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The Journals of Gerontology Series A: Biological Sciences and Medical Sciences 56:B172-B179 (2001)
© 2001 The Gerontological Society of America

Telomere Erosion and Senescence in Human Articular Cartilage Chondrocytes

James A. Martina and Joseph A. Buckwaltera

a Iowa City Veterans Administration Medical Center and University of Iowa Department of Orthopaedics, Iowa City

James A. Martin, Department of Orthopaedic Surgery, Biochemistry Laboratory, 1182 ML, The University of Iowa, Iowa City, IA 52242 E-mail: james-martin{at}uiowa.edu.

Decision Editor: John A. Faulkner, PhD


   Abstract
Top
 Abstract
Materials and Methods
 Results
 Discussion
 References
 
Aging and the degeneration of articular cartilage in osteoarthritis are distinct processes, but a strong association exists between age and the incidence and prevalence of osteoarthritis. We hypothesized that this association is due to in vivo replicative senescence, which causes age-related declines in the ability of chondrocytes to maintain articular cartilage. For this hypothesis to be tested, senescence-associated markers were measured in human articular chondrocytes from donors ranging in age from 1 to 87 years. These measures included in situ staining for senescence-associated ß-galactosidase activity, 3H-thymidine incorporation assays for mitotic activity, and Southern blots for telomere length determinations. We found that senescence-associated ß-galactosidase activity increased with age, whereas both mitotic activity and mean telomere length declined. These findings indicate that chondrocyte replicative senescence occurs in vivo and support the hypothesis that the association between osteoarthritis and aging is due in part to replicative senescence. The data also imply that transplantation procedures performed to restore damaged articular surfaces could be limited by the inability of older chondrocytes to form new cartilage after transplantation.

ARTICULAR cartilage stability depends on the biosynthetic activities of chondrocytes, which counteract normal degradation of matrix macromolecules. In most young people, the timely synthesis of appropriate extracellular matrix (ECM) molecules prevents the progressive loss of articular cartilage associated with the clinical syndrome of osteoarthritis; however, the incidence and prevalence of synovial joint degeneration increases dramatically in middle age, suggesting that age-related cartilage changes render the tissue incapable of adequately maintaining the ECM. This phenomenon has been attributed to the harmful effects of mechanical and chemical stress that are thought to impair maintenance activity by killing chondrocytes outright or by inducing apoptosis (1)(2)(3). Although such environmental factors undoubtedly contribute to degenerative disease in some individuals, they do not explain the seemingly irreversible age-dependent decline in chondrocyte growth factor responsiveness and ECM synthesis found by a number of investigators (4)(5)(6)(7)(8)(9)(10)(11)(12)(13)(14)(15). These changes, which persist in cell culture, are more likely a reflection of aging processes intrinsic to chondrocytes and other somatic cells (4)(5)(12)(13)(14).

One recently formulated hypothesis suggests that cell aging is regulated by an intrinsic genetic "clock" associated with changes in DNA structures termed telomeres. Telomeres are DNA sequences at the ends of chromosomes that are necessary for chromosomal replication (16). Mean telomere length can be estimated from preparations of genomic DNA by using terminal restriction fragment analysis. This Southern blot approach uses a telomere sequence-specific probe and takes advantage of the fact that common restriction enzyme cleavage sites, which are frequent in most genomic DNA, are not found within telomere repeats. Chromosomes from young, normal somatic cells show relatively long terminal restriction fragments of >9 kilobase pairs (kbp), but these are eroded at the rate of 100–200 base pairs (bp) with each cell division cycle (17)(18). Erosion beyond the minimum critical length necessary for DNA replication (5–7.6 kbp) results in cell cycle arrest, a condition referred to as replicative senescence (19)(20). Most cell types reach cell cycle arrest after a characteristic number of population doublings. This fundamental barrier to unbridled growth, termed the Hayflick limit, is common to somatic cells that lack an enzyme responsible for replacing telomere sequences (16)(21). The Hayflick limit for human fibroblasts has been estimated at ~60 population doublings (22), whereas the estimated limit for human chondrocytes is ~35 doublings (23). In contrast, germ cell lines and cancer cell lines, in which the "telomerase" enzyme is active, are virtually immortal (18)(24)(25). Furthermore, transfection with the telomerase gene is sufficient to greatly extend the replicative life span of normal somatic cells (26)(27). In telomerase-negative cells, telomere length can be viewed as a cumulative history of preceding cell division as well as a predictor of future capacity to divide (19)(20).

Comparisons of telomeres from young and old donors show a significant correlation between telomere length and donor age for some cell types, including T-cells, dystrophic skeletal muscle cells, kidney cells, and vascular smooth muscle cells, indicating significant cell turnover and the absence of telomerase expression in these tissues (28)(29)(30)(31). In the case of vascular smooth muscle cells, telomere shortening was directly associated with replicative senescence and degenerative disease. Senescent smooth muscle cells accumulate with age and mechanical stress exposure in blood vessel walls, where high stress levels continuously stimulate demand for new cells. Senescent or near-senescent cells from these sites fail to proliferate in culture and bear shortened telomeres compared with those of their counterparts from low stress sites. Finally, abnormal metabolism and gene expression by senescent cells appears to contribute to atherosclerotic plaque development (31)(32). These data confirm that cell turnover-driven telomere erosion occurs in vivo and leads to senescence and degenerative disease.

Declining protein synthesis, altered growth factor and cytokine responses, and longer population doubling times are senescence-like phenotypic changes that begin to appear in continuously grown somatic cell cultures long before Hayflick limits are reached (28)(33)(34)(35)(36). This suggests that cell populations begin to drift toward senescence relatively early in their replicative life spans, before telomeres have eroded to critical lengths. Declines in cartilage ECM synthesis in serially passaged chondrocyte cultures support the idea that early replicative history is important for chondrocytic gene expression. Chondrocyte growth in a monolayer culture generally results in the rapid loss of the chondrogenic phenotype or "dedifferentiation." Up to a point, these changes are reversible by subculturing the monolayer-grown cells in three-dimensional gels. The ability to return to chondrogenic gene expression in gel culture depends on the number of preceding passages in the monolayer: After approximately five monolayer passages, the maximum rate of protein synthesis and cartilage matrix production (Type II collagen synthesis) in gel culture declines by twofold to fourfold, compared to primary cultures or to cultures passaged only one to four times (37). These chondrocyte cultures were capable of growth beyond passage 5, indicating that they were not yet senescent. These results suggest that replication-induced phenotypic drift occurs in chondrocytes before their entry into senescence. These observations also suggest that declining ECM expression associated with chondrocyte turnover could gradually undercut cartilage maintenance activities with aging.

The relevance of telomeres to cartilage aging and disease rests on proof that in vivo chondrocyte turnover rates are sufficient to cause telomere erosion. Short-term DNA labeling studies indicate that chondrocyte mitoses are present but relatively rare in normal cartilage (38)(39)(40). Although this apparent rate of turnover is too slow to result in significant telomere erosion over the short term, decades of turnover might well be sufficient. Furthermore, mitotic activity increases severalfold following cartilage injury, which could significantly accelerate telomere erosion in some individuals (3)(38). Increased mitotic activity during cartilage degeneration may also speed up the accumulation of senescent, growth-arrested chondrocytes in end-stage osteoarthritis (39)(40). These findings suggest that, in many cases, in vivo chondrocyte turnover is sufficient to result in biologically significant telomere erosion.

The role of chondrocyte turnover in cartilage aging and disease has not been systematically studied, partly because of the difficulty of assessing the in vivo replicative history of chondrocytes. Terminal restriction fragment length analysis of telomeres offers a simple means to overcome this problem as cell turnover should be detectable as an age-related decline in average telomere length. If telomere erosion causes senescence, telomere length should correlate with phenotypic measures of senescence. With the use of these rationales, we hypothesized that telomere length in human articular cartilage chondrocytes declines as a function of donor age as phenotypic measures of senescence increase. Southern blot analyses of telomere restriction fragments were used to determine mean terminal restriction fragment lengths in cells isolated from donors aged 1 to 85 years. In addition, we investigated phenotypic changes associated with cell senescence, including declines in DNA synthesis rates, and expression of senescence-associated ß-galactosidase activity.


   Materials and Methods
Top
 Abstract
 Materials and Methods
Results
 Discussion
 References
 
Human articular cartilage samples were harvested, chopped, and digested overnight in Dulbecco's modified Eagle medium (DMEM), containing 10% fetal calf serum (FCS; Gibco-BRL, Rockville, MD) and 0.5 mg/ml of Pronase E (Sigma, St. Louis, MO), and 0.5 mg/ml of Collagenase Type 1A (Sigma). The resulting single cell suspension was filtered through 70-µm mesh nylon cloth, and the cells were counted by using a hemocytometer. Cells were then plated in a monolayer culture and incubated for 1–5 days in DMEM–10% FCS without enzymes. Human chondrosarcoma cells from a tumor excised at the University of Iowa Department of Orthopaedic surgery, and a human fibrosarcoma cell line, HT1080 (ATCC), were thawed from frozen stocks and cultured as monolayers for one to three passages.

Genomic DNA was isolated from ~1 x 106 to 5 x 106 cells by using a DNEasy kit (Qiagen, Valencia, CA), according to the manufacturer's directions. The DNA concentration of each sample was determined by ultraviolet spectrophotometry and 2 µg was digested to completion with 10 units each of Rsa I and Hinf I (New England Biolabs, Beverly, MA) in a 60-µl reaction. The reactions were electrophoresed on 0.5% SeaKem Gold agarose (FMC Bioproducts, Rockland, ME) in parallel with digoxigenin-labeled {lambda} Hind III size standards (Roche, Indianapolis, IN). The gels were transferred by capillary action to Hybond-N+ (Amersham, Piscataway, NJ) nylon membranes in 20x standard sodium citrate (SSC) and baked for 2 hours at 80°C. Nonradioactive methods were used to detect telomere sequences according to Genius system directions published by the manufacturer (Roche). In brief, the membranes were prehybridized for 4–16 hours at 37°C in hybridization buffer (50% formamide, 5x SSC, 0.1% sodium lauryl sulfate, 0.02% sodium dodecyl sulfate, or SDS, and 2% block). A synthetic oligonucleotide complimentary to human telomeric repeat sequences, (CCCTAA)3, labeled at the 3' end with digoxigenin (Genosys, Sigma) was diluted to 50 pM in hybridization buffer, and the membrane was probed for 16–24 hours at 37°C. Excess probe was removed by washing the membranes twice in 2x SSC with 0.1% SDS at ambient temperature (2 x 15 minutes), then in 0.5x SSC with 0.1 SDS at 37°C (2 x 15 minutes). A goat antidigoxigenin alkaline phosphatase-conjugated antibody and a chemiluminescent substrate, CDP-Star (Roche), were used to detect the digoxigenin-labeled probe. Autoradiograms of the blots (15- to 120-minute exposures) were digitized by using a flat bed scanner (ScanJet II CX, Hewlett Packard, Palo Alto, CA). Optical density scans of each lane were performed by using Scion Image (Scion Corp., Frederick, MD) on a personal computer. The positions of the {lambda} Hind III standard bands were plotted (log molecular weight versus relative migration distance) and the data were fitted by using a linear regression analysis (Microcal Origin). Mean telomere terminal restriction fragment lengths (MTLs) were derived as decribed (22). In brief, the standard regression line was used to calculate the molecular weight at each pixel row from the origin and to demarcate the region corresponding to 3–17 kbp. The MTL was then calculated for each lane as {Sigma}ODi/({Sigma}ODi/Li), where OD is the optical density at position i and L is the length in kilobase pairs at position i. All DNA samples were digested and analyzed on at least two gels.

Total RNA was prepared from one T-25 flask as described (41) and used as a template for reverse transcription polymerase chain reaction (RT-PCR). cDNA reactions were done by using a cDNA Cycle kit (Invitrogen, Carlsbad, CA) with polydT as a primer. Oligonucleotide primers for amplification of human GAPDH (1635'GACCCCTTCATTGAC- CTCAAC3'/4215'TGATGACCCTTTTGCTCCC3'), collagen type II (36435'AGACCTGAAACTCTGCCAC3'/41395'ACA- GTCTTGCCCCACTTAC3'), and telomerase catalytic subunit (29615'TGCGTTCTTGGCTTTCAG3'/32115'AACATGC- GTCGCAAACTC3') were used for PCR reactions, which were performed by using a "hot start" protocol and 36 cycles of 94°C (1 minute), 55°C (1 minute), and 72° (2 minutes). The products were electrophoresed on 1.2% agarose gels in parallel with {phi}X174 Hae III size standards.

Senescence-associated ß-galactosidase activity assays were performed essentially as described (42). In brief, chondrocytes were transfered to four-well chamber slides (65,000 cells per well) and incubated overnight. The cell layers were washed twice by using phosphate-buffered saline (PBS), and then fixed for 2 minutes in 2% paraformaldehyde. After three PBS washes the cell layers were overlaid with assay solution (2.0 mg/ml of X-gal in 40 mM of citric acid-sodium phosphate, pH 6.0, 5 mM of K ferricyanide, 150 mM of NaCl, 2 mM of MgCl2) and were incubated in a sealed chamber at 37°C without CO2 supplementation for 6–10 hours. The reactions were stopped by removal of the substrate and repeated washing in cold PBS. The slides were mounted and viewed on a Olympus BX60 (Olympus America, Lake Success, NY) microscope fitted with differential interference contrast optics. At least four images taken were recorded on color slide film by using a 20x objective (25–50 cells/field), and the percentage of positively stained cells in the field were scored by an observer who was unaware of sample donor age.

Incorporation of 3H-thymidine was measured by establishing three replicate cultures in 24-well plates at a concentration of 130,000 cells/well. The cells were incubated overnight in DMEM/10% FCS before the addition of fresh medium containing 5 µCi/ml of 3H-thymidine (Amersham). After 24 hours the medium was removed and the wells were washed three times for 5 minutes in PBS at 4°C before trypsinization with 0.25% trypsin, ethylenediamine tetra-acetic acid in Hanks balanced salt solution (Gibco-BRL). Cells in the trypsinized suspension were counted by using a hemacytometer, and then they were pelleted by centrifugation at 200 x g. Cell pellets were extracted by boiling for 2 minutes in 7.7 M of urea with 1% SDS. An aliquot of the extract was added to a scintillation cocktail and counted on a Beckman LSII liquid scintillation counter (Arlington Heights, IL). The total counts in the extracts were normalized to the cell number.


   Results
Top
 Abstract
 Materials and Methods
 Results
Discussion
 References
 
Primary first or second passaged chondrocyte cell strains were cultured as monolayers after isolation from articular cartilage specimens. Donor ages and sources of each of the 27 cartilage specimens used are given in Table 1 . In order to avoid in vitro telomere erosion, the cells were harvested for analyses as soon as possible after their isolation. Isolated cells (~1.0 x 106 to 4 x 106) were plated initially in T-75 flasks and then passed to three T-25 flasks after 1–5 days. Total RNA was prepared for PCR analysis from one of the three T-25 flasks. Amplification reactions for GAPDH, Type II collagen, and the telomerase catalytic subunit (hTERT) were performed, and the products were analyzed by agarose electrophoresis. Photographs of typical ethidium bromide-stained agarose gels are shown in Fig. 1. All of the primary chondrocyte cultures analyzed were positive for expression of GAPDH (258-bp product) and Type II collagen (496-bp product) but not for hTERT expression. A fibrosarcoma cell line (HT1080) did express detectable levels of hTERT message (250-bp product) but did not express Type II collagen. These analyses confirmed that the primary cultures were chondrocytic and lacked significant telomerase activity.


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  		Table 1. Sources of Articular Cartilage for Chondrocyte Cell Strains

 


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  		Figure 1. Collagen and hTERT expression in chondrocytes and fibrosarcoma cells. Ethidium bromide stained agarose gels show RT PCR reaction products for five chondrocyte strains from donors of different ages (60, 52, 1, 87, and 40 years old), a human fibrosarcoma line (HT1080), and a human chondrocarcoma cell line (CS). Samples were analyzed for expression of hTERT (t, the telomerase catalytic subunit), collagen Type II (c), and glyceraldehyde 3-phosphate dehydrogenase (GAPDH, g). Molecular weight standards ({phi}X174 Hae III digest) are shown on each gel. Sizes of selected standard bands are indicated in base pairs to the left of the first gel.

 
Chondrocyte genomic DNA was extracted from a second T-25 flask and analyzed by Southern blotting to determine mean terminal restriction fragment length (MTL). A typical Southern blot with results for 14 chondrocyte cultures is shown in Fig. 2. Each lane of the pictured blot is labeled with donor age together with the results of senescence-associated ß-galactosidase (SA ß-gal), and 3H-thymidine incorporation assays. Telomere signals from the Rsa I/Hinf 1 digested DNA samples appear as a broad smear of densities ranging in size from ~20 kbp to ~3 kbp. This heterogeneity of sizes within each sample arises in part from variation in the proximity of restriction enzyme sites to telomere sequences in different chromosomes (22). MTLs from digitized blots were calculated and plotted as a function of donor age (Fig. 3). Each data point in the plot represents the means from at least two different digestions analyzed on different gels. Error bars show standard deviations for samples analyzed at least three times (on three different blots). Coefficients of variation for replicate determinations, which were in the range of 10%, showed that the measurements were reproducible. Absolute MTL values varied from a maximum of 11,759 bp (13 year old) to a minimum of 8,731 bp (87 year old). These values fell in the range calculated for human fibroblasts and smooth muscle cells (19)(20)(31). A regression analysis showed a significant linear correlation between MTL and age (r = -.71, p = .0004).



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  		Figure 2. Telomere Southern blot. Autoradiograph of a typical Southern blot used to determine mean terminal restriction fragment lengths. Data above each lane indicates the age in years of each donor represented on the autoradiograph and the results of ß-galactosidase (ß-Gal) and 3H-thymidine (3H-Thy) incorporation assays for that donor. CPM stands for counts per minute; ND indicates that the parameter was not determined. Results for 14 primary chondrocyte strains are shown. Molecular weight standards ({lambda} Hind III digests) flank the sample lanes. Sizes of the standard bands are indicated in kilobase pairs.

 


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  		Figure 3. Mean terminal restriction fragment lengths. Mean telomere lengths (MTLs) for 27 chondrocyte strains are plotted as a function of donor age. Each data point shows the mean of at least two determinations. For clarity, error bars indicate only positive standard deviations (means from three or more determinations). The results of a regression analysis are shown (inset) and the line of best fit is drawn. Here kbp stands for kilobase pairs.

 
SA ß-gal assays were performed to determine if senescent chondrocytes accumulate with age. These assays were performed on replicate cultures of the same first or second passage chondrocytes used for telomere analyses. Blue-staining cells (SA ß-gal positives) were scored as a percentage of all cells in four or more randomly chosen 20x microscopic fields. A series of light micrographs showing staining in three different cultures is presented in Fig. 4. A cell identified as positive at 20x is pictured at 100x in Fig. 4. Examples of 20x images used for cell counting are shown in Fig. 4. Positively stained cells can be readily identified in all the images as a result of the intense blue product formed by X-gal catabolism. At high magnification (4A) the stain appears intracellularly in a punctate pattern, consistent with its lysosomal localization. A strongly positive cell is embedded in a group of at least four other cells that contained only scattered blue particles and were scored as unstained. Additional positive cells intersect the right and bottom edges of the image. The series of three 20x images shows staining patterns for the 8-year-old donor (B), the 52-year-old donor (C), and the 87-year-old donor (D). The series shows the increase in frequency of positively stained cells with increasing donor age. Means and standard deviations based on four different 20x fields were plotted as a function of donor age (Fig. 5). The minimum value across all samples (N = 15) was 4.5% positive (1 year old) and the maximum value was 55% (77 year old). A regression analysis of these data revealed a significant linear relationship between the stain and donor age (r = .80, p = .0001); the percentage of positive staining cells increased at an average rate of 4% per decade. These results indicated that ß-galactosidase expressing chondrocytes accumulate in articular cartilage as a function of age.



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  		Figure 4. In situ staining for senescence-associated ß-galactosidase activity. A, typical appearance of the stain at high magnification (100x objective); strong blue punctate staining is seen in three of the eight cells in the micrograph (52-year-old donor). B, C, and D show one of four low-magnification fields (20x objective) used to quantitate staining in each cell strain: B, 8-year-old donor (9% positive); C, 52-year-old donor (33% positive); D, 87-year-old donor (55% positive).

 


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  		Figure 5. Senescence-associated ß-galactosidase (SA ß-gal) scores. SA ß-gal activity, shown as SA ß-gal positive (%), is plotted as a function of donor age. Each point represents means and standard deviations (error bars) from four microscope fields. Linear regression parameters are shown (inset) and the line of best fit is drawn.

 
3H-thymidine incorporation assays were used to measure DNA synthesis in subconfluent cultures of first or second passaged chondrocytes. Triplicate cultures labeled for 24 hours were trypsinized and the cells counted prior to DNA extraction. Tritium counts in the extract were normalized to number of cells in each culture (CPM/cell, where CPM stands for counts per minute) and the data were plotted as a function of donor age (Fig. 6). Incorporation values ranged from 2.93 CPM/cell (1-year-old donor) to 0.62 CPM/cell (87-year-old donor). A regression analysis of the plot showed a significant negative linear relationship between the age of the donor and DNA synthesis activity (r = -.77, p = .001).



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  		Figure 6. 3H-Thymidine incorporation data plotted against donor age. The data (CPM/cell where CPM stands for counts per minute) are the means and standard deviations (error bars) from triplicate cultures. The best-fit line from a linear regression analysis is shown together with regression parameters (inset).

 
MTL data shown in Fig. 1 were plotted as a function of SA ß-gal activity (Fig. 7) or as a function of 3H-thymidine incorporation (Fig. 7) to determine if MTL correlated with these senescence markers. The plot for ß-galactosidase expression revealed a negative linear relationship (r = -.62) that was statistically significant (p = 0.01). These findings indicated that the proportion of senescent chondrocytes increases as MTL declines. Similar results were found when MTL was plotted against 3H-thymidine incorporation: A regression analysis showed a significant positive, linear relationship (r = .77, p = .001) between the two variables, indicating that incorporation declines with decreasing MTL.



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  		Figure 7. Mean telomere length (MTL) versus senscence markers. A, linear regression parameters (inset) for MTL versus senescence-associated ß-galactosidase activity, shown as SA ß-gal (% positive). B, Linear regression parameters (inset) for MTL versus 3H-thymidine incorporation; CPM = counts per minute.

 

   Discussion
Top
 Abstract
 Materials and Methods
 Results
 Discussion
References
 
We hypothesized that replicative senescence brought on by a lifetime of cell turnover contributes to the age-related changes in chondrocyte phenotype that decrease the ability of the cells to maintain articular cartilage and thereby increase the risk of osteoarthritis. Our results show that human articular cartilage chondrocytes become senescent with increasing age. In particular, we found that telomere erosion increased with the chronological age of articular chondrocyte donors and that telomere changes were linked to changes in phenotype associated with cell senescence.

Because our goal was to evaluate age-related changes that occurred in vivo, we avoided in vitro expansion of the primary chondrocyte populations used in this study. The chondrocytic nature of these populations was confirmed by RT PCR analysis for Type II collagen expression. In addition, it was essential to confirm that chondrocytes do not express the telomerase catalytic subunit that blocks telomere erosion in germ line cells as well as in many transformed cells (25)(43)(44)(45). We found that although hTERT mRNA was readily detectable in a fibrosarcoma cell line by RT PCR, it was undetectable in all 12 chondrocyte strains we tested. These data provided evidence that even young chondrocytes lack significant telomerase activity and thus, like most other somatic cells, suffer telomere erosion with each cell cycle.

Southern blot analyses of articular chondrocyte DNA from a broad range of ages (1 to 87 years) showed a significant correlation between MTL and age (p = .0004). These data ranged from 11.8 kbp for 13-year-old chondrocytes to 8.7 kbp for 87-year-old chondrocytes. The slope of the line suggests that the rate of telomere erosion for articular chondrocytes is ~22 bp/year whereas the difference between maximum and minimum values was 3028 bp/74 years, suggesting a rate approaching 40 bp/year. These rates are somewhat lower than those of vascular endothelial cells, which ranged from 47 bp/year for iliac vein cells (14- to 49-year old donors) to 102 bp/year for iliac artery cells (14- to 58-year old donors). From these results the authors concluded that hemodynamic stress in the iliac artery leads to excessive cell turnover, a high rate of telomere erosion, senescent cell accumulation, and age-related atherosclerosis. By analogy we expect that cell turnover in cartilage depends on mechanical stress exposure and injury. Because stress and the frequency of injuries vary across cartilage surfaces, even within the same joint, we expect local variations in telomere erosion rates. The relatively modest average telomere erosion we observed may reflect the focal nature of cell turnover in cartilage: Locally high concentrations of cells with very short telomeres would be expected to have only a modest impact on average telomere lengths when measurements are based on cells taken from an entire joint surface. Thus, though our chondrocyte results are not as dramatic as results for endothelial cells, the data are consistent with the hypothesis that chondrocyte turnover over the course of several decades is sufficient to induce senescence.

Chondrocyte populations used for telomere analysis were also tested for evidence of phenotypic changes associated with senescence. We found that the percentage of chondrocytes that expressed SA ß-gal, a senescence marker enzyme rose 10-fold over eight decades of donor age. These data showed a significant linear correlation between activity and age (p = .0001). SA ß-gal was expressed by some cells (<10%) in young chondrocyte populations (<15-year-old donors), a result that agrees with previous studies that reported the presence of a small number of senescent cells in somatic cell populations regardless of age (46). Mitotic activity, as measured by 3H-thymidine incorporation, also correlated with donor age (p = .0013), declining ~10-fold between the youngest (1-year-old) chondrocytes, which incorporated 2.9 CPM/cell, and the oldest (87-year-old) chondrocytes, which incorporated 0.26 CPM/cell. Significant correlations between MTL and SA ß-gal activity (p = .010) and between MTL and 3H-thymidine incorporation (p = .0046) showed that MTL erosion parallels an apparent phenotypic shift toward senescence.

Although the telomere erosion and phenotypic changes we observed appeared to be correlated with donor age, these data must be interpreted with caution. First, the apparant linear relations we found between senescence markers and donor age may be due in part to the uneven age distribution of the donors. Relatively few young and middle-aged donors were analyzed, and the resulting clustering of points at very young and old ages could lead to a false impression of linearity over the entire age range. Second, other processes such as oxidative stress and damage to DNA may induce senescence. Thus, some of the senescence we observed in chondrocyte strains, particularly those harvested from osteoarthritic donors following inflammatory episodes, may have been due to processes other than telomere erosion (47). Third, ß-galactosidase activity at pH 6.0 is a somewhat controversial senescence marker. Although our results with articular chondrocytes are similar to findings for vascular smooth muscle cells, which appear to exhibit the same strong correlation between age and activity, investigators studying human fibroblasts have been unable to observe a relationship between activity and donor age. This suggests that senescent cells accumulate in different tissues at different rates. Moreover, although ß-galactosidase activity is strongly associated with replicative senescence, it is also present in quiescent cells (48)(49), which may be common in some cartilage samples. Lastly, the apparent age-related changes we observed might have been due to other ongoing disease processes in osteoarthritic donors that were clustered in the old age range. Potentially relevant disease processes include chondrocyte "cloning," a classic histologic feature of osteoarthritis (OA) that refers to isolated clusters of chondrocytes formed by clonal expansion of a single cell. Cells within such clusters show accelerated mitotic activity (39)(40), suggesting a cause for rapid telomere erosion. Thus the rapid decline in mitotic activity and ECM synthesis typical of end-stage OA chondrocytes may reflect replicative senescence brought on by cloning, a hypothesis that might explain why OA samples typically showed shorter telomeres than nonosteoarthritic samples. Although this hypothesis indicates that replicative senescence is a result rather than a cause of OA, it also implies that the phenomenon plays an important role in the progression from early to end-stage degeneration.

The results summarized here demonstrate for the first time that chondrocyte telomeres erode in vivo in parallel with phenotypic changes associated with senescence. How these processes contribute to degenerative disease is not yet clear; however, our data strongly suggest that replicative senescence contributes to either the development or progression of OA. These observations also suggest that telomere erosion could lead to senescence in chondrocyte populations used in transplantation procedures performed to replace lost or damaged articular surfaces (50)(51)(52)(53)(54). Most protocals require in vitro expansion of the initial populations obtained from biopsy, and several population doublings may be necessary to furnish sufficient cells. Moreover, articular cartilage chondrocytes from older individuals may be less capable of proliferating and forming new tissue. Future studies in our laboratory will focus on the underlying molecular mechanisms that link telomere length, cell cycle timing, and gene expression in chondrocytes. These studies will help to elucidate the role of replicative senescence in cartilage degeneration and may shed light on the reasons for the notoriously short replicative life span of chondrocytes, which has sharply limited their availability for the treatment of cartilage defects.


   Acknowledgments
 
This work was funded by the Veterans Administration (Merit Review) and the Department of Orthopaedic Surgery at the University of Iowa. We thank Aaron Schroeder and Stacy Smith for technical assistance.

Received October 23, 2000

Accepted November 1, 2000


   References
Top
 Abstract
 Materials and Methods
 Results
 Discussion
 References
 

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