Background
Age is commonly associated with increased prevalence of tendinosis and injury [
1‐
3], and degenerative changes are commonly found in the tendons of people over 35 years of age [
4]. The most common pathology observed during surgery for chronic painful Achilles tendon is degeneration or tendinosis [
2]. In addition, most pathological changes in spontaneously ruptured tendons are degenerative [
4].
Little is known about the roles of mechanisms responsible for aging in the degeneration of tendons, but biophysical investigations have implicated a role for imbalanced homeostatic turnover of the extracellular matrix (ECM) of the tendon [
5‐
7]. Accumulated physical damage on the rotator cuff increased cleavage of matrix components in aging tendons [
8]. It appears that both insufficient synthesis and increased degradation of ECM might contribute to the mechanical deterioration of tendons.
The degree of ECM breakdown is controlled by the release of matrix metalloproteinases (MMPs) and their inhibition by tissue inhibitor of metalloproteinases (TIMPs) [
9]. Several MMPs have been implicated in chronic tendon pathologies, with increased levels of expression of MMP-1, MMP-2, MMP-9, MMP-19, MMP-23 and MMP-25, and decreased levels of expression of MMP-3, MMP-10, MMP-12, MMP-27 and TIMP-2 in either ruptured or painful tendons [
5‐
8,
10]. However, there is currently no direct evidence of an association between age and the activities of MMPs. Gelatinases (MMP-2 and −9) cleave soluble type-IV collagen [
11], as well as both native and reconstituted type-I collagen [
12]. Cyclic strain may increase the levels of both MMP-2 and MMP-9 in horse superficial digital flexor tendons and human Achilles tendons [
5]. Moreover, aging enhances this mechanically induced MMP activity [
13]. Therefore, it is crucial to investigate whether aging affects the enzymatic activities of MMP-2 and −9 and their physiologic inhibitors, TIMP-2 and −1 directly, as this could ultimately improve our understanding of the mechanism that accounts for the increasing incidence of tendinopathy in aging populations.
The transforming growth factor (TGF)-β gene family consists of at least five homologous genes that encode proteins with a wide range of effects on the differentiation and activity of many cell types [
14]. Three homodimeric isoforms (TGF-β1, -β2, and -β3) exist in mammalian cells [
15]. Studies reveal that TGF-β1 selectively stimulates the synthesis of connective tissue matrix components both
in vivo and
in vitro, to control the formation and degradation of connective tissues [
16,
17]. These effects might be augmented by reducing the synthesis of proteinases (MMPs) [
18‐
20], or by increasing the expression of tissue inhibitors of MMP (TIMPs) [
20,
21]. A study on the effects of aging on the synthesis of rabbit fibroblast matrix showed that the fibroblasts from aging rabbits produced significantly less collagen in response to TGF-β1 than fibroblasts from young rabbits did [
22]. However, whether aging alters the secretion of TGF-β in tenocytes has not yet been investigated.
The present study was undertaken to assess the effects of aging on the expression of six mRNAs, the enzymatic activities of MMP-2 and −9, and the secretion of TGF-β1 from tenocytes.
Methods
All procedures were approved by the Institutional Animal Care and Use Committee of Chung Gung Memorial Hospital, Taiwan.
Primary culture of rat Achilles tenocytes
Tenocytes were obtained from Sprague–Dawley rats, as previously described [
23]. The animals were divided into 3 groups by age: young (2 months), middle-aged (12 months), and near senescence (old, 24 months). Samples from passages 2–4, which contained fibroblasts with normal growth rates and shapes, were used. Similar cell densities were used in each group at the start of the experimental process, and all experiments were performed at least in triplicate.
3-[4,5-Dimethylthiazol-2-yl]-2,5-diphenyltetrazolium bromide (MTT) assay
Tenocytes from all age groups were cultured, and cell viability was measured by MTT assay both 24 h and 48 h after plating. After the addition of MTT (50 μg/ml), the mixture was incubated at 37°C for 1 h. Next, the MTT solution was discarded, and 1 ml of dimethyl sulfoxide (DMSO) was added to dissolve the formazan crystals. The optical density of the aliquots was measured at 570 nm OD570 nm using a spectrophotometer (VICTOR™X3 Multilabel Plate Reader; PerkinElmer Inc, Waltham, MA). Fold changes in the OD570 nm values for the middle-aged and senescent tenocytes were calculated relative to the values for young tenocytes.
Isolation of RNA, reverse transcription, and quantitative real-time polymerase chain reaction (PCR)
Tenocytes were lysed by using a guanidine isothiocyanate buffer. Subsequently, total RNA was extracted with phenol and chloroform/isoamyl alcohol (49:1) to remove proteins and genomic DNA. One microgram of total RNA was reverse-transcribed into complementary DNA (cDNA) by incubating it with 200 units of reverse transcriptase in 20 μl of reaction buffer containing 0.25 μg of random primers and 0.8 mM dNTPs at 42°C for 1 h. Quantitative real-time PCR was performed using an SYBR Green and Mx3000P™ QPCR system (Stratagene, La Jolla, CA). Aliquots (20 ng) of cDNA were used for each quantitative PCR, and each reaction was run in triplicate. The primers used are shown in Table
1. Relative gene expressions between experimental groups were determined using MxPro software (Stratagene, La Jolla, CA), and the mRNA that encodes glyceraldehyde-3-phosphate dehydrogenase (GAPDH) was used as an internal control.
Table 1
Primer sequences of target genes for real-time PCR
GAPDH
| 5’ AGTCTACTGGCGTCTTCA 3’ forward |
| 5’ TTGTCATATTTCTCGTGGT 3’ reverse |
COL1A1
| 5’ TACAGCACGCTTGTGGATG 3’ forward |
| 5’ TTGGGATGGAGGGAGTTTA 3’ reverse |
MMP-2
| 5’ GGAAGCATCAAATCGGACTG 3’ forward |
| 5’ GGGCGGGAGAAAGTAGCA 3’ reverse |
MMP-9
| 5’ CCCACTTACTTTGGAAACG 3’ forward |
| 5’ GAAGATGAATGGAAATACGC 3’ reverse |
TIMP-1
| 5’ GCCTCTGGCATCCTCTTG 3’ forward |
| 5’ CTGCGGTTCTGGGACTTG 3’ reverse |
TIMP-2
| 5’ CCAAAGCAGTGAGCGAGAA 3’ forward |
| 5’ CCCAG GGCAC AATAA AGTC 3’ reverse |
TGF-beta-1
| 5’ AGAGATTCAAGTCAAACTGTGGAG 3’ forward |
| 5’ CCAAGGTAACGCCAGGAA 3’ reverse |
Gelatin zymography
The presence of MMP-2 and MMP-9 in conditioned medium was detected using gelatin zymography, which was performed under non-reducing conditions in a 7.5% SDS-polyacrylamide gel containing 2 mg/ml gelatin (Mini-PROTEAN II system; Bio-Rad Laboratories Ltd, Hempstead, UK). Gels were washed in 2.5% Triton X-100 to remove SDS and allow renaturation of MMPs, before they were transferred to a solution containing 50 mM Tris (pH 7.5), 5 mM CaCl2, and 1 mM ZnCl2, followed by incubation at 37°C for 18 h. After staining with Coomassie brilliant blue R250 (Bio-Rad Laboratories, Hercules, CA), pro-MMPs and active MMPs were observed as white lysis bands produced by gelatin degradation. To quantify MMP-2 and MMP-9 activities, densitometric analysis was performed using 1D Digital Analysis Software (Kodak Digital Science; Eastman Kodak Company, Rochester, NY). The values of MMP-2 and MMP-9 were normalized relative to viable cell numbers determined from the MTT assay.
Enzyme-linked immunosorbent assay (ELISA)
An ELISA was used to measure the concentration of TGF-β1 in conditioned medium (culture supernatant) of tendon cells. The medium was aspirated and transferred to the wells of a 96-well ELISA plate that was pre-coated with mouse anti-TGF-β1 antibody (360 μg/ml, 100 μl/well; R&D Systems, Minneapolis, MN) overnight at room temperature, according to the manufacturer’s procedures. The plate was then read using a microplate reader set to measure absorbance at 450 nm (VICTOR™X3 Multilabel Plate Reader; PerkinElmer Inc, Waltham, MA). Recombinant TGF-β1 was serially diluted from 0 to 2000 pg/ml, and the readings were plotted to generate a standard curve. The amount of TGF-β1 production was normalized relative to viable cell numbers determined from the MTT assay after subtracted the value of culture medium.
Statistical analysis
All data from the MTT assay and densitometric analysis were expressed as mean ± SEM values. The analysis was performed with SPSS 18.0 software for Windows (SPSS Inc., Chicago, IL). Tenocytes among the three age groups were compared using the nonparametric Kruskal-Wallis test. The Mann–Whitney U-test was used for comparisons between any two groups. P values less than 0.05 were considered significant.
Discussion
Tenocytes—the basic cellular component of tendons—produce collagens, other proteins, and matrix proteoglycans [
24]. Healing of injured tendons proceeds via three overlapping stages: inflammation, regeneration, and remodeling [
24,
25]. Each stage prepares the healing process for the following stage, so the impairment of one stage may negatively impact the next one. Tenocyte proliferation is one of the principal steps in the regeneration phase of tendon healing. The results of this study indicate that tenocyte viability (which may compose of cell proliferation ability and metabolism) decreases with aging. This might partially account for the poor healing observed in aging tendons. Similar results have been obtained in studies of wound healing, where the proliferative capacity of fibroblast progressively decreases over time [
26,
27].
Matrix turnover, which involves both the synthesis and degradation of matrix components, is important for the maintenance and repair of tendons. Type-I collagen constitutes around 60% of the dry mass of the tissue and approximately 95% of total collagen [
6]. It appears that highly stressed tendons show increased levels of collagen turnover [
28]. A study of pathologic human Achilles tendon showed that levels of collagen type-I and -III mRNAs were significantly higher in tendons with chronic pain or spontaneous rupture than in normal tendons [
10]. However, the present study demonstrated that aging did not affect the level of the mRNA that encodes type-I collagen. The expression of type-I collagen mRNA is not expected to be a response of aging-related collagen degradation.
The tendon matrix is constantly remodeled throughout life. A relatively high level of matrix remodeling is common in tendons such as the supraspinatus tendon, and this process is linked to degenerative pathology [
8]. It appears that MMPs play a key role in regulating matrix remodeling, as they are considered responsible for the degradation of collagens and proteoglycans [
7,
29]. The results of our present study reveal that the activities of both MMP-2 and MMP-9 are higher in the tendons of aging rats than in the tendons of young rats. To the best of our knowledge, this is the first study to measure gelatinase activities in aging tenocytes. However, a similar age-dependent increase in MMP-2 or MMP-9 activity was reported for samples of the skin, heart, articular cartilage, and even plasma [
30‐
33]. It is reasonable to postulate that tendon degeneration may result from the aging-induced over-expression of gelatinase activity. Regarding TIMPs, our data revealed that both
TIMP-1 and
TIMP-2 mRNA expressions were decreased in old tenocytes, suggesting the activities of MMP-2 and −9 in old tenocytes, under less inhibitory effect from TIMP-1 and −2, may further have a more negative impact on the integrity of tendon matrix. These findings provide a molecular mechanism that accounts for the effect of aging on tendon healing. The over-expression of gelatinase activities may impair the turnover of matrix, which could lead to a qualitatively different and mechanically less stable tendon that is more susceptible to damage.
The transforming growth factor-β is active during almost all stages of tendon healing. During inflammation, TGF-β has a variety of effects on the regulation of cellular migration and proliferation, as well as on the stimulation of collagen production [
15]. During tendon synthesis, TGF-β1 significantly promoted the accumulation of
COL1A1 mRNA in cultured tendon fibroblasts [
34]. For tendon remodeling, TGF-β1 regulates
MMP-2 expression at the transcriptional and post-transcriptional levels by inducing an early increase in
MMP-2 transcription and an increase in the half-life of
MMP-2 mRNA [
21]. It is also thought that TGF-β exerts a selective effect on ECM deposition by modulating the action of other growth factors on metalloproteinase and
TIMP expression [
35]. Increased synthesis of TGF-β1 has also been demonstrated for tendon fibroblasts subjected to strain as well as in tendinosis fibroblast cultures [
36,
37]. However, our study demonstrated that although aging could increase the activities of MMP-2 and −9, aging is not significantly associated with
TGF-β1 expression. These observations suggest that TGF-β1 does not play a major role in either the aging process related to tendinopathy or the age-related regulation of gelatinase expression.
As for other cell culture models, there are several limitations of the model used in the present study. For instance, the behavior of explanted tendon cells is not identical to the behavior of tendon cells in their natural matrix environment
in vivo [
38,
39]. Therefore, one should always be cautious about translating culture data directly to the
in vivo situation. Further animal studies are needed to assess the physiological relevance of our findings. Aging may alter cell activity, but likely also alters the biochemical environment [
40]. It may be speculated that using a reduced level of fetal bovine serum (FBS) in culture medium might better simulate the aging condition. Although the design of the present study did not address the effects of different biochemical environment, in previous investigations, it was clearly shown that there was a decreased proliferation rate when lower level of FBS was used. Besides, independent of the levels of FBS in culture medium, there was a better proliferation in cells from young donors than cells from old donors at all times assessed [
41]. Meanwhile, immobilization has been demonstrated for an increase of catabolic process of extracellular matrix by increasing the expression of MMPs [
42,
43]. It is possible that differences in physical activity between the age groups might partly account for the findings in this study. Further study may be performed to compare the MMPs expression between the effects of inactivity and aging.
Competing interests
The authors declare that they have no competing interests.
Authors’ contributions
TY: main investigator, drafting and revision of the manuscript. JHS: main investigator, data interpretation and revision of the manuscript. KPS: data interpretation and proofreading. MCL: statistical analysis. CH: data interpretation. WC: chief supervisor, study design and revision of the manuscript. All authors read and approved the final manuscript.