VI) chain (40). The elimination of the α1(VI) chain resulted in the absence of triple helical collagen VI molecules in the ECM (40). Mice were sacrificed at 1, 3, 6, and 11 months time points. All procedures had been approved by the Duke University Institutional Animal Care and Use Committee.

Fluorescence immunohistochemistry

Collagen VI immunostaining was performed using a polyclonal anti-collagen VI antibody raised against a peptide mapping near the amino terminus of murine α1(VI) chain (Santa Cruz Biotechnology, CA). Cryostat sections of the coronal plane were obtained from decalcified femoral heads of mice using standard histological methods.

Skeletal Staining

One-month-old mice were sacrificed, skinned, and eviscerated. Alcian blue and alizarin red were used to stain the cartilage and bone respectively using standard skeletal staining techniques.

Histological analysis and morphometric grading

Fixed cryostat sections were stained with Toluidine blue and Hematoxylin and Eosin (H&E) using standard histological techniques. Osteoarthritic and developmental changes were assessed using quantitative histomorphometric grading schemes on the femoral head of 60 mice at ages of 1, 3, 6, and 11 months (41, 42). Osteoarthritic grading was based on the sum score of surface fibrillation (0 to 4), toluidine blue intensity staining (0 to 3), and fibrocartilage presence (0 to 2), which was reported as percentage of the 0 to 9 scale where 0% corresponds to no histological signs of osteoarthritis and 100% corresponds to the most severe changes (grade 9). Specimens were classified as non-OA (score 0–1), mild OA (15), and severe OA (5 to 9).

For developmental changes, the histological grading scale was based on the presence of the growth plate and the extension of the secondary ossification center (42), which was graded from 0 to 5 (0 corresponded to normal 1-month-old mice and 5 corresponded to normal 11-month-old mice). The grading scale was based on the assessment of the percentage of the ossified area relative to the total area: cartilage with growth plate and no secondary ossification area present (grade 0), area of secondary ossification less than 10% (grade 1), area of secondary ossification more than 10% and less than 50% (grade 2), area of secondary ossification more than 50% and less than 90% (grade 3), area of secondary ossification more than 90% (grade 4), no presence of growth plate (grade 5). The secondary ossification was reported as a percentage of the 0 to 5 scale where 0% corresponds to grade 0 (no secondary ossification) and 100% corresponds to the fully developed femoral head (grade 5).

Bone Mineral Density

Dual energy X-ray absorptiometry (DXA, PIXImus, Lunar Corp, Madison, WI) was used for measuring bone mineral density (43). The mice were weighed and placed in supine position in the DXA unit and the whole body except the skull was measured. A total of 82 mice were analyzed at ages of 1, 3, 6, and 11 months.

Mechanical testing of articular cartilage

A total of 18 right hip joints from one-month-old mice (Col6a1−/−: n=7, Col6a1+/+: n=7, Col6a1+/−: n=4) were tested in indentation using an electromechanical test system (ELF 3200, EnduraTEC, Minnetonka, MN) instrumented with a low capacity load-cell (250g, Sensotec, Columbus, OH) and extensometer (1mm, Epsilon, Jackson, WY) (44). Plane-ended microindenters were machined from glass fibers (diameter: 110 µm, Thorlabs, Newton, NJ). A dual-angle camera system was used to optically align the indenter tip perpendicular to the cartilage surface (Fig. 8). After applying a tare load of 0.3 grams-force and allowing it to equilibrate, four consecutive indentation displacements (5 µm/step with a ramping speed of 1 µm/sec) were applied to the cartilage surface and allowed to equilibrate for 200 sec per step. The time, reaction force, and displacement data were collected throughout the test at 1 Hz. The equilibrium force vs. displacement curve was obtained from the linear region of the curve. After mechanical tests, the thickness of cartilage from the tissue surface to the calcified cartilage was measured at a site adjacent to the test site using routine histology (5 µm sections labeled with Safranin-O and fast green). The Young’s modulus of mouse cartilage was calculated using an elastic indentation model (45) with assumed Poisson’s ratio of 0.25 (44).

Mechanical testing of the pericellular matrix

Chondrons were mechanically isolated (n=93 from 26 donors) from the femoral articular cartilage of 1-month-old mice as described previously with a custom-built “microaspirator”, which applies suction pressure to the cartilage surface with a modified syringe (46). The micropipette aspiration technique (4749) was used to measure the mechanical properties of the PCM, as described previously (5052). With this technique, the surface of the PCM is aspirated into a glass micropipette (12 µm diameter) by the application of a series of controlled pressures up to 18 kPa, and the ensuing equilibrated aspiration length is measured using video microscopy (Fig. 8). The Young’s modulus of the PCM was determined using a theoretical model that represents the chondron as an elastic, compressible layer (i.e., PCM) overlying an elastic half-space (i.e., chondrocyte) (46).

Statistical analysis

Statistical analysis was performed using a multi factorial Analysis of Variance (ANOVA) setup in Statistica (StatSoft, Tulsa, OK, USA). Categorical predictors were considered only AGE (1, 3, 6, 9, and 11) and GENOTYPE (+/+, +/−, and −/−). We assumed those variable were able to predict 4 dependent measurements namely weight, OA score, BMD, and ossification score. Full factorial design revealed that both AGE, GENOTYPE and the AGE*GENOTYPE effects were significant contributors, and thus post hoc comparison was performed using the Fisher LSD method. AGE effects were significant for all 4 measurements as expected. GENOTYPE effects between Col6a1+/+ and Col6a1−/− were significant for all 4 measurements (Figure 3, Figure 4, and Figure 5). GENOTYPE effects between Col6a1+/+, Col6a1+/−, and Col6a1−/− were significant only for OA and ossification scores. AGE*GENOTYPE effects within the same age groups were not significant for OA. For ossification scored, significant difference was observed between Col6a1+/+ and Col6a1−/− at 3 months only. For BMD, a significant difference was observed between Col6a1+/+ and Col6a1−/− for all ages except for 1 month.

Figure 3
Lack of collagen VI results in delayed growth and ossification. (A–C) Toluidine blue staining of the femoral head of 3-month-old mice. (A) In the wild type mice, the secondary ossification process is almost complete by 3 months, while (B) Col6a1 ...
Figure 4
Lack of collagen VI results in age-related osteoarthritis in the hip. (A–C) Histological sections of the femoral cartilage of 11-month-old mice stained with hematoxylin-eosin revealed significant progression of osteoarthritis in the knockout mice ...
Figure 5
Bone mineral density of Col6a1+/+, Col6a1+/−, and Col6a1−/− mice, as measured by micro-DXA. Bone mineral density depended on age (p<0.001) and was significantly lower in mice that lack type VI collagen (p<0.001). ...

Results

Histological Evaluation

Mice lacking collagen VI exhibited no apparent abnormalities and all animals survived until sacrifice at 11 months. Wild type and heterozygous mice showed extensive pericellular labeling for type VI collagen in the articular cartilage, whereas knockout mice revealed no presence of type VI collagen (Fig. 1). Intense labeling of type VI collagen was also observed in the growth plate of one-month-old wild type mice (Fig. 1a).

Figure 1
Immunostaining revealed the pericellular distribution of type VI collagen in the cartilage of one-month-old Col6a1+/+ and Col6a1+/− mice (panel A and B). Collagen VI was also abundant in the ossification area (A). No type VI collagen was present ...

Skeletal staining of bone and cartilage (Fig. 2) in one-month-old mice indicated that Col6a1−/− mice are smaller in size and exhibit a slower ossification process of the upper (Fig. 2 C,D) and lower (Fig. 2 E,F) extremities that the wild type counterparts. The smaller size was also consistent with a trend toward lower body weight of one month old Col6a1−/− mice (16.5±2.4 g) as compared to Col6a1+/+ mice (versus the 18.3±1.99 g) (p=0.11, two-tailed t-test).

Figure 2
Skeletal analysis of one-month-old Col6a1+/+ (A) and Col6a1−/− (B) mice using alcian blue (cartilage) and alizarin red (bone) staining. Homozygous mutants are smaller, with slower ossification progress in the upper (C–D) and lower ...

To better evaluate the developmental process, we measured the secondary ossification process of the femoral head (Fig. 3A–C). Col6a1−/− mice showed significantly delayed ossification at 3 months (grade 2.2±1.8) as compared to Col6a1+/+ mice (grade 4.1±0.2) (p<0.02) (Fig. 3D) (see also Materials and Methods for grading scale and statistical analysis). For the 3 months old mice, only one out of 6 Col6a1−/− mice showed ossification grade above 4, whereas 3 out of 3 Col6a1+/+ showed an ossification grade above or equal to 4. The ossification process was complete for all mice after the 6th month.

Semi-quantitative histological analysis of cartilage degeneration revealed significant age-dependent osteoarthritic changes in the Col6a1−/− mice. Osteoarthritic changes depended on age (p<0.001) and genotype (p<0.05) (Fig. 4). For the 6- to 11-month-old mice, only 2/12 of the Col6a1+/+, 3/7 of the Col6a1+/−, and 11/16 Col6a1−/− scored above 1 and are characterized with OA (either mild or severe, see Materials and Methods for grading scale). However, 0/12 of the Col6a1+/+, 0/7 of the Col6a1+/−, and just 2/16 Col6a1−/− scored above 5 and were characterized as exhibiting severe OA.

Bone Mineral Density

DXA revealed that wild type mice have significantly higher bone mineral density than the knockout counterparts at 3 and 6 months (p<0.001), although these differences were no longer present by 11 months (Fig. 5)

Mechanical properties of articular cartilage and PCM

The PCM exhibited linear elastic behavior, and the Young’s modulus of the PCM of chondrons isolated from Col6a1+/+ mice was significantly higher than those of heterozygous Col6a1+/− mice, which was further reduced in the knockout Col6a1−/− mice (Fig. 6C). Microindentation tests revealed no significant differences in the mechanical properties of the femoral head articular cartilage Col6a1+/+, Col6a1+/−, or Col6a1−/− mice (Fig. 6D).

Figure 6
Mechanical testing of articular cartilage and PCM. (A) A microindentation system comprised by a plane-ended glass indenter was employed to assess the mechanical properties of murine articular cartilage. (B) The micropipette aspiration technique was used ...

Discussion

The findings of this study provide new evidence of significant musculoskeletal changes in Col6a1−/− mice. Primarily, our findings show that mice lacking collagen VI exhibit accelerated development of hip osteoarthritis, as well as a delayed secondary ossification process and lower bone mineral density. Lack of type VI collagen resulted in a loss of the stiffness (decreased modulus) of the PCM of the articular cartilage prior to any detectable histological changes. However, no differences in ECM properties were observed. These findings provide indirect evidence of a role for type VI collagen in regulating the physiology of the chondrocyte, potentially due to alterations in the biological and mechanical environment of the chondrocytes in articular cartilage due to changes in biomechanical properties of the PCM or due to increased joint laxity associated with a deficiency in type VI collagen.

The mechanical environment of the chondrocytes is one of several environmental factors that influence the normal balance between the synthesis and breakdown of articular cartilage and is an important factor in etiopathogenesis of osteoarthritis (13, 53, 54). Thus, changes in the mechanical interactions between the cell and ECM may have a significant influence on the regulatory response of the chondrocyte. While a biomechanical function for the PCM has long been hypothesized (5, 6, 9), there is growing evidence from both theoretical modeling and experimental studies that the PCM plays a significant role in regulating the biomechanical signals perceived by the chondrocyte (17, 55, 56). In normal cartilage, the mechanical properties of the PCM are relatively uniform with depth (57) but are significantly altered with osteoarthritis, exhibiting reduced stiffness and increased fluid permeability (46, 58). The PCM appears to function by providing a relatively uniform cellular microenvironment despite large inhomogeneities in local tissue strain (17, 59). Thus, a compromised PCM could significantly affect the mechanical environment of the chondrocytes in articular cartilage, leading to increased strain at the cellular level (56), which may affect catabolic responses at the level of single cells (60). In other tissues such as bone, however, the pericellular region (i.e., the glycocalyx) can serve as a strain amplifier by coupling fluid drag forces to the actin cytoskeleton within the processes of osteocytes (6163). In the present study, Col6a1−/− mice showed significantly reduced PCM stiffness at 1 month of age, preceding any histological or biomechanical changes in the overall articular cartilage. With age, these mice exhibited accelerated development of osteoarthritis. These findings provide indirect evidence that early alterations in the mechanical properties of the PCM are associated to the progression of osteoarthritis.

In normal articular cartilage, type VI collagen is exclusively present in the PCM and it has been characterized as a discrete marker of chondron anatomy (36). For this reason, it has been hypothesized that type VI collagen is necessary for providing the structural integrity and mechanical properties of the PCM. Contrary to our hypotheses, though, Col6a1−/− mice exhibited intact chondrons that could be isolated despite the lack of type VI collagen. This finding suggests that proteins other than collagen VI provide some of the structural integrity of cartilage PCM. Nonetheless, the Young’s modulus (stiffness) of the PCM of Col6a1−/− mice was dramatically decreased to nearly one-third of the wild-type controls indicating the important role of type VI collagen in the properties of the PCM.

An important issue that must be considered is the link between collagen VI deficiency and changes in muscle physiology displayed by Col6a1−/− mice (40). Such link has also been observed in humans, where mutations of collagen VI genes have been shown to play a causal role in two inherited disorders of muscle, Bethlem myopathy and Ullrich congenital muscular dystrophy (UCMD) (64, 65). It is possible that some features of UCMD, particularly joint laxity or predisposition to hip dislocation, may also contribute to the accelerated hip degeneration observed in Col6a1−/− mice. Since joint laxity and PCM mechanical alterations are both inheritably coupled in Col6a−/− mice and both lead to altered mechanical environment in chondrocytes, both factors can contribute to the development of OA. While the present study clearly shows an association between Col6a1 deficiency and OA, presumably via mechanical alterations caused by joint laxity or altered PCM properties, further studies aimed at developing and characterizing conditional or tissue-specific knockouts may be required to fully understand the mechanisms by which Col6a1 deficiency leads to OA. Nonetheless, our results are consistent with the hypothesized role of type VI collagen as an integrating molecule in the structure of cells and tissues; downregulation of collagen VI is associated with tissue laxity and wasting (e.g., Bethlem myopathy, UCMD, joint hyperlaxity), whereas collagen VI upregulation results in increased fibrosis and tissue stiffness (e.g., Bullous keratopathy, scleroderma) (40, 6673).

In our experiments we found no gross morphologic differences between wild type and collagen VI knockout chondrons other than reduced skeletal size of Col6a1−/− mice. Skeletal changes were apparent as a retardation of the developmental process until 11 months of age. During development, histogenesis of long bones occurs via endochondral ossification of cartilage tissue. During this process, chondrocytes in the epiphyseal plate differentiate into mature hypertrophic cells and finally are eliminated from the growth plate (74). The hypertrophic cell lacunae are invaded by vessels carrying mesenchymal and osteogenic cells that differentiate into osteoclasts and synthesize a bony matrix. A similar procedure, known as secondary ossification, takes place at the end of the bone where the formation of the bony epiphysis occurs. Our experimental results point to a slowing of secondary ossification changes and decreased bone mineral density in the Col6a1−/− mice. While there is no known direct mechanism coupling type VI collagen deficiency to endochondral ossification, type VI collagen may provide a scaffold for osteoblasts, preosteoblasts and chondrocytes to proceed to osteochondral ossification (75). In addition, type VI collagen has been linked to the early events of chondrocyte differentiation (76), to the regulation of mesenchymal cell proliferation in vitro (77), and to ECM stabilization during development (36). It has also been hypothesized that collagen VI is important for chondrocyte proliferation and hypertrophy in cartilage (36, 78, 79). These studies in conjunction with our observation of the ubiquitous presence of type VI collagen in the growth plate (Fig. 1A) support the hypothesis that collagen VI deficiency may delay cell differentiation and proliferation, resulting in delayed development and decreased bone formation. Interestingly, the COL6A1 gene was recently identified as the locus for ossification of the posterior longitudinal ligament of the spine (75) and has been also associated with increased systemic bone mineral density and diffuse idiopathic skeletal hyperostosis (80). These findings point to a role for collagen VI in diseases associated with high-bone-mass, consistent with the lower bone mineral density we observed in Col6a1−/− mice. While it is beyond the scope of the present study to analyze the mechanisms resulting in altered bone mineral density, these changes may also be biomechanical in origin, as collagen VI deficiency causes muscular dystrophy (40, 68), which can lead to abnormal mechanical loading regime of the musculoskeletal systems.

In this study, the role of an abnormal mechanical environment on chondrocytes was investigated by using collagen type VI knockout mice. Our findings suggest that collagen VI plays a major role in the mechanical properties of the PCM, and thus, the mechanical environment of the chondrocytes. Col6a1−/− mice showed accelerated development of osteoarthritis that may be “biomechanical” in nature, either via altered properties of the PCM or inheritable joint laxity. In addition, our findings provide direct evidence that collagen type VI might have a significant role on the osteochondral ossification process by modulating the chondrocyte and mesenchymal cell differentiation and proliferation activities. This model may provide a valuable tool to better understand how changes in the mechanical environment of the chondrocytes may lead to abnormal skeletal development and development of osteoarthritis.

Acknowledgments

The authors would like to thank Dr. David Birk for his important advice and Gregory Williams and Jason Perera for their assistance with the project. This study was supported by the National Institutes of Health grants AG15768, AR48182, AR48852, and AR50245, and by Telethon grant GGP04113.

References

1. Stockwell RA. Structure and function of the chondrocyte under mechanical stress. In: Helminen HJ, Kiviranta I, Tammi M, Saamanen AM, KP, Jurvelin J, editors. Joint Loading: Biology and Health of Articular Structures. Bristol: Wright and Sons; 1987. pp. 126–148.
2. van Campen GPJ, van de Stadt RJ. Cartilage and chondrocytes responses to mechanical loading in vitro. In: Helminen HJ, Kiviranta I, Tammi M, Saamanen AM, KP, Jurvelin J, editors. Joint Loading: Biology and Health of Articular Structures. Bristol: Wright and Sons; 1987. pp. 112–125.
3. Guilak F, Sah RL, Setton LA. Physical regulation of cartilage metabolism. In: Mow VC, Hayes WC, editors. Basic Orthopaedic Biomechanics. 2nd ed. Philadelphia: Lippincott-Raven; 1997. pp. 179–207.
4. Poole CA. Chondrons, the chondrocyte and its pericellular microenvironment. In: Kuettner KE, Schleyerbach R, Peyron JG, Hascall VC, editors. Articular Cartilage and Osteoarthritis. New York: Raven Press; 1992. pp. 201–220.
5. Poole CA. Articular cartilage chondrons: form, function and failure. Journal of Anatomy. 1997;191(Pt 1):1–13. [PMC free article] [PubMed]
6. Szirmai JA. The concept of the chondron as a biomechanical unit. In: Hartmann F, editor. Biopolymer und Biomechanik von Bindegewebssystemen. Berlin: Academic Press; 1974. p. 87.
7. Benninghoff A. Form und bau der Gelenkknorpel in ihren Beziehungen Zur Funktion. Zweiter Teil: der Aufbau des Gelenkknorpels in sienen Bezienhungen zur Funktion. 1925;2:783.
8. Poole CA, Gilbert RT, Herbage D, Hartmann DJ. Immunolocalization of type IX collagen in normal and spontaneously osteoarthritic canine tibial cartilage and isolated chondrons. Osteoarthritis and Cartilage. 1997;5(3):191–204. [PubMed]
9. Guilak F, Alexopoulos LG, Upton ML, Youn I, Choi JB, Cao L, et al. The pericellular matrix as a transducer of biomechanical and biochemical signals in articular cartilage. Ann N Y Acad Sci. 2006;1068:498–512. [PubMed]
10. Adams JC, Watt FW. Regulation of development and differentiation by the extracellular matrix. Development. 1993;117:1183–1198. [PubMed]
11. Boudreau N, Myers C, Bissel MJ. From laminin to lamin: regulation of tissue-specific gene expression by the ECM. Trends in Cell Biology. 1995;5:1–4. [PubMed]
12. Loeser RF. Growth factor regulation of chondrocyte integrins. Differential effects of insulin-like growth factor 1 and transforming growth factor beta on alpha 1 beta 1 integrin expression and chondrocyte adhesion to type VI collagen. Arthritis Rheum. 1997;40(2):270–276. [PubMed]
13. Ruoslahti E, Yamaguchi Y. Proteoglycans as modulators of growth factor activities. Cell. 1991;64(867–69):867–869. [PubMed]
14. Sandy JD, O'Neill JR, Ratzlaff LC. Acquisition of hyaluronate-binding affinity in vivo by newly synthsized cartilage proteoglycans. Biochemical Journal. 1989;258:875–880. [PMC free article] [PubMed]
15. Poole CA, Flint MH, Beaumont BW. Chondrons extracted from canine tibial cartilage: preliminary report on their isolation and structure. Journal of Orthopaedic Research. 1988;6(3):408–419. [PubMed]
16. Poole CA. Chondrons: the chondrocyte and its pericellular microenvironment. In: Kuettner KE, Schleyerbach R, Peyron JG, Hascall VC, editors. Articular Cartilage and Osteoarthritis. New York: London: Academic Press; 1992. pp. 201–220.
17. Choi JB, Youn I, Cao L, Leddy HA, Gilchrist CL, Setton LA, et al. Zonal changes in the three-dimensional morphology of the chondron under compression: the relationship among cellular, pericellular, and extracellular deformation in articular cartilage. J Biomech. 2007;40(12):2596–2603. [PMC free article] [PubMed]
18. Lee V, Cao L, Zhang Y, Kiani C, Adams ME, Yang BB. The roles of matrix molecules in mediating chondrocyte aggregation, attachment, and spreading. Journal of Cellular Biochemistry. 2000;79(2):322–333. [PubMed]
19. Loeser RF, Sadiev S, Tan L, Goldring MB. Integrin expression by primary and immortalized human chondrocytes: evidence of a differential role for alpha1beta1 and alpha2beta1 integrins in mediating chondrocyte adhesion to types II and VI collagen. Osteoarthritis and Cartilage. 2000;8(2):96–105. [PubMed]
20. Knudson W, Loeser RF. CD44 and integrin matrix receptors participate in cartilage homeostasis. Cellular and Molecular Life Sciences. 2002;59(1):36–44. [PubMed]
21. McDevitt CA, Marcelino J, Tucker L. Interaction of intact type VI collagen with hyaluronan. FEBS Letters. 1991;294(3):167–170. [PubMed]
22. Buschmann MD, Gluzband YA, Grodzinsky AJ, Hunziker EB. Mechanical compression modulates matrix biosynthesis in chondrocyte/agarose culture. Journal of Cell Science. 1995;108(Pt 4):1497–1508. [PubMed]
23. Timpl R, Engel J. Type VI collagen. In: Mayne R, Burgeson RE, editors. Structure and Function of Collagen Types. New York: London: Academic Press; 1987. pp. 105–143.
24. Wiberg C, Hedbom E, Khairullina A, Lamande SR, Oldberg A, Timpl R, et al. Biglycan and decorin bind close to the n-terminal region of the collagen VI triple helix. J Biol Chem. 2001;276(22):18947–18952. [PubMed]
25. Bidanset DJ, Guidry C, Rosenberg LC, Choi HU, Timpl R, Hook M. Binding of the proteoglycan decorin to collagen type VI. J Biol Chem. 1992;267(8):5250–5256. [PubMed]
26. Tillet E, Wiedemann H, Golbik R, Pan TC, Zhang RZ, Mann K, et al. Recombinant expression and structural and binding properties of alpha 1(VI) and alpha 2(VI) chains of human collagen type VI. Eur J Biochem. 1994;221(1):177–185. [PubMed]
27. Specks U, Mayer U, Nischt R, Spissinger T, Mann K, Timpl R, et al. Structure of recombinant N-terminal globule of type VI collagen alpha 3 chain and its binding to heparin and hyaluronan. Embo J. 1992;11(12):4281–4290. [PMC free article] [PubMed]
28. McDevitt CA, Marcelino J, Tucker L. Interaction of intact type VI collagen with hyaluronan. FEBS Lett. 1991;294(3):167–170. [PubMed]
29. Bonaldo P, Russo V, Bucciotti F, Doliana R, Colombatti A. Structural and functional features of the alpha 3 chain indicate a bridging role for chicken collagen VI in connective tissues. Biochemistry. 1990;29(5):1245–1254. [PubMed]
30. Pfaff M, Aumailley M, Specks U, Knolle J, Zerwes HG, Timpl R. Integrin and Arg-Gly-Asp dependence of cell adhesion to the native and unfolded triple helix of collagen type VI. Exp Cell Res. 1993;206(1):167–176. [PubMed]
31. Aumailley M, Mann K, von der Mark H, Timpl R. Cell attachment properties of collagen type VI and Arg-Gly-Asp dependent binding to its alpha 2(VI) and alpha 3(VI) chains. Exp Cell Res. 1989;181(2):463–474. [PubMed]
32. Burg MA, Tillet E, Timpl R, Stallcup WB. Binding of the NG2 proteoglycan to type VI collagen and other extracellular matrix molecules. J Biol Chem. 1996;271(42):26110–26116. [PubMed]
33. Lamande SR, Morgelin M, Adams NE, Selan C, Allen JM. The C5 domain of the collagen VI alpha3(VI) chain is critical for extracellular microfibril formation and is present in the extracellular matrix of cultured cells. J Biol Chem. 2006;281(24):16607–16614. [PubMed]
34. Buckwalter JA, Mankin HJ. Articular cartilage: tissue design and chondrocyte-matrix interactions. Instr Course Lect. 1998;47:477–486. [PubMed]
35. Marcelino J, McDevitt CA. Attachment of articular cartilage chondrocytes to the tissue form of type VI collagen. Biochim Biophys Acta. 1995;1249(2):180–188. [PubMed]
36. Sherwin AF, Carter DH, Poole CA, Hoyland JA, Ayad S. The distribution of type VI collagen in the developing tissues of the bovine femoral head. Histochem J. 1999;31(9):623–632. [PubMed]
37. Keene DR, Engvall E, Glanville RW. Ultrastructure of type VI collagen in human skin and cartilage suggests an anchoring function for this filamentous network. J Cell Biol. 1988;107(5):1995–2006. [PMC free article] [PubMed]
38. Kielty CM, Whittaker SP, Grant ME, Shuttleworth CA. Type VI collagen microfibrils: evidence for a structural association with hyaluronan. J Cell Biol. 1992;118(4):979–990. [PMC free article] [PubMed]
39. Chang J, Nakajima H, Poole CA. Structural colocalisation of type VI collagen and fibronectin in agarose cultured chondrocytes and isolated chondrons extracted from adult canine tibial cartilage. J Anat. 1997;190(Pt 4):523–532. [PMC free article] [PubMed]
40. Bonaldo P, Braghetta P, Zanetti M, Piccolo S, Volpin D, Bressan GM. Collagen VI deficiency induces early onset myopathy in the mouse: an animal model for Bethlem myopathy. Hum Mol Genet. 1998;7(13):2135–2140. [PubMed]
41. Carlson CS, Guilak F, Vail TP, Gardin JF, Kraus VB. Synovial fluid biomarker levels predict articular cartilage damage following complete medial meniscectomy in the canine knee. J Orthop Res. 2002;20(1):92–100. [PubMed]
42. Rivas R, Shapiro F. Structural stages in the development of the long bones and epiphyses: a study in the New Zealand white rabbit. J Bone Joint Surg Am. 2002;84-A(1):85–100. [PubMed]
43. Fink C, Cooper HJ, Huebner JL, Guilak F, Kraus VB. Precision and accuracy of a transportable dual-energy X-ray absorptiometry unit for bone mineral measurements in guinea pigs. Calcif Tissue Int. 2002;70(3):164–169. [PubMed]
44. Cao L, Youn I, Guilak F, Setton LA. Compressive properties of mouse articular cartilage determined in a novel micro-indentation test method and biphasic finite element model. J Biomech Eng. 2006;128(5):766–771. [PubMed]
45. Hayes WC, Keer LM, Herrmann G, Mockros LF. A mathematical analysis for indentation tests of articular cartilage. Journal of Biomechanics. 1972;5:541–551. [PubMed]
46. Alexopoulos LG, Haider MA, Vail TP, Guilak F. Alterations in the mechanical properties of the human chondrocyte pericellular matrix with osteoarthritis. J Biomech Eng. 2003;125(3):323–333. [PubMed]
47. Hochmuth RM. Micropipette aspiration of living cells. Journal of Biomechanics. 2000;33(1):15–22. [PubMed]
48. Trickey WR, Vail TP, Guilak F. The role of the cytoskeleton in the viscoelastic properties of human articular chondrocytes. J Orthop Res. 2004;22(1):131–139. [PubMed]
49. Guilak F, Erickson GR, Ting-Beall HP. The effects of osmotic stress on the viscoelastic and physical properties of articular chondrocytes. Biophysical Journal. 2002;82(2):720–727. [PMC free article] [PubMed]
50. Guilak F, Alexopoulos LG, Haider MA, Ting-Beall HP, Setton LA. Zonal uniformity in mechanical properties of the chondrocyte pericellular matrix: Micropipette aspiration of canine chondrons isolated by cartilage homogenization. Annals of Biomedical Engineering. 2005;33(10):000–000. [PubMed]
51. Alexopoulos LG, Haider MA, Vail TP, Guilak F. Alterations in the mechanical properties of the human chondrocyte pericellular matrix with osteoarthritis. Journal of Biomechanical Engineering. 2003;125(3):323–333. [PubMed]
52. Alexopoulos LG, Williams GM, Upton ML, Setton LA, Guilak F. Osteoarthritic changes in the biphasic mechanical properties of the chondrocyte pericellular matrix in articular cartilage. Journal of Biomechanics. 2005;38(3):509–517. [PubMed]
53. Guilak F, Ratcliffe A, Lane N, Rosenwasser MP, Mow VC. Mechanical and biochemical changes in the superficial zone of articular cartilage in canine experimental osteoarthritis. Journal of Orthopaedic Research. 1994;12(4):474–484. [PubMed]
54. Setton LA, Mow VC, Muller FJ, Pita JC, Howell DS. Mechanical properties of canine articular cartilage are significantly altered following transection of the anterior cruciate ligament. Journal of Orthopaedic Research. 1994;12(4):451–463. [PubMed]
55. Guilak F, Mow VC. The mechanical environment of the chondrocyte: a biphasic finite element model of cell-matrix interactions in articular cartilage. Journal of Biomechanics. 2000;33(12):1663–1673. [PubMed]
56. Alexopoulos LG, Setton LA, Guilak F. The biomechanical role of the chondrocyte pericellular matrix in articular cartilage. Acta Biomater. 2005;1(3):317–325. [PubMed]
57. Guilak F, Alexopoulos LG, Haider MA, Ting-Beall HP, Setton LA. Zonal uniformity in mechanical properties of the chondrocyte pericellular matrix: micropipette aspiration of canine chondrons isolated by cartilage homogenization. Ann Biomed Eng. 2005;33(10):1312–1318. [PubMed]
58. Alexopoulos LG, Williams GM, Upton ML, Setton LA, Guilak F. Osteoarthritic changes in the biphasic mechanical properties of the chondrocyte pericellular matrix in articular cartilage. J Biomech. 2005;38(3):509–517. [PubMed]
59. Youn I, Choi JB, Cao L, Setton LA, Guilak F. Zonal variations in the three-dimensional morphology of the chondron measured in situ using confocal microscopy. Osteoarthritis Cartilage. 2006;14(9):889–897. [PubMed]
60. Leipzig ND, Athanasiou KA. Static compression of single chondrocytes catabolically modifies single-cell gene expression. Biophys J. 2008;94(6):2412–2422. [PMC free article] [PubMed]
61. Han Y, Cowin SC, Schaffler MB, Weinbaum S. Mechanotransduction and strain amplification in osteocyte cell processes. Proc Natl Acad Sci U S A. 2004;101(47):16689–16694. [PMC free article] [PubMed]
62. Weinbaum S, Guo P, You L. A new view of mechanotransduction and strain amplification in cells with microvilli and cell processes. Biorheology. 2001;38(2–3):119–142. [PubMed]
63. You L, Cowin SC, Schaffler MB, Weinbaum S. A model for strain amplification in the actin cytoskeleton of osteocytes due to fluid drag on pericellular matrix. J Biomech. 2001;34(11):1375–1386. [PubMed]
64. Pepe G, Lucarini L, Zhang RZ, Pan TC, Giusti B, Quijano-Roy S, et al. COL6A1 genomic deletions in Bethlem myopathy and Ullrich muscular dystrophy. Ann Neurol. 2006;59(1):190–195. [PubMed]
65. Lampe AK, Bushby KM. Collagen VI related muscle disorders. J Med Genet. 2005;42(9):673–685. [PMC free article] [PubMed]
66. Griffiths MR, Shepherd M, Ferrier R, Schuppan D, James OF, Burt AD. Light microscopic and ultrastructural distribution of type VI collagen in human liver: alterations in chronic biliary disease. Histopathology. 1992;21(4):335–344. [PubMed]
67. Higuchi I, Shiraishi T, Hashiguchi T, Suehara M, Niiyama T, Nakagawa M, et al. Frameshift mutation in the collagen VI gene causes Ullrich's disease. Ann Neurol. 2001;50(2):261–265. [PubMed]
68. Pan TC, Zhang RZ, Sudano DG, Marie SK, Bonnemann CG, Chu ML. New molecular mechanism for Ullrich congenital muscular dystrophy: a heterozygous in-frame deletion in the COL6A1 gene causes a severe phenotype. Am J Hum Genet. 2003;73(2):355–369. [PMC free article] [PubMed]
69. Ljubimov AV, Burgeson RE, Butkowski RJ, Couchman JR, Wu RR, Ninomiya Y, et al. Extracellular matrix alterations in human corneas with bullous keratopathy. Invest Ophthalmol Vis Sci. 1996;37(6):997–1007. [PubMed]
70. Mollnau H, Munkel B, Schaper J. Collagen VI in the extracellular matrix of normal and failing human myocardium. Herz. 1995;20(2):89–94. [PubMed]
71. Rauch A, Pfeiffer RA, Trautmann U. Deletion or triplication of the alpha 3 (VI) collagen gene in three patients with 2q37 chromosome aberrations and symptoms of collagen-related disorders. Clin Genet. 1996;49(6):279–285. [PubMed]
72. Specks U, Nerlich A, Colby TV, Wiest I, Timpl R. Increased expression of type VI collagen in lung fibrosis. Am J Respir Crit Care Med. 1995;151(6):1956–1964. [PubMed]
73. Takasaki S, Fujiwara S, Shinkai H, Ooshima A. Human type VI collagen: purification from human subcutaneous fat tissue and an immunohistochemical study of morphea and systemic sclerosis. J Dermatol. 1995;22(7):480–485. [PubMed]
74. Ballock RT, O'Keefe RJ. Physiology and pathophysiology of the growth plate. Birth Defects Res Part C Embryo Today. 2003;69(2):123–143. [PubMed]
75. Tanaka T, Ikari K, Furushima K, Okada A, Tanaka H, Furukawa K, et al. Genomewide linkage and linkage disequilibrium analyses identify COL6A1, on chromosome 21, as the locus for ossification of the posterior longitudinal ligament of the spine. Am J Hum Genet. 2003;73(4):812–822. [PMC free article] [PubMed]
76. Quarto R, Dozin B, Bonaldo P, Cancedda R, Colombatti A. Type VI collagen expression is upregulated in the early events of chondrocyte differentiation. Development. 1993;117(1):245–251. [PubMed]
77. Atkinson JC, Ruhl M, Becker J, Ackermann R, Schuppan D. Collagen VI regulates normal and transformed mesenchymal cell proliferation in vitro. Exp Cell Res. 1996;228(2):283–291. [PubMed]
78. Mylona P, Kielty CM, Hoyland JA, Aplin JD. Expression of type VI collagen mRNAs in human endometrium during the menstrual cycle and first trimester of pregnancy. J Reprod Fertil. 1995;103(1):159–167. [PubMed]
79. Sloan P, Carter DH, Kielty CM, Shuttleworth CA. An immunohistochemical study examining the role of collagen type VI in the rodent periodontal ligament. Histochem J. 1993;25(7):523–530. [PubMed]
80. Tsukahara S, Miyazawa N, Akagawa H, Forejtova S, Pavelka K, Tanaka T, et al. COL6A1, the candidate gene for ossification of the posterior longitudinal ligament, is associated with diffuse idiopathic skeletal hyperostosis in Japanese. Spine. 2005;30(20):2321–2324. [PubMed]
PubReader format: click here to try

Formats:

Related citations in PubMed

See reviews...See all...

Cited by other articles in PMC

See all...

Links

Recent Activity

Your browsing activity is empty.

Activity recording is turned off.

Turn recording back on

See more...