HomeLarge Type Edition
HOME ARCHIVE SEARCH TABLE OF CONTENTS
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Services
Right arrow Download to citation manager
PubMed
Right arrow PubMed Citation
The Journals of Gerontology Series A: Biological Sciences and Medical Sciences 59:B324-B336 (2004)
© 2004 The Gerontological Society of America

Effects of Oxidative Damage and Telomerase Activity on Human Articular Cartilage Chondrocyte Senescence

James A. Martin1, Aloysius J. Klingelhutz2, Farid Moussavi-Harami1 and Joseph A. Buckwalter1,

1 Department of Orthopaedics and Rehabilitation
2 Department of Microbiology, University of Iowa, Iowa City.

Address correspondence to Joseph A. Buckwalter, 01008 Pappajohn Pavilion, Department of Orthopaedics, University of Iowa College of Medicine, Iowa City, IA 52242. E-mail: joseph-buckwalter{at}uiowa.edu


   Abstract
Top
 Abstract
Methods
 Results
 Discussion
 References
 
Senescence compromises the ability of chondrocytes to maintain and repair articular cartilage. We hypothesized that oxidative stress and telomere loss contribute to chondrocyte senescence. To test this hypothesis, we compared the growth of human articular cartilage chondrocytes incubated in 5% O2 and 21% O2. Cells grown in 5% O2 reached 60 population doublings (PD) before senescing, but growth in 21% O2 induced DNA damage and premature senescence at less than 40 PD. Human telomerase reverse transcriptase (hTERT)-transduction failed to prevent chondrocyte senescence in 21% O2, but allowed 1 of 3 chondrocyte strains to exceed 90 PD in 5% O2. These results show that oxidative stress causes premature chondrocyte senescence. They may help explain the increased risk of osteoarthritis with age and after joint trauma and inflammation, and suggest that minimizing oxidative damage will help produce optimal results for chondrocyte transplantation.

OSTEOARTHRITIS, the joint pain and dysfunction caused by degeneration of the articular surface, is one of the most prevalent age-related diseases (1). It is also among the most common causes of impairment for middle-aged and older people (2). Epidemiologic investigations have shown that advancing age, joint injury and inflammation, and excessive mechanical demands on the articular surfaces all increase the risk of joint degeneration (3–7). Yet, the pathogenesis of this disorder remains poorly understood (3,4,7).

With increasing age, articular cartilage chondrocyte function deteriorates and the ability of the cells to maintain or restore the tissue declines (8). The cells synthesize smaller aggrecans and less functional link proteins leading to the formation of smaller more irregular proteoglycan aggregates (9–13). Chondrocyte mitotic and synthetic activities decline with age (13–15), and they show age-related declines in response to anabolic cytokines (16–19). The counteracting effect of transforming growth factor-beta on responses to the catabolic cytokine interleukin-1 is reduced as chondrocytes age, a change that interferes with cartilage repair activities (20). In parallel with these phenotypic changes, chondrocyte telomere length declines and senescence-associated beta-galactosidase (SAß-Gal) expression increases with advancing age (14,21), suggesting that senescence contributes to the age-related deterioration of chondrocyte function (8). Two proposed mechanisms of chondrocyte senescence are telomere erosion and oxidative damage (8). Evidence that mitochondrial DNA degrades with successive cell divisions in vitro and is lost in growth-arrested chondrocytes supports the hypothesis that oxidative damage induces chondrocyte senescence (8,21).

Although the overriding risk factor for osteoarthritis is age, synovitis, joint injuries, and repetitive mechanical demands on joints increase the risk of joint degeneration and cause the early onset of this disorder (6,22–32). It is not certain how synovitis increases the risk of joint degeneration, but synovitis increases degradation of the cartilage matrix (33–36) and may lead to the release of free radicals, suggesting a link with oxidative stress (36–40). Joint injuries directly damage the articular surface (41). After a joint injury, apparently normal articular cartilage may degenerate, especially in joints with residual instability, surface incongruity, and malalignment, conditions that increase loading of the articular surface (42). The cause of joint degeneration associated with mechanical loading is unknown, but loading of articular surfaces increases the metabolic demands on chondrocytes, and in vitro mechanical stress stimulates free radical production in cartilage explants (43–45). These observations suggest that synovitis and joint injury accelerate joint degeneration by promoting oxidative damage in the cartilage matrix and in chondrocytes.

Previous studies performed in our laboratory showed that human chondrocytes senesced at 25–30 population doublings (PD) under standard culture conditions (21% O2). Chondrocytes transduced with human telomerase reverse transcriptase (hTERT) maintained long telomeres but senesced at the same time as controls, indicating that growth arrest at 25–30 PD was a form of telomere-independent, stress-induced senescence. In contrast, chondrocytes transduced with the E6 and E7 protooncogenes from human papilloma virus 16 (HPV 16) were able to bypass early senescence and grow to 60 PD. Since E6 and E7 are known to interfere with p53, and p16ink4a function, we concluded that signaling through one, or both, of these tumor suppressor activities induced early senescence. E6/E7-transduced chondrocytes were effectively immortalized by hTERT, indicating that growth arrest at 60 PD was induced by telomere-dependent replicative senescence (46). These findings were consistent with the results of human chondrosarcoma cell growth studies, which also showed that telomerase activity and tumor suppressor loss were both required for immortalization. In particular, we found that most chondrosarcoma cell strains began to senesce after less than 30 PD in 21% O2. However, in some senescent cultures, telomerase-positive cells emerged and rapidly overgrew senescent cells, forming populations that continued to proliferate for more than 120 PD. Cells in these long-lived populations remained positive for telomerase activity but did not express p16ink4a (47). Taken together, these findings indicated that human chondrocytes and chondrosarcoma cells are subject to premature, stress-induced senescence under standard culture conditions and that growth arrest at this stage depended on signaling through p16ink4a, and possibly p53.

Typical chondrocyte culture conditions use an ambient oxygen level of 21%, considerably higher than the estimated 1%–10% levels prevailing in synovial joints (48–50). A number of investigators have shown that increased ambient oxygen levels increase the generation of oxidants in cultured cells (51–53). Thus, high oxygen levels might induce sufficient oxidative stress to impair growth. A number of previous in vitro studies have investigated the effects of oxygen tension on cell growth and senescence. An early paper by Packer and Fuehr noted a 25% increase in the proliferative capacity of WI-38 and IMR-90 cells grown in 10% versus 20% O2 (54). More recently, von Zglinicki and coauthors showed that exposure of WI-38 cultures to 40% O2 for 2 weeks induced premature senescence and led to accelerated telomere erosion (55). Working with the fetal skin fibroblast line AG02258, Balin and colleagues showed that growth in 5% O2 allowed an extension of approximately 10 PD over growth in 20% O2 (56). In addition, Parrinello and colleagues showed that mouse embryo fibroblasts (MEFs) suffered premature senescence in 20% O2 but not in 3% O2 (57). This was in contrast to human fibroblasts, which did not undergo premature senescence under either condition, a result that was attributed in part to the superior DNA repair capabilities of human versus murine fibroblasts (57). The effects of O2 level on chondrocytes were investigated by Grimshaw and Mason, who found that the proliferation of bovine chondrocytes in suspension culture did not differ in 5% and 20% O2. However, this study did not investigate the long-term effects of O2 level on population growth kinetics or senescence (50). Nevertheless, the findings from this study support the hypothesis that, like MEFs, human chondrocytes are more susceptible to oxidative stress than human fibroblasts, which reach their full replicative potential (60 PD) in 21% O2.

We hypothesized that a 21% oxygen atmosphere causes sufficient oxidative stress to induce premature senescence in serially passaged chondrocytes. Moreover, we hypothesized that ectopic hTERT expression would immortalize chondrocytes grown under conditions of minimal oxidative stress. To test these hypotheses, we compared the replicative potential of chondrocytes cultured in 5% O2 versus 21% O2 and tested the effects of hTERT transduction on growth under both conditions.


   METHODS
Top
 Abstract
 Methods
Results
 Discussion
 References
 
Cell Culture
Human articular cartilage was harvested from the distal tibial or talar surfaces of normal ankle joints. Chondrocytes were incubated in a Hereaus model BB 6060 incubator (Kendro Laboratory Products, Asheville, NC) equipped with atmospheric controls (5% O2, 5% CO2, 90% N2). Chondrocytes from approximately 2 g of cartilage (2–5 x 106 cells) were seeded in primary monolayer cultures in growth medium containing 40% Dulbecco's modified Eagle medium, 40% Modified Eagle Medium-alpha, 10% Ham's F12, 10% fetal calf serum, 0.1 U/ml insulin, 25 µg/ml ascorbate, 5 µM hydrocortisone, and antibiotics (Life Technologies, Carlsbad, CA). Three separate cultures were established from 2 patients. Culture "PL" was established from distal tibial chondrocytes from a 21-year-old patient. Cultures "PLK8" and "TLK8" were established from distal tibia and talar cartilage, respectively, from a 43-year-old patient. When these primary cultures grew to confluence, the cells were passaged by sequential digestion with collagenase and trypsin as follows: Incubation at 37°C for 20–30 minutes in 0.5 mg/ml collagenase type I (Sigma, Ronkonkoma, NY) dissolved in growth medium, followed by washes with Hank's balanced salts solution (HBSS), then incubation at 37°C for 5 minutes with trypsin solution (0.25% trypsin, 1.0 mM EDTA [edetic acid] in HBSS). Trypsinized cells were recovered in growth medium and replated in 100 mm dishes at a density of ~200,000 cells/dish for viral transduction. The chondrocytes were then transduced with feline immunodeficiency virus (FIV)-hTERT or FIV beta galactosidase (control). The hTERT cDNA was isolated from pBABE-Hygro-hTERT (a gift from Dr. Robert Weinberg, Whitehead Institute, Cambridge, MA) by EcoR1/Sal 1 digestion. The fragment was subcloned using the same restriction sites in pVETL (58). pVETL-hTERT was pseudotyped with vesicular stomatitis virus-G (VSV-G), purified and produced at the University of Iowa Vector Core (B. Davidson, Director). The construction of pVETL-beta galactosidase was as described (59). Chondrocytes were transduced at a multiplicity of infection of ~10. After 24 hours of exposure, the viral infection medium was replaced with fresh growth medium, and the cells were incubated an additional 24 hours. Growth studies were initiated by passaging the transduced cultures using collagenase and trypsin and seeding 100,000 cells in 100 mm dishes. The cultures were placed in incubators with either a low O2 atmosphere (5% O2, 5% CO2, 90% N2) or high O2 atmosphere (21% O2, 5% CO2, 74% N2) and were fed once every 2–3 days with growth medium that was preequilibrated in low or high O2 incubators. This procedure was repeated just prior to reaching confluence, or every 5–7 days, for several weeks.

Senescence-Associated Beta-Galactosidase Activity
Chondrocytes were seeded in plastic 4-well chamber slides (Nalge Nunc International, Rochester, NY) at a density of 20,000 cells per well (~40% confluent). The cultures were incubated overnight before starting the SAß-Gal activity assay. The assay was performed essentially as described by Dimri and colleagues (60). In brief, the cell layers were washed twice using phosphate-buffered saline (PBS), then fixed for 2 minutes in 2% paraformaldehyde. After 3 PBS washes, the cell layers were overlaid with X-Gal assay solution 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 in Vectashield with DAPI (Vector Laboratories, Burlingame, CA) and imaged on an Olympus BX60 microscope using a digital camera (Optronics, Goleta, CA). Each field was imaged first using epifluorescence optics with ultraviolet illumination to detect DAPI (4'6-diamidino-2-phenylindole dihydrochloride)-stained nuclei, and again with transmitted light to detect SAß-Gal staining. At least 6 fields of each culture were imaged using a 20x objective (15–50 cells/field). The number of SAß-Gal-staining cells in each field was divided by the number of DAPI-stained nuclei (total cells per field) to derive % SAß-Gal positive. We noted SAß-Gal activity was at background levels in early-passaged pVETL-ß-gal transductants, indicating that prokaryotic ß-galactosidase reporter gene product was not active in the senescence assay.

Southern Blot Analyses for Mean Telomere Length and Mitochondrial DNA Deletion
DNA was isolated from ~1 x 106 to 2 x 106 cells using a DNEasy kit according to the manufacturer's directions (Qiagen, Valencia, CA). Southern blots for mean telomere length (MTL) determination were performed as previously described (14). Mitochondrial DNA Southern blots were performed using Pvu II-digested total cellular DNA as described (61), except that nonradioactive methods were used according to Genius system directions published by the manufacturer (Roche Diagnostics Corp., Indianapolis, IN). The digoxigenin-dUTP-labeled probe MP2, complimentary to human mitochondrial DNA sequences from nucleotides 526–1768 (62), was generated by polymerase chain reaction (PCR) amplification of human mitochondrial DNA with the forward primer 5'-AACCAAACCCCAAAGACAC-3', and the reverse primer 5'-CTGCTAAATCCACCTTCGAC-3' (IDT). The probe hybridizes to mitochondrial sequences common to intact chromosomes and to a 5 kbp deletion product (11.5 kbp band) commonly associated with oxidative damage (63–65). An antidigoxigenin alkaline phosphatase-conjugated antibody and a chemiluminescent substrate, CDP-Star (Roche), were used to detect the digoxigenin-labeled probe. All DNA samples were digested and analyzed on at least two gels.

Mitochondrial Density
The membrane potential-sensitive fluorescent probe DiOC6 (66) was used to assess the density of functional mitochondria in chondrocytes grown in 21% and 5% O2. A total of 20,000 cells were seeded in 4-well chamber cover slips 2 days prior to staining. DiOC6 was added to each well to a concentration of 100 nM, and the cells were incubated at 37°C for 60 minutes before imaging on an Olympus BX60 epifluorescence microscope using an Optronics digital camera.

Oxidant Production
The oxidation-sensitive dye dihydroethidium was used to detect reactive oxygen species in situ (67). TLK8 chondrocytes grown in 5% or 21% O2 for ~20 PD were stained by incubation for 30 minutes at 37°C with 5 µM dihydroethidium (Molecular Probes). One low-O2 culture was treated with 1.0 mM H2O2 for 30 minutes at 37°C, then exposed to dihydroethidium. After staining, the cultures were washed briefly with PBS, and mounted in Vectashield with DAPI (Vector Labs). The stained cells were imaged using an Olympus BX60 epifluorescence microscope and an Optronics digital camera. The same fields were imaged first using 530–550 nm excitation (for oxidized ethidium) and then using 360–370 nm excitation (for DAPI-stained nuclei). All three images of the dihydroethidium stain were taken using the same exposure time.

Comet Assays
A modified comet assay, the Fragment-Length-Analysis-using-Repair-Enzymes, or FLARE assay, was performed using a commercial kit according to the supplier's instructions (Trevigen, Gaithersburg, MD). The assay derives specificity for oxidative DNA damage through the use of the Escherichia coli DNA repair endonuclease, formamidopyrimidine-DNA glycosylase (Fpg). Fpg introduces single-strand breaks at open-ring forms of 7-methylguanine, at 8-oxoguanine, and at other oxidatively damaged nucleotides. Chondrocytes were passaged after 60–70 days of growth in either 21% or 5% O2, using collagenase, and trypsin as described above. The cells were allowed to recover after passaging by incubating the suspension in growth medium at 37°C for 30 minutes. Individual comets (n = 40–60) were imaged using 488 nm illumination and an Optronics digital camera attached to an Olympus BX60 epifluorescence microscope. Comet analysis was performed using commercially available software (CASP), which quantifies DNA degradation in terms of comet tail moment. The program was operated by setting pixel value thresholds to identify the area of the comet head (intact DNA that remains in the cell's nucleus) and comet tail (degraded DNA that electrophoresed from the nucleus). Tail moment is the product of the % DNA in the tail and tail length. Comets from cells treated with 0 U/ml Fpg (control), 0.34 U/ml Fpg, or 3.4 U/ml Fpg were analyzed. These data showed that tail moments from 0 U/ml Fpg treatment averaged ~2 units, a low value indicating a low background of DNA breakpoints in the cell preparations. A 3.4 U/ml Fpg treatment resulted in high tail moment values (~20 units) for all cells regardless of culture conditions, indicating nonspecific DNA degradation. In contrast, the lower 0.34 U/ml concentration gave a range of tail moment values for the different cell strains and was adopted for the analysis reported here.

Telomerase Activity Assay
Telomerase activity was determined using a telomere repeat amplification protocol (TRAP) assay kit (Roche) according to the supplier's directions. This assay allows relative quantitation of telomerase activity if extracts are prepared from equal numbers of cells. Accordingly, we used a standard number of cells (2500 cells per µl lysis buffer) for each determination. Relative telomerase activity (RTA) was calculated as follows: [(ODsample – ODheat inactivated sample)/ODinternal standard]/[(ODpositive control extract – ODextract buffer)/ODinternal standard].

Western Blot Analysis of p16 Expression
Western blot analysis was performed using extracts from subconfluent cultures. Cells in 100 mm dishes were extracted in 0.25 ml lysis buffer (25 mM Tris-HCl pH 7.5, 125 mM NaCl, 2.5 mM EDTA, 0.05% sodium dodecyl sulfate, 0.5% Nonidet P-40, 0.5% deoxycholate, 10% glycerol) containing protease inhibitors. Total protein concentration was determined by bicinchoninic acid assay using a commercial kit (Pierce Biotechnology, Rockford, IL). Ten micrograms of total protein in reducing buffer was fractionated on SDS-polyacrylamide gels, using a discontinuous buffer system. Western blots were performed using Immobilon-P nylon membranes (Millipore Corp., Billerica, MA). The blots were probed with the following mouse monoclonal antibodies: antihuman p16ink4a (clone G175-1239) diluted to 1.0 µg/ml (BD Pharmingen, San Diego, CA); antihuman pRb (AB-11) diluted to 1.0 µg/ml (Oncogene Research Products/Calbiochem, San Diego, CA); antihuman p53 (AB-6) diluted to 1.0 µg/ml (Calbiochem, San Diego, CA). Blots for actin were probed with a goat anti-ß-actin polyclonal antibody (I-19) diluted to 10 µg/ml (Santa Cruz Biotehnology, Inc., Santa Cruz, CA). After washing, the blots for tumor suppressors were incubated with a goat-antimouse alkaline phosphatase-conjugated secondary antibody (Promega, Madison, WI), then developed in CDP-Star (Roche) and exposed to Biomax MR autoradiography film (Eastman Kodak, Rochester, NY) for 2–15 minutes. The actin blot was washed and incubated with donkey antigoat IgG-HRP (Santa Cruz) then developed in ECL substrate (Amersham, Piscataway, NJ).

Statistical Analysis
Analysis of variance (ANOVA) with the Tukey test for multiple comparisons was used to test for significant differences in SAß-Gal activity and tail moment. The mitochondrial deletion data was analyzed using Kruskal-Wallace one-way ANOVA on ranks with Dunn's test for multiple comparisons.


   RESULTS
Top
 Abstract
 Methods
 Results
Discussion
 References
 
Human ankle chondrocytes were serially passaged in either a high-O2 atmosphere (21%) or a low-O2 atmosphere (5%), to determine the effects of O2 level on population doubling limit (PDL). Three different chondrocyte strains (PL, PLK8, and TLK8) were tested. Until ~20 PD growth rates were similar under the 2 atmospheric conditions, but after 20 PD, rates of growth in high O2 began to decline and cells from all 3 strains stopped proliferating entirely at 30–40 PD. The same cells cultured in low O2 grew at a constant rate until ~60 PD, when they too entered growth arrest (Figure 1).



View larger version (18K):
[in this window]
[in a new window]
  		Figure 1. Effects of oxygen level and human telomerase reverse transcriptase (hTERT) on population growth. Population doublings versus time-in-culture for 3 different chondrocyte strains. A, Distal tibial chondrocytes from a 21-year-old (PL). B, Distal tibial chondrocytes from a 43-year-old (PLK8). C, Talar chondrocytes from the same 43-year-old (TLK8). Cells from each strain were transduced with hTERT (TERT) or control vector (control) and incubated during serial passaging in a 5% O2 atmosphere (Low O2) or a 21% O2 atmosphere (High O2)

 
Chondrocytes were transduced with hTERT to determine if growth arrest in low-O2 or high-O2 environments was blocked by telomerase expression. Telomerase activity had a relatively modest effect on population growth in high O2: The growth of PL-hTERT cells exceeded PL control cells by ~12 PD, but TLK8-hTERT and PLK8-hTERT cells stopped growing at virtually the same time as controls (Figure 1). hTERT effects in low O2 were mixed. The strongest hTERT effect was in the PL strain, which grew to over 90 PD, or ~30 PD beyond controls, with no signs of impending growth arrest. In contrast, hTERT extended PDL by ~10 PD in PLK8 cells and had no effect in TLK8 cells. TRAP assays and southern blots were performed at 25–30 PD to confirm telomerase expression and activity in hTERT transductants. As expected, telomerase activity was detectable by TRAP assay only in hTERT-transduced cells (Figure 2A). hTERT-transduced cells also showed greater MTL than controls at 25–30 PD (p <.05), confirming telomerase function in these cells (Figure 2B). Notably, PL-hTERT cells showed longer telomeres (7.48–7.97 kbp) than were found in either the PLK8-hTERT strain (7.00–7.18 kbp) or the TLK8-hTERT strain (6.50–6.80 kbp). While the presence of hTERT increased MTL significantly over controls, there was no significant difference in MTL between cells grown in low O2 versus high O2.



View larger version (26K):
[in this window]
[in a new window]
  		Figure 2. Telomerase activity and mean telomere length. A, Telomere repeat amplification protocol assay of relative telomerase activity (RTA) in 3 chondrocyte strains (PL, PLK8, TLK8) with or without transgenic human telomerase reverse transcriptase (hTERT). B, Mean telomere length (MTL) in the same 3 strains after 25–30 PD of growth in either 5% or 21% oxygen

 
Senescence-associated ß-Gal activity was measured in all cultures after 40–50 days in culture (~30 PD) (Figure 3). Activity levels were significantly greater in high O2 within each cell strain, and the mean activity for all high-O2 cultures (38.7% positive) was significantly greater than for their low-O2 counterparts (7.61 % positive). At this stage, there was no significant difference between hTERT transductants and controls. However, SAß-Gal activity increased to 25% positive in low-O2 control cells after 70–80 days in culture (50–60 PD), just as their growth rates began to decline. Activity in hTERT-transductants remained low at this stage (6.7% positive) (Figure 4). Later assays of PLK8-hTERT and TLK8-hTERT cells at ~70 PD showed increased activity (30.5% positive and 41.0% positive, respectively) coincident with entry into growth arrest. PL-hTERT cells, which were still growing at the time, continued to express low SAß-Gal activity levels (8.3%) at 70 PD. These data showed that growth arrest in both high O2 and low O2 was associated with the accumulation of senescent chondrocytes.



View larger version (27K):
[in this window]
[in a new window]
  		Figure 3. Senescence-associated ß-Galactosidase (SAß-Gal) activity at ~30 population doublings. A, means and standard errors for individual cultures; B, pooled means and standard errors for high-O2 and low-O2 cultures. The p value above the columns indicates a significant difference between the overall means for cells grown in 21% O2 versus 5% O2. Con = control; TERT = human telomerase reverse transcriptase

 


View larger version (21K):
[in this window]
[in a new window]
  		Figure 4. Senescence-associated ß-Galactosidase (SAß-Gal) activity at ~60 population doublings. SAß-Gal activity in cells grown in 5% O2. A, Means and standard errors for hTERT-transduced and control cultures from PL, PLK8, and TLK8 strains. B, Pooled means and standard errors for hTERT and control cultures. The p value above the columns indicates a significant difference between the overall means for hTERT-transduced versus control cells

 
Age-related declines in the energy produced by liver, skeletal, and cardiac muscle cells have been strongly associated with oxidative damage to mitochondrial DNA, resulting in the accumulation of deletion products representing truncated forms of the 16.5 kbp mitochondrial chromosome. Thus, we expected that oxidative stress exposure associated with long-term growth in high O2 might have resulted in the accumulation of such deletions in serially passaged chondrocytes. To test this, we analyzed DNA from chondrocyte strains after ~50 days of growth (~30 PD), just when cells grown in high O2 began to show signs of senescence. Damage was assessed using Southern blots of total cellular DNA probed with sequences designed to detect both the intact form and common deletion mutants of the mitochondrial chromosome (Figure 5). Samples from both low-O2 and high-O2 cultures showed bands at 16.5 kbp, representing the intact chromosome, and 4 additional discreet bands (15.5, 14.2, 11.5, and 9.5 kbp), representing deletions of 1.0, 2.3, 5.0, and 7.0 kbp. Overall DNA deletion levels were estimated by calculating the proportion of the signal in the 16.5 kbp band to the sum of the signals in all 5 bands (% intact). Although high-O2 cultures as a whole showed more deletions than low-O2 cultures (61% intact vs 77% intact), the difference was not statistically significant (p =.086). Moreover, despite the apparent trend toward higher values for individual deletion products in high-O2 samples, the differences were not significant (p >=.1), indicating a relatively weak O2 effect on mitochondrial DNA integrity. In addition, one low-O2 culture (TLK8) which subsequently grew to > 60 PD, showed a low intact DNA level (54%), similar to high-O2 cultures that senesced at ~30 PD. Conversely, PL cells from high O2 entered senescence at 30 PD despite very low deletion levels (90% intact). These cases demonstrated that growth potential did not correlate with mitochondrial DNA deletion status.



View larger version (33K):
[in this window]
[in a new window]
  		Figure 5. Effects of oxygen level on mitochondrial DNA integrity. A, Southern blot showing mitochondrial DNA deletion patterns for the indicated cell strains (C = control, T = hTERT-transduced) grown in high O2 (High) or low O2 (Low) for ~50 days (~30 PD). The bands representing the intact chromosome (16.5 kbp), and 4 deletion mutants, are labeled on the right (15.5, 14.2, 11.5, and 9.5 kbp). Molecular weight markers ({lambda}Hind III) are shown on the left. The proportion of the signal in the intact, 16.5 kbp band, derived from densitometric analysis of the blot, is given at the top of each lane (% Intact). B, DNA deletion levels (% of total) from analysis of the blot shown in (A)

 
Since chondrocyte mitochondrial function could have been impaired by oxidation-induced point mutations that would not have been detected by deletion analysis, we probed living chondrocytes with DiOC6, a fluorescent indicator of mitochondrial membrane potential. Representative images of PL cells are shown (Figure 6). Micrographs of the stained cultures revealed brightly stained short strings or dots in the cytoplasm, a staining pattern characteristic of mitochondria. All chondrocyte cultures showed a similar staining pattern and intensity, indicating that the density of functional mitochondria was not affected by O2 level or hTERT. However, preincubation of cells in 10 µM sodium azide for 16 hours completely suppressed the signal, indicating specificity for mitochondrial membrane potential (data not shown). These findings suggested that mitochondrial energy production was not markedly impaired by exposure to chronic oxidative stress in high-O2-grown chondrocytes.



View larger version (126K):
[in this window]
[in a new window]
  		Figure 6. Effects of oxygen level on mitochondrial density. DiOC6-stained PL chondrocytes after ~50 days in culture (~30 PD) imaged using a 40x objective (A and B) or 80x objective (C and D). A and C: cells grown in 5% O2; B and D: cells grown in 21% O2. Mitochondria are distinguishable in the 80x image as short strings or dots in the cytoplasm

 
Oxidative damage to cellular DNA results in the accumulation of mutagenic adducts such as 7-methylguanine, and 8-oxoguanine, which might induce growth arrest in chondrocytes. We used the FLARE comet assay to determine if growth in high O2 resulted in an increase in these DNA adducts. Chondrocytes were suspended in agarose and treated with the E. coli DNA repair enzyme Fpg, which generates single-strand breaks at 7-methylguanine, and 8-oxoguanine residues. The extent of the resulting DNA fragmentation is related to the number of oxidative adducts present in cellular DNA at the time of analysis. Following Fpg treatment, the agarose-suspended cells were subjected to electrophoresis to quantify DNA fragmentation in single cells. Fluorescence images of SYBR green-stained cells taken after electrophoresis revealed a smear or "tail" of DNA streaming away from the nucleus towards the cathode (Figure 7A–C). Tail length and staining intensity derived from CASP image analysis were used to calculate tail moment, a value that increases with increasing DNA fragmentation. O2-related differences in tail moment were detected within each cell stain (Figure 7D) and between high- and low-O2 groups, which gave an overall average of 5.1 units for low-O2 cells, and 13.7 units for high-O2 cells (Figure 7E). The difference between the overall means was highly significant (p <.05), as were the differences between high O2 and low O2 within each of the cell strains (p <.05). These data demonstrated that chondrocytes sustained significantly more oxidative DNA damage in high O2 than in low O2.



View larger version (26K):
[in this window]
[in a new window]
  		Figure 7. Fragment length analysis using repair enzymes (FLARE) comet assay analysis of oxidative damage to cellular DNA. A, B, and C: Representative fluorescence images of SYBR green-stained single cells after electrophoresis. Calculated tail moments (TM) are given for each image. The cell shown in (A) was from a low-O2 PL culture. The cells shown in (B) and (C) were from a high-O2 PL culture. D, Tail moments for individual cultures (means and standard errors based on 40–60 cells). E, Overall tail moment means for all cells grown in 21% or 5% O2. The p value above the columns indicates a statistically significant difference (21% vs 5%)

 
TLK8 cells were stained with dihydroethidium (DHE) to compare oxidant generation by cells grown under high- and low-O2 conditions (Figure 8). Cells from the low-O2 culture showed low DHE staining intensity (Figure 8A). In contrast, cells in the high-O2 culture stained strongly (Figure 8C), with intensity similar to a positive control culture treated with H2O2 prior to DHE staining (Figure 8E). Images of the same fields taken under ultraviolet illumination to reveal DAPI-stained nuclei confirmed that there were similar cell densities in each field (Figure 8B, 8D, and 8F). These analyses indicated that substantially more oxidants were generated in high O2 than in low O2.



View larger version (108K):
[in this window]
[in a new window]
  		Figure 8. Generation of oxidants in high O2 and low O2. TLK8 cells stained with dihydroethidium after 20 population doublings in low O2 (A), high O2 (C), or after pretreatment with 1.0 mM H2O2 (E). The panels on the right show DAPI staining of cell nuclei in the same fields shown on the left (B, D, and F). The bar in the bottom right panel indicates 200 µm

 
Previous population growth studies indicated that the tumor suppressor p16ink4a played a crucial role in chondrocyte senescence. These studies suggested that loss of p16 was necessary for chondrocytes to bypass premature senescence at ~30 PD. Here, we analyzed p16ink4a, pRb, and p53 expression in low-O2-grown cells to determine if their ability to grow past 30 PD was associated with loss of p16ink4a or other tumor suppressors. Western blots with extracts from cells at ~50 PD (at least 10 PD after high-O2 cells had senesced) detected tumor suppressor expression in all cell strains, indicating that the extended replicative life span of cells in low O2 did not depend on loss of tumor suppressor expression (Figure 9).



View larger version (39K):
[in this window]
[in a new window]
  		Figure 9. Tumor suppressor levels in late-passaged hTERT transductants. Western blots for tumor suppressors (p16, pRb, p53) in cells grown in low O2 for 70 population doublings. Cell strains (PL, PLK8, TLK8) were either controls [C] or hTERT transduced [T]. The blot for ß-actin is included as a loading control

 

   DISCUSSION
Top
 Abstract
 Methods
 Results
 Discussion
References
 
This study showed that oxygen level is an important determinant of in vitro chondrocyte growth potential. We found that chondrocytes cultured in a 21% O2 atmosphere entered senescence prematurely at about 30 PD, much earlier than cells cultured in 5% O2. Without hTERT, low-O2 cells reached 60 PD, the doubling limit attributed to replication-dependent telomere erosion in human fibroblasts (68–71). The levels of reactive oxygen species were greater in high O2 than in low O2, and this was associated with increased oxidative damage to cellular DNA, which was twofold higher in high O2.

In high O2, hTERT transduction prevented telomere erosion but had no effect on the growth of the PLK8 or TLK8 cultures, which senesced at virtually the same time as controls. These findings demonstrated that growth arrest in high O2 was telomere-independent. However, hTERT transduction added ~10 PD to the growth of the PL culture in high O2. A number of studies suggest that telomerase can prevent premature senescence by blocking oxidative damage to telomeres. The data from these studies showed that polyguanosine sequences in telomere DNA are hypersensitive to oxidative depolymerization (72). Furthermore, oxidative stress was shown to promote telomere attrition, while the antioxidant ascorbic acid slowed the rate of age-related telomere loss in telomerase-deficient cells (73,74). We did not find a significant effect of oxidative stress on chondrocyte MTL in the current study, due perhaps to the presence of ascorbic acid in the growth medium. In any case, our data suggest that oxidation-induced telomere attrition was not a major factor in the premature senescence we observed at 30 PD. Thus, the role of telomerase in forestalling this process is still uncertain.

Under low-O2 conditions, hTERT exerted a strong growth-extending effect in PL cells, which were still proliferating after almost 90 PD, or 30 PD beyond the control PL culture. In contrast, low-O2-grown PLK8-hTERT, and TLK8-hTERT cells senesced at close to the same PD as controls, indicating a telomere-independent process. These data suggest that, although oxidative stress was undoubtedly reduced in low O2, sufficient oxidative damage accumulated over time to induce growth arrest after more than 60 PD. The basis of the extreme longevity of the PL-hTERT strain relative to the PLK8 and TLK8 strains is not known. Though it is tempting to speculate that the relatively young PL cells outperformed older PLK8 and TLK8 cells due to aging factors, our study was not designed to test this hypothesis, which would require analysis of many more cell stains from a much broader age range of samples.

Importantly, growth enhancement in low O2 did not come at the expense of abnormal tumor suppressor phenotype: p16ink4a, pRb, and p53 were present to some degree in all strains of late-passaged, low-O2-grown cells. These findings are consistent with the hypothesis suggested by Herbert and colleagues (75) that suboptimal culture conditions limit the growth-enhancing effects of telomerase. These authors showed that hTERT was insufficient to immortalize human mammary epithelial cells (HMECs) grown on plastic without concomitant loss of p16 activity. In contrast, when HMECs were grown on feeder layers to better simulate in vivo conditions, they were immortalized by hTERT without loss of p16. Similarly, our data suggest that the replicative potential of normal chondrocytes can be markedly enhanced by hTERT when the cells are cultured in low O2, a situation that approximates in vivo conditions in cartilage better than conventional high-O2 culture.

With increasing age, human chondrocytes decrease their synthetic and mitotic activity and their responsiveness to anabolic stimuli. These changes are associated with progressive telomere erosion and increased SAß-Gal activity (14,21,46). Age-related damage to chondrocyte nuclear and mitochondrial DNA and mitochondrial function may also increase the risk of apoptosis (76). Using in situ TUNEL (Tdt-mediated dUTP nick end labeling) assays we found a low frequency (<2%) of apoptotic cells in senescent cultures regardless of oxygen level (data not shown). Although this suggests that apoptosis was not a major factor in postmitotic chondrocytes, analyses of cells at earlier stages of growth might have revealed significant oxygen-related increases in apoptosis that could have contributed to growth limitations. Thus, additional work will be needed to determine the role of apoptosis in chondrocyte responses to oxidative stress.

We used monolayer cultures to study the PD limits of chondrocytes. This approach is conducive to growth studies, but not for maintenance of the differentiated chondrocyte phenotype, which requires suspension culture. Thus, we did not attempt to assess chondrocyte-specific gene expression in this study. However, such studies will be necessary to better understand the impact of stress, growth, and senescence on chondrocyte function, and to link these factors with age-related deterioration of articular cartilage.

Understanding mechanisms of chondrocyte senescence will be important in devising new approaches to the prevention and treatment of osteoarthritis. Current studies focus on two mechanisms of cell senescence: replicative senescence due to progressive telomere erosion from repeated cell division and premature senescence associated with stress due to such factors as oxidative damage to telomeres and other cell components. Although most authors have assumed that chondrocyte mitotic activity in normal human articular cartilage occurs rarely after skeletal maturity (3,4,7), some cell division has been observed (77,78), and even a slow rate of cell division might be sufficient to cause telomere erosion over decades. Furthermore, chondrocyte mitotic activity increases following cartilage injury (3,79); therefore, repetitive intense joint loading or joint trauma could accelerate telomere erosion (27). Oxidative damage provides an alternative, or complimentary, explanation for chondrocyte senescence (57,80,81). Oxidative stress-induced senescence is manifested in age-related degeneration of mitochondria (82). The respiratory activity of tissues such as bone, skeletal muscle, liver, and brain decline with age (83) due to progressive age-dependent decrease in the number and activity of mitochondria (64,84,85), a phenomenon related to alterations in the mitochondrial genome: deletions, duplications, and point mutations that disrupt expression of electron transport proteins and accumulate with age (63,65,83). These events lead to increasing free radical production as a consequence of faulty electron transport. Oxidative damage to mitochondria can initiate apoptosis through caspase activation but may also lead to irreversible growth arrest similar to replicative senescence (80). In this study, we found a slight increase in mitochondrial DNA deletions in high-O2 cultures but deletion levels did not correlate with the growth potential of the cell strains. Moreover, direct visualization of mitochondria using the membrane potential-sensitive dye DiOC6 showed a similar density of functional mitochondria in high- and low-O2 cells. On the other hand, the FLARE comet assay revealed a significant increase in oxidative damage to cellular DNA in high-O2 cultures. These data suggest that premature senescence in high O2 was attributable to point mutations induced by nuclear DNA damage rather than to mitochondrial damage. In any event, our study shows that chondrocytes are highly susceptible to oxidative stress and suggest that antioxidant treatment could be used to significantly delay senescence in vivo.

Articular cartilage may be especially vulnerable to the effects of cell senescence. After the completion of skeletal growth, no new cells enter the articular cartilage for the remainder of life (7). The cells adapt to an in vivo tissue oxygen tension of about 5% (48–50,86) and sustain the low level of metabolic activity necessary to maintain articular cartilage. In the absence of stress, they commonly succeed in preserving articular surfaces for many decades. Increased mechanical stress and tissue injuries force the cells to increase their metabolic activity (3), but the lack of a vascular supply prevents an increase in the supply of nutrients. Under stress, chondrocytes produce reactive oxygen species (87,88), and the hemarthrosis that occurs in many joint injuries (89), synovitis (39,40,90), and certain patterns of mechanical loading (45) of articular cartilage expose chondrocytes to reactive oxygen species. Progressive senescence of the limited population of highly differentiated cells adapted to low oxygen tension and low level of metabolic activity may make it increasingly difficult for these cells to maintain the tissue.

Our study shows that cultured chondrocytes are sensitive to oxygen level, and that replicative life span can be greatly extended by growth in low oxygen levels that approximate in vivo conditions. Moreover, our data show that, at least in one case, ectopic hTERT expression further enhanced growth potential to beyond 90 PD, far in excess of previously reported limits. These encouraging findings may prove useful for tissue engineering projects involving autologous chondrocytes or marrow stromal cells that have limited replicative potential. Additional investigation is needed to determine if chondrocyte senescence has an important role in the joint degeneration responsible for osteoarthritis, and if synovitis, joint injury, and excessive joint loading accelerate chondrocyte senescence. However, our results thus far suggest that chondrocyte senescence contributes to the risk of cartilage degeneration by decreasing the ability of the cells to maintain and repair the tissue (8,14,21,45), and that oxidative damage is a major cause of human chondrocyte senescence.


   Acknowledgments
 
The work reported in this article was supported by award P50 AR48939, National Institutes of Health Specialized Center of Research for OA, http://poppy.obrl.uiowa.edu/Specialized Center of Research/SCOR.htm

The authors thank Dr. Robert Weinberg, Calista Mitchell, Dr. Beverly Davidson, and the staff of Gene Transfer Vector Core at the University of Iowa for assistance with retroviral constructs. We also wish to thank Gilbert Falcon, and Joshua Boehm for their expert technical assistance.


   Footnotes
 
Estela E. Medrano,, PhD, Decision Editor

Received October 14, 2003

Accepted January 16, 2004


   References
Top
 Abstract
 Methods
 Results
 Discussion
 References
 

  1. Buckwalter JA, Heckman JD, Petrie DP., An AOA critical issue. Aging of the North American population: new challenges for orthopaedics. J Bone Joint Surg Am.. 2003;85A:748-758.
  2. Praemer A, Furner S, Rice DP. Musculoskeletal Conditions in the United States. Rosemont IL: American Academy of Orthopaedic Surgeons; 1999.
  3. Buckwalter JA, Mankin HJ. Articular cartilage II. Degeneration and osteoarthrosis, repair, regeneration and transplantation. J Bone Joint Surg.. 1997;79A:612-632.
  4. Buckwalter JA, Martin JA, Mankin HJ. Synovial joint degeneration and the syndrome of osteoarthritis. Instructional Course Lectures.. 2000;49:481-489.[Medline]
  5. Buckwalter JA, Lappin DR. The disproportionate impact of chronic arthralgia and arthritis among women. Clin Orthop Rel Res.. 2000;372:159-168.
  6. Felson DT. The epidemiology of osteoarthritis: prevalence and risk factors. In: KE Kuettner, VM Goldberg, eds. Osteoarthritic Disorders. Rosemont, IL: American Academy of Orthopaedic Surgeons; 1995:13–24.
  7. Buckwalter JA, Mankin HJ. Articular cartilage I. Tissue design and chondrocyte-matrix interactions. J Bone Joint Surg.. 1997;79A:600-611.
  8. Martin JA, Buckwalter JA. The role of chondrocyte senescence in the pathogenesis of osteoarthritis and in limiting cartilage repair. J Bone Joint Surg Am.. 2003;85A:(Suppl 2): B106-110.
  9. Buckwalter JA, Choi H, Tang L, Rosenberg L, Ungar R. The effect of link protein concentration of proteoglycan aggregation. Trans 32nd Meeting Ortho Res Soc.. 1986;11:98.
  10. Buckwalter JA, Rosenberg LC. Electron microscopic studies of cartilage proteoglycans. Elec Microsc Rev.. 1988;1:87-112.
  11. Buckwalter JA, Roughley PJ, Rosenberg LC. Age-related changes in cartilage proteoglycans: quantitative electron microscopic studies. Micros Res Tech.. 1994;28:398-408.[Medline]
  12. Buckwalter JA, Rosenberg LC. Electron microscopic studies of cartilage proteoglycans: Direct evidence for the variable length of the chondroitin sulfate rich region of the proteoglycan subunit core protein. J Biol Chem.. 1982;257:9830-9839.[Free Full Text]
  13. Verbruggen G, Cornelissen M, Almqvist KF, Wang L, Elewaut D, Broddelez C, de Ridder L, Veys EM. Influence of aging on the synthesis and morphology of the aggrecans synthesized by differentiated human articular chondrocytes. Osteoarthritis Cartilage.. 2000;8:170-179.[Medline]
  14. Martin JA, Buckwalter JA. Telomere erosion and senescence in human articular cartilage chondrocytes. J Gerontol Biol Sci.. 2001;56A:B172-B179.
  15. Bolton MC, Dudhia J, Bayliss MT. Age-related changes in the synthesis of link protein and aggrecan in human articular cartilage: implications for aggregate stability. Biochem J.. 1999;337:77-82.
  16. Martin JA, Ellerbroek SM, Buckwalter JA. The age-related decline in chondrocyte response to insulin-like growth factor-I: the role of growth factor binding proteins. J Ortho Res.. 1997;15:491-498.[Medline]
  17. Martin JA, Buckwalter JA. The role of chondrocyte-matrix interactions in maintaining and repairing articular cartilage. Biorheology.. 2000;37:129-140.[Medline]
  18. Loeser RF, Shanker G, Carlson CS, Gardin JF, Shelton BJ, Sonntag WE. Reduction in the chondrocyte response to insulin-like growth factor 1 in aging and osteoarthritis: studies in a non-human primate model of naturally occurring disease. Arthritis Rheum.. 2000;43:2110-2120.[Medline]
  19. Guerne PA, Blanco F, Kaelin A, Desgeorges A, Lotz M. Growth factor responsiveness of human articular chondrocytes in aging and development. Arthritis Rheum.. 1995;38:960-968.[Medline]
  20. Scharstuhl A, van Beuningen HM, Vitters EL, van der Kraan PM, van den Berg WB. Loss of transforming growth factor counteraction on interleukin 1 mediated effects in cartilage of old mice. Ann Rheum Dis.. 2002;61:1095-1098.[Abstract/Free Full Text]
  21. Martin JA, Buckwalter JA. Human chondrocyte senescence and osteoarthritis. Biorheology.. 2002;39:145-152.[Medline]
  22. Felson DT, Hannan MT, Naimark A, Berkley J. Occupational physical demands, knee bending, and knee osteoarthritis: results from the Framingham study. J Rheumatol.. 1991;18:1587-1592.[Medline]
  23. Felson DT, Zhang Y, Anthony JM, Naimark A, Anderson JJ. Weight loss reduces the risk for symptomatic knee osteoarthritis in women. Ann Intern Med.. 1992;116:535-539.
  24. Felson DT. Occupation and knee osteoarthritis: evidence from two population-based studies. Acta Orthop Scand.. 1993;64:(Suppl 253): B56-57.
  25. Buckwalter JA. Osteoarthritis and articular cartilage use, disuse and abuse: experimental studies. J Rheumatol.. 1995;22:(Suppl 43): B13-15.
  26. Buckwalter JA, Lane NE. Aging, sports and osteoarthritis. Sports Med Arth Rev.. 1996;4:276-287.
  27. Buckwalter JA, Lane NE. Athletics and osteoarthritis. Am J Sports Med.. 1997;25:873-881.[Abstract/Free Full Text]
  28. Axmacher B, Lindberg H. Coxarthrosis in farmers. Acta Orthop Scand.. 1993;64:(Suppl 253): B59.
  29. Lindberg H, Montgomery F. Heavy labor and the occurrence of gonarthrosis. Clin Orthop Rel Res.. 1987;214:235-236.
  30. Lindberg H, Roos H, Gardsell P. The relationship between soccer and coxarthrosis. Acta Orthop Scand.. 1993;64:(Suppl 253): B60.
  31. Roos H, Lindberg H, Gardsell P, Wingstrand H. The prevalence of gonarthrosis in former soccer players and its relationship to menisectomy. Acta Orthop Scand.. 1993;64:(Suppl 253): B60.
  32. Maxian TA, Brown TD, Weinstein SL. Chronic stress tolerance levels for human articular cartilage: two nonuniform contact models applied to long-term follow-up of CDH. J Biomech.. 1995;28:159-166.[Medline]
  33. Elsaid KA, Jay GD, Chichester CO. Detection of collagen type II and proteoglycans in the synovial fluids of patients diagnosed with non-infectious knee joint synovitis indicates early damage to the articular cartilage matrix. Osteoarthritis Cartilage.. 2003;11:673-680.[Medline]
  34. Ishiguro N, Ito T, Oguchi T, Kojima T, Iwata H, Ionescu M, Poole AR. Relationships of matrix metalloproteinases and their inhibitors to cartilage proteoglycan and collagen turnover and inflammation as revealed by analyses of synovial fluids from patients with rheumatoid arthritis. Arthritis Rheum.. 2001;44:2503-2511.[Medline]
  35. Goldbach-Mansky R, Lee JM, Hoxworth JM, et al. Active synovial matrix metalloproteinase-2 is associated with radiographic erosions in patients with early synovitis. Arthritis Res.. 2000;2:145-153.[Medline]
  36. Smith MD, Triantafillou S, Parker A, Youssef PP, Coleman M. Synovial membrane inflammation and cytokine production in patients with early osteoarthritis. J Rheumatol.. 1997;24:365-371.[Medline]
  37. Harris ED, Jr. The rationale for combination therapy of rheumatoid arthritis based on pathophysiology. J Rheumatol Suppl.. 1996;44:2-4.[Medline]
  38. Edmonds SE, Blake DR, Morris CJ, Winyard PG. An imaginative approach to synovitis—the role of hypoxic reperfusion damage in arthritis. J Rheumatol Suppl.. 1993;37:26-31.[Medline]
  39. Mapp PI, Grootveld MC, Blake DR. Hypoxia, oxidative stress and rheumatoid arthritis. Br Med Bull.. 1995;51:419-436.[Abstract/Free Full Text]
  40. Nagai H, Kumamoto H, Fukuda M, Takahashi T. Inducible nitric oxide synthase and apoptosis-related factors in the synovial tissues of temporomandibular joints with internal derangement and osteoarthritis. J Oral Maxillofac Surg.. 2003;61:801-807.[Medline]
  41. Buckwalter JA. Mechanical injuries of articular cartilage. In: G Finerman G, ed. Biology and Biomechanics of the Traumatized Synovial Joint. Park Ridge, IL: American Academy of Orthopaedic Surgeons; 1992:83–96.
  42. Marsh JL, Buckwalter JA, Gelberman R, et al. AOA Symposium. Articular fractures: does an anatomic reduction really change the result? J Bone Joint Surg.. 2002;84A:1259-1271.
  43. Heiner AD, Martin JA, Brown TD. Cartilage responses to stress in a novel triaxial mechanical stress culture system. Transcript of the 4th Combined Orthopaedic Research Societies Meeting, 2001:141.
  44. Heiner AD, Martin JA. Cartilage responses to a novel triaxial mechanostimulatory culture system. J Biomech. In Press.
  45. Martin JA, Brown T, Heiner A, Buckwalter JA. Post-traumatic osteoarthritis: the role of accelerated chondrocyte senescence. Biorheology. In Press.
  46. Martin JA, Mitchell CJ, Klingelhutz AJ, Buckwalter JA. Effects of telomerase and viral oncogene expression on the in vitro growth of human chondrocytes. J Gerontol Biol Sci.. 2002;57A:B48-B53.
  47. Martin JA, Forest E, Block JA, et al. Malignant transformation in human chondrosarcoma cells supported by telomerase activation and tumor suppressor inactivation. Cell Growth Diff.. 2002;13:397-407.[Abstract/Free Full Text]
  48. Grimshaw MJ, Mason RM. Modulation of bovine articular chondrocyte gene expression in vitro by oxygen tension. Osteoarthritis Cartilage.. 2001;9:357-364.[Medline]
  49. Murphy CL, Sambanis A. Effect of oxygen tension and alginate encapsulation on restoration of the differentiated phenotype of passaged chondrocytes. Tissue Eng.. 2001;7:791-803.[Medline]
  50. Grimshaw MJ, Mason RM. Bovine articular chondrocyte function in vitro depends upon oxygen tension. Osteoarthritis Cartilage.. 2000;8:386-392.[Medline]
  51. DeGroot H, Littauer A. Hypoxia, reactive oxygen, and cell injury. Free Radic Biol. Med.. 1989;173:541-551.
  52. Yusa T, Crapo JD, Freeman BA. Hyperoxia enhances lung and liver nuclear superoxide generation. Biochim Biophys Acta.. 1984;798:167-174.[Medline]
  53. Turrens JF, Freeman BA, Levitt JG, Crapo JD. The effect of hyperoxia on superoxide production by lung submitochondrial particles. Arch Biochem Biophys.. 1982;217:401-410.[Medline]
  54. Packer L, Fuehr K. Low oxygen concentration extends the lifespan of cultured human diploid cells. Nature.. 1977;267:423-425.[Medline]
  55. von Zglinicki T, Saretzki G, Docke W, Lotze C. Mild hyperoxia shortens telomeres and inhibits proliferation of fibroblasts: a model for senescence? Exp Cell Res.. 1995;220:186-193.[Medline]
  56. Balin AK, Fisher AJ, Anzelone M, Leong I, Allen RG. Effects of establishing cell cultures and cell culture conditions on the proliferative life span of human fibroblasts isolated from different tissues and donors of different ages. Exp Cell Res.. 2002;274:275-287.[Medline]
  57. Parrinello S, Samper E, Krtolica A, Goldstein J, Melov S, Campisi J. Oxygen sensitivity severely limits the replicative lifespan of murine fibroblasts. Nat Cell Biol.. 2003;5:741-747.[Medline]
  58. Johnston JC, Gasmi M, Lim LE, et al. Minimum requirements for efficient transduction of dividing and nondividing cells by feline immunodeficiency virus vectors. J Virol.. 1999;73:4991-5000.[Abstract/Free Full Text]
  59. Hughes SM, Moussavi-Harami F, Sauter SL, Davidson BL. Viral-mediated gene transfer to mouse primary neural progenitor cells. Mol Ther.. 2002;5:16-24.[Medline]
  60. Dimri G, Lee X, Basile G, et al. A biomarker that identifies senescent human cells in culture and in aging skin in vivo. Proc Natl Acad Sci U S A.. 1995;92:9363-9367.[Abstract/Free Full Text]
  61. Varanasi SS, Datta HK. Southern analysis of mitochondrial DNA in cortical bone of elderly patients undergoing knee and hip arthroplasty. J Pathol.. 2001;193:557-562.[Medline]
  62. Tengan CH, Moraes CT. Detection and analysis of mitochondrial DNA deletions by whole genome PCR. Biochem Mol Med.. 1996;58:130-134.[Medline]
  63. Kang CM, Kristal BS, Yu BP. Age-related mitochondrial DNA deletions: effect of dietary restriction. Free Rad Biol Med.. 1998;24:148-154.[Medline]
  64. Kopsidas G, Kovalenko SA, Kelso JM, Linnane AW. An age-associated correlation between cellular bioenergy decline and mtDNA rearrangements in human skeletal muscle. Mutation Res.. 1998;421:27-36.
  65. Lee CM, Weindrich R, Aiken JM. Age-related alterations of the mitochondrial genome. Free Rad Biol Med.. 1997;22:1259-1269.[Medline]
  66. Rajpurohit R, Mansfield K, Ohyama K, Ewert D, Shapiro IM. Chondrocyte death is linked to development of a mitochondrial membrane permeability transition in the growth plate. J Cell Physiol.. 1999;179:287-296.[Medline]
  67. Li W, Miller F, Brown M, et al. Enhanced H2O2-induced cytotoxicity in "epithelioid" smooth muscle cells implications for neointimal regression. Arterioscler Thromb Vasc Biol.. 2000;20:1473-1479.[Abstract/Free Full Text]
  68. Hayflick L, Moorhead PS. The serial cultivation of human diploid cell strains. Exp Cell Res.. 1961;25:585-621.
  69. Hayflick L. The limited in vitro lifetime of human diploid cell strains. Exp Cell Res.. 1965;37:614-636.[Medline]
  70. Hayflick L. The longevity of cultured human cells. J Am Geriatr Soc.. 1974;22:1-12.[Medline]
  71. Allsopp RC, Chang E, Kashefi-Aazam M, et al. Telomere shortening is associated with cell division in vitro and in vivo. Exp Cell Res.. 1995;220:194-200.[Medline]
  72. Liu L, Trimarchi JR, Navarro P, Blasco MA, Keefe DL. Oxidative stress contributes to arsenic-induced telomere attrition, chromosome instability and apoptosis. J Biol Chem. 2003;278:31998–2004.
  73. Furumoto K, Inoue E, Nagao N, Hiyama E, Miwa N. Age-dependent telomere shortening is slowed down by enrichment of intracellular vitamin C via suppression of oxidative stress. Life Sci.. 1998;63:935-948.[Medline]
  74. Szalai VA, Singer MJ, Thorp HH. Site-specific probing of oxidative reactivity and telomerase function using 7,8-dihydro-8-oxoguanine in telomeric DNA. J Am Chem Soc.. 2002;124:1625-1631.[Medline]
  75. Herbert BS, Wright WE, Shay JW. p16(INK4a) inactivation is not required to immortalize human mammary epithelial cells. Oncogene.. 2002;21:7897-7900.[Medline]
  76. Mobasheri A. Role of chondrocyte death and hypocellularity in ageing human articular cartilage and the pathogenesis of osteoarthritis. Med Hypotheses.. 2002;58:193-197.[Medline]
  77. Aigner T, Hemmel M, Neureiter D, et al. Apoptotic cell death is not a widespread phenomenon in normal aging and osteoarthritis human articular knee cartilage: a study of proliferation, programmed cell death (apoptosis), and viability of chondrocytes in normal and osteoarthritic human knee cartilage. Arthritis Rheum.. 2001;44:1304-1312.[Medline]
  78. Kusuzaki K, Sugimoto S, Takeshita H, et al. DNA cytofluorometric analysis of chondrocytes in human articular cartilages under normal aging or arthritic conditions. Osteoarthritis Cartilage.. 2001;9:664-670.[Medline]
  79. Buckwalter JA, Mow VC, Hunziker EB. Concepts of cartilage repair in osteoarthritis. In: R Moskowitz, VM Goldberg, D Howell, R Altman, JA Buckwalter, eds. Osteoarthritis: Diagnosis and Medical/Surgical Management. 3rd Ed. Philadelphia, PA: Saunders; 2001:101–114.
  80. Toussaint O, Medrano E, Zglinicki Tv. Cellular and molecular mechanisms of stress-induced premature senescence (sips) of human diploid fibroblasts and melanocytes. Exp Gerontol.. 2000;35:927-945.[Medline]
  81. Dumont P, Chen QM, Burton M, et al. Induction of replicative senescence biomarkers by sublethal oxidative stress in normal human fibroblasts. Free Rad Biol Med.. 2000;28:361-373.[Medline]
  82. Golden T, Melov S. Mitochondrial DNA mutations, oxidative stress, and aging. Mech Ageing Dev.. 2001;122:1577-1589.[Medline]
  83. Arnheim N, Cortopassi G. Deleterious mitochondrial DNA mutations accumulate in aging human tissues. Mutation Res.. 1992;275:157-167.
  84. Beckman KB, Ames BN. Endogenous oxidative damage of mtDNA. Mutat Res.. 1999;424:51-58.[Medline]
  85. Beckman KB, Ames BN. Oxidative decay of DNA. J Biol Chem.. 1997;272:19633-19636.[Free Full Text]
  86. Rajpurohit R, Koch CJ, Tao Z, Teixeira CM, Shapiro IM. Adaptation of chondrocytes to low oxygen tension: relationship between hypoxia and cellular metabolism. J Cell Physiol.. 1996;168:424-432.[Medline]
  87. Henrotin Y, Deby-Dupont G, Deby C, De Bruyn M, Lamy M, Franchimont P. Production of active oxygen species by isolated human chondrocytes. Br J Rheumatol.. 1993;32:562-567.[Abstract/Free Full Text]
  88. Mazzetti I, Grigolo B, Pulsatelli L, et al. Differential roles of nitric oxide and oxygen radicals in chondrocytes affected by osteoarthritis and rheumatoid arthritis. Clin Sci (Lond).. 2001;101:593-599.[Medline]
  89. Roosendaal G, Vianen ME, Marx JJ, van den Berg HM, Lafeber FP, Bijlsma JW. Blood-induced joint damage: a human in vitro study. Arthritis Rheum.. 1999;42:1025-032.[Medline]
  90. Schett G, Tohidast-Akrad M, Steiner G, Smolen J. The stressed synovium. Arthritis Res.. 2001;3:80-86.[Medline]




This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Services
Right arrow Download to citation manager
PubMed
Right arrow PubMed Citation

HOME ARCHIVE SEARCH TABLE OF CONTENTS