Keywords

Epigenetic reprogramming, chondrocyte rejuvenation, osteoarthritis, small chemical molecules, cartilage regeneration

Aging is a complex biological process characterized by the gradual decline of tissue and organ function. Among the various theories of aging, the epigenetic theory plays a key role, proposing that the accumulation of changes in the cellular epigenetic profile—such as DNA methylation and histone modifications—leads to dysregulation of gene expression essential for maintaining tissue homeostasis [1-5]. These changes are particularly critical in tissues with low regenerative capacity, such as articular cartilage, where epigenetic dysregulation has been implicated in the progression of osteoarthritis (OA) [6, 7]. OA is the most common joint disease, significantly reducing quality of life, especially in older individuals. Its pathogenesis involves hyaline cartilage degradation, chronic inflammation, and synovitis [8, 9]. One of the major risk factors for OA in women is postmenopausal estrogen deficiency, which exacerbates inflammatory and degenerative processes in cartilage and accelerates disease progression [10, 11].
Current therapeutic approaches primarily aim to relieve pain and improve joint function but fail to prevent disease progression or address its underlying causes [12-14]. As a result, strategies targeting epigenetic reprogramming of chondrocytes—the primary cells responsible for maintaining cartilage integrity—are gaining increased attention [15]. Several studies have demonstrated the potential of epigenetic rejuvenation in cell cultures undergoing replicative senescence, using small chemical molecules (SCMs) [16, 17]. These molecules modulate key epigenetic mechanisms, including chromatin remodeling and DNA methylation, thereby restoring a transcriptional profile characteristic of a more youthful cellular state. In vitro, epigenetic modifiers have been shown to enhance cell proliferation and reduce the expression of pro-inflammatory markers [18, 19].
While most evidence to date comes from in vitro systems, there is a growing need to validate these findings in vitro, particularly in the context of degenerative diseases such as OA. To address this, we developed a preclinical rat model of age-related OA to assess the regenerative potential of intra-articularly administered SCMs. To better simulate the human condition, we selected female rats over two years of age, roughly equivalent to humans over 60. This choice was based on their accelerated aging profile and the high relevance of postmenopausal OA in women. The model allowed investigation of epigenetic interventions in a physiologically relevant setting within a manageable experimental timeframe. To further explore the role of hormonal status, we used the estrogen receptor blocker clomiphene citrate. As previously demonstrated [20], this agent effectively induces postmenopausal conditions in rats. Compared to surgical ovariectomy, this pharmacological method offers the advantage of preserving animal mobility and reducing potential confounding effects on study outcomes. Estrogen receptors mediate hormonal effects across various tissues [21-23], and their blockade enabled assessment of the interaction between SCM-induced epigenetic reprogramming and estrogen deficiency—a critical factor in the pathogenesis of age-related OA.
The aim of this study was to evaluate the therapeutic potential of SCMs in promoting cartilage regeneration through epigenetic reprogramming in an in vitro model of age-related OA. Special attention was given to the structural and biochemical restoration of cartilage, as well as to the modulation of chondrocyte activity and the immune response under conditions simulating human aging and hormonal decline.

Laboratory animals

Experiments were performed on female white rats (Rattus norvegicus albinus) weighing 250–270 g. The selected animals were fertile females, classified as at least class 2 (i.e., having birthed no fewer than five pups per litter), with regular estrous cycles every 6–10 days. All animals were housed and fed under standard vivarium conditions in accordance with the recommendations of the Ethical Committee of Uzbekistan (2014) and the Ethical Guidelines for the Use of Animals in Research (2019). Ethical approval and a completed ARRIVE checklist are included in the supplementary materials.

Study design

The experiment was conducted in two stages:
Stage I: Model Development
To determine the optimal inducer of knee joint OA, 34 female rats were randomly divided into four groups: three experimental groups (n = 8 each) and one control group (n = 10). Group 1 received 0.5 mL of a 10% talc suspension, Group 2 received 0.1 mL of 0.1% trypsin, and Group 3 received 0.1 mL of 1% papain. All injections were administered once intra-articularly under sterile conditions. The control group received 0.5 mL of sterile saline to account for injection-related stress.
Stage II: Main Experiment
Fifty-six rats were randomly allocated into six groups: Group I – Intact control (n = 6), Group II – Intact + SCM, Group III – OA + postmenopause, Group IV – OA + postmenopause + SCM, Group V – OA + postmenopause + chondroitin sulfate, and Group VI – OA + SCM. OA was induced by intra-articular injection of trypsin. Postmenopausal state was pharmacologically modeled using clomiphene citrate (10 mg/kg/day orally, 2 courses of 5 days).

Small chemical molecule set (SCM)

SCMs were administered intra-articularly at concentrations previously validated in in vitro VC6TF experiments for epigenetic reprogramming (Supplementary Table 1) [16].

Modeling the postmenopausal state

To simulate postmenopausal conditions, clomiphene citrate—a selective estrogen receptor modulator—was used. As previously demonstrated [20], clomiphene competitively binds to estrogen receptors on chondrocytes, effectively inducing a postmenopausal-like state in experimental animals.

Biochemical and cytokine analysis

Venous blood from the sublingual vein was collected in vacutainers with a coagulation activator (silicon dioxide), then centrifuged for 15 min at 3000 rpm. The level of cytokine expression in the serum of experimental animals was determined by the ELISA method (test systems of ZAO Vector-Best in KT-I, KT-II and KT-III). ALT and ALP on a biochemical analyzer (BioChem SA) with a built-in thermostat ((High Technology, Inc). Vitamin D immunochemiluminescent assay ELISA (CTK Biotech, Inc, USA). Calcium, Phosphorus and Alkaline Phosphatase were determined by a colorimetric photometric method (Human ELISA, Germany). C-reactive protein was determined by the agglutination method (kit CRP Latex Cypress Diagnostics, Belgium)

Histological and histochemical methods

Following euthanasia, the right knee joints, including the distal femur and proximal tibia, were dissected for histomorphological analysis of cartilage and bone [24, 25].
Three staining techniques were used: hematoxylin and eosin (H&E), Van Gieson staining, and the periodic acid–Schiff (PAS) reaction. These techniques were selected to highlight structural features relevant to evaluating epigenetic reprogramming. Collected tissue samples (right lower limbs) were coded and processed group-wise. Samples were embedded in Histomix paraffin (BioVitrum, Russia), and 5–8 μm sections were cut using a microtome. Microscopic evaluation was performed using a ZEISS Primo Star light microscope. A blinded histopathologist acquired digital microphotographs. Further image analysis was conducted using a NanoZoomer Hamamatsu system (Japan) at 50x, 100x, 200x, and 400x magnification. Digital images were captured with NanoZoomer software (REF C13140-21.S/N000198/HAMAMATSU PHOTONICS/431-3196 JAPAN). A morphometric study of tissue architecture was performed using a multiplex confocal system. The QuPath-0.5.0 software was used to generate two-dimensional images. These were analyzed along X and Y axes to calculate surface areas and lesion extent. Specific zones of articular cartilage were identified to assess the degree of tissue involvement.

Severity assessment: Modified Mankin scale

Cartilage degeneration was scored using the Modified Mankin scale [26], with total scores ranging from 0 (normal) to 12 (severe damage). The following parameters were assessed: surface structure (1—irregularity/erosion, 2—cracks, 3—delamination), cellularity (1—slight reduction in chondrocyte number, 2—significant reduction, 3—complete absence), staining intensity (1—mild loss, 2—marked loss, 3—no staining), and cell proliferation (1—isogenic groups with two cells, 2—two to three cells per group, 3—proliferation foci with more than three cells per group). Nuclear–Cytoplasmic Index (N:C ratio) was measured in superficial and middle zones of the articular cartilage. For each sample, 100 chondrocytes per zone were measured using calibrated image analysis software.

Statistical analysis

Quantitative data were expressed as mean ± standard deviation. Statistical analyses were performed using MS Excel 2007 and STATISTICA (Windows 10). Within-group and between-group comparisons were made using Student’s t-test. For correlation analyses, Pearson’s coefficient was used for normally distributed data and Spearman’s rank coefficient for non-parametric data. A P-value of < 0.05 was considered statistically significant.

To study the effect of SCM on joint tissues in OA, a series of experiments were conducted in two stages: Stage I—modeling OA in rats, and Stage II—evaluating the effect of SCMs on cartilage tissue in aged, postmenopausal rats (Figure 1).

Stage I: optimization of the OA model

The general scheme of the first-stage experiment is presented in Supplementary Figure 1.For 60 days following the administration of OA inducers, the clinical condition of the animals was monitored, including body weight, behavioral responses (activity, mobility, lameness, gait), functional tests ("open field" test, sprint test), and biochemical and immunological markers (calcium, phosphorus, vitamin D, alkaline phosphatase, IL-1β, TNF-α, IL-6, IL-8, and C-reactive protein).
The first clinical signs of developing gonarthrosis were observed as early as the second week, in the form of decreased activity, noticeable lameness, dragging of the right hind limb, and unsteady gait. Functional and motor activity, assessed using the "open field" and "sprint" tests, was most reduced in Group 3 (trypsin), where the lowest levels of horizontal and vertical activity, as well as movement speed, were recorded.
Among the biochemical parameters, increased serum calcium and phosphorus levels were observed in all groups with induced osteoarthrosis compared to the control group. In addition, alkaline phosphatase levels were significantly elevated in the papain and trypsin groups (P < 0.05), but not in the talc group—likely due to the presence of aluminum in talc (Supplementary Figure 2). Vitamin D concentrations remained unchanged across all groups. C-reactive protein levels significantly increased in all experimental groups (P < 0.05), indicating an acute phase of inflammation. In this study, CRP levels rose sharply by day 30, reaching a plateau by the end of the experiment (Supplementary Figure 3).
By day 30, serum levels of IL-6 and TNF-α peaked in the trypsin group (P < 0.01), reflecting an acute inflammatory response. IL-8 levels showed no significant difference in Group II (talc) compared to the control, but Groups III and IV exhibited a significant increase. A substantial elevation in IL-1β levels was recorded in all experimental groups (Supplementary Figure 4).
Thus, immunobiochemical analysis revealed a marked increase in proinflammatory interleukins by day 30, especially in the enzyme-induced OA groups (Groups III and IV). By day 60, however, cytokine and CRP levels stabilized, with only a slight further increase, suggesting that a stable OA phenotype had developed by day 30.
Following euthanasia, 34 rats were removed from the experiment for histomorphological examination of cartilage tissue. Histological images from the same anatomical regions in different groups were compared. Representative images are shown in Supplementary Table 2. In Group III, the medial, lateral, and central areas of the hyaline cartilage surface exhibited erosion and desquamation, along with marked chondrocyte proliferation and variable surface roughness and thickness (Supplementary Figure 5). Evidence of pannus formation was observed in the form of proliferating chondroblasts, surrounded by isogenic chondrocyte groups with uneven vacuolar degeneration. The intercellular matrix showed reduced homogeneity, with erosive-destructive changes extending to a depth of 1.025 mm. In the joint cavity, synoviocyte desquamation and accumulation of free-floating synoviocytes were observed, along with narrowing of the joint space, consistent with reactive synovitis. Dystrophic and degenerative changes were also found in the epimetaphyseal region, with resorptive cystic lesions in the metaphyseal trabeculae.
Given that spontaneous OA does not naturally develop in rats [27], it is reasonable to conclude that the observed joint damage was directly caused by OA inducer administration. Histological analysis confirmed that the most pronounced tissue alterations occurred in Groups III (trypsin) and IV (papain). These included surface erosions, fissures, ulcerations, uneven hyaline cartilage thickness, matrix vacuolization, reduced staining intensity, disrupted collagen matrix organization, and blurred zonal boundaries—findings characteristic of advanced cartilage destruction and pannus development.
Based on the extent and consistency of cartilage damage, trypsin was selected as the optimal inducer for subsequent experiments, as it most closely mimicked the chronic degenerative joint changes observed in human OA.

Stage II: effect of SCM on aged OA model

At Stage II, 56 female rats meeting the same selection criteria as in Stage I were included in the study. All animals were divided into six groups of 10 animals each, including six intact rats (Figure 2).

Throughout the 60-day experiment, a detailed observation diary was maintained, recording the animals' body weight, motor activity, and periodic blood sampling for biochemical and immunological analyses.

Clinical observations and weight dynamics

Body weight remained stable in Groups I and II but declined significantly in Groups III and V (P < 0.05), correlating with behavioral signs of OA. Groups IV and VI, which received SCM, demonstrated partial preservation of body weight and motor activity (Supplementary Figure 6).

Biochemical Parameters

ALT and AST levels were highest in Group III, reaching 92.98 ± 4.3 U/L and 366.15 ± 18.7 U/L, respectively. SCM treatment in Groups IV and VI significantly reduced these enzyme levels (P < 0.01), suggesting hepatoprotective and anti-inflammatory effects. ALP levels, which reflect bone metabolism, were significantly lower in SCM-treated groups compared to untreated OA animals (P < 0.05).
Trypsin-induced OA significantly altered serum biochemical markers, including ALP, ALT, and AST. The elevation of these enzymes reflects activation of inflammatory pathways and a secretory phenotype in senescent cells, associated with increased cytokine and protease production.
A gradual increase in ALP was observed in the control group (Group I) over time, consistent with age-related changes. However, in SCM-treated groups (Groups II, IV, and VI), ALP levels were significantly lower (P < 0.05), indicating a potential slowdown in both aging processes and OA progression. A similar reduction in ALP was also seen in Group III (OA + estrogen receptor blockade) (Supplementary Figure 7).
ALT and AST elevations in Group III (reaching 92.98 ± 4.3 U/L and 366.15 ± 18.7 U/L, respectively) were significantly reduced following SCM treatment in Groups IV and VI (P < 0.01) (Supplementary Figures 8 & 9). A correlation was observed between ALP and ALT levels, reflecting their shared roles in bone and liver metabolism. An increase in ALT (indicating inflammation and cellular damage) was accompanied by a reduction in ALP compared to the control group, suggesting impaired bone metabolism.
Notably, the group receiving chondroitin sulfate (Group V) did not exhibit significant improvements in any of the biochemical parameters, underscoring the need for new therapeutic strategies, particularly for postmenopausal OA.

Cytokine profile

SCM-treated groups (Groups IV and VI) showed reduced levels of IL-1β, IL-6, TNF-α, and IL-8 compared to Group III (P < 0.01), indicating suppression of pro-inflammatory signaling. CRP levels also declined in these groups, consistent with a reduction in systemic inflammation (Supplementary Figure 10).

Histological assessment

Various staining methods were employed to assess cartilage integrity, including quantitative evaluation of chondrocytes and nuclear-cytoplasmic ratios. The obtained data were analyzed using the international Mankin scoring system and processed statistically. A comprehensive assessment of joint condition included key markers of chronic osteoarthritis, such as cartilage degeneration, pannus formation, joint space narrowing, synovial membrane alterations, bone tissue changes, and inflammation severity (Figure 3).

Histological analysis was conducted to evaluate the extent of cartilage degeneration and regenerative changes across all experimental groups. Group I (Intact control) displayed well-preserved cartilage with a smooth articular surface, normal zonal architecture, and evenly distributed chondrocytes at various stages of maturation. The cartilage matrix was dense, with no signs of erosion or pannus formation. The Mankin score was 1, indicating minimal histological alterations. Group II (Healthy + SCM) showed a slight increase in chondrocyte clustering and mucopolysaccharide content, as evidenced by intense matrix staining. Isogenic groups were aligned along a uniform trajectory within cartilage of consistent thickness. These findings suggest early anabolic activity without evident degeneration, with a Mankin score of 2. Group III (iOA + Postmenopause) exhibited the most severe histopathological changes. The cartilage surface was irregular with deep erosions and substantial chondrocyte loss. Cystic resorption cavities filled with adipose tissue were observed, and the bone trabeculae appeared thin and fragmented. Osteoclast activity was apparent, indicating advanced joint degeneration. The Mankin score was 12. Group IV (iOA + Postmenopause + SCM) demonstrated marked improvement compared to Group III. The cartilage surface was more regular, with evidence of chondrocyte proliferation and moderate vacuolar dystrophy. Isogenic clusters were organized, and the extracellular matrix was moderately preserved. The Mankin score was 4. Group V (iOA + Postmenopause + comparison drug): Showed intermediate cartilage damage. While the articular surface appeared smooth, PAS staining revealed a reduction in Schiff-positive matrix components, indicating partial degradation. Chondrocyte density was lower than in SCM-treated groups. Mankin score = 7. Group VI (iOA + SCM): Showed clear signs of regeneration. Cartilage layers were uniform in thickness, and large isogenic groups of chondrocytes were observed. The matrix was rich in mucopolysaccharides, suggesting enhanced biosynthetic activity. Mankin score = 6.

These histological findings confirm that SCM treatment supports chondrocyte activity and matrix repair in osteoarthritic cartilage, with particularly favorable outcomes in postmenopausal animals receiving SCM compared to untreated controls. To enable comparative evaluation across groups, morphometric analysis and the Mankin scale were applied [26, 28]. A quantitative assessment of cartilage and epiphyseal zones in the tibia and femur was performed (Figure 4).

Notably, Group VI demonstrated significant thickening of the tibial epiphyseal zone, confirming a robust regenerative effect of SCM on OA-damaged cartilage.
To assess chondrocyte maturation, the nuclear-cytoplasmic index (NCI) was calculated in several mitotically active zones of the cartilage (Figure 5).

NCI values were significantly higher in Group I and moderately elevated in SCM-treated Groups II, IV, and VI (P < 0.05 vs. Group III), indicating reactivation of chondrocyte metabolism without evidence of hyperproliferation.
Staining methods enabled the identification of cartilage damage and impaired metabolic function in OA groups, as well as restoration of metabolic activity in chondrocytes after SCM administration (Supplementary Figure 11). Group III (OA + postmenopause) showed severe cartilage degradation, including erosions, thinning, and indistinct zonal boundaries. In contrast, Groups IV and VI, which received SCM, exhibited significant histological improvement, indicating reactivation of chondrocyte metabolism.
Quantitative histological scoring ranged from 0 to 3 per parameter, with a total score from 0 (healthy tissue) to 12 (maximum cartilage damage), as summarized in Supplementary Table 3.
In our study, the lowest score (1) was observed in control Group I, while the highest (12) was recorded in untreated Group III (OA + postmenopause). Cartilage condition in Group II was slightly worse than Group I, while Group IV showed significant improvement over Group III, and Group VI demonstrated better results than Group V.
A comparative analysis across SCM-treated groups—II (Healthy + SCM), IV (OA + Postmenopause + SCM), and VI (OA + SCM)—revealed a 2:4:6 scoring pattern. This suggests a complex and possibly modulatory role of estrogen receptor blockade in postmenopausal OA (Supplementary Figure 12 and Supplementary Table 3).
Together, these findings confirm the regenerative and anti-inflammatory efficacy of SCMs in a rat model of age-related osteoarthritis.

Acknowledgments

Baratov Kuzizhon Rabbim ugli (PhD) from the Laboratory of Pharmacology and Screening of Biologically Active Substances of the Institute of Biological Chemistry of the Academy of Sciences of the Republic of Uzbekistan for assistance in working with animals; Alloberganov Dilshod Shavkatovich (PhD), Associate Professor of the Republican Center for Pathological Anatomy (Tashkent) for assistance in histological evaluation of biomaterial; Grigoryants Karina Eduardovna from the Department of Cell Therapy of the Institute of Human Immunology and Genomics (Tashkent) for assistance in photographing and videotaping experiments; Ryskulov Farukh Tursunbaevich, Department of Cell Therapy of the Institute of Human Immunology and Genomics (Tashkent)for assistance in conducting laboratory work.

Authors contribution

Saodat Muratkhodjaeva performed all experimental work, mathematical data processing and graphs, and made significant contributions to the writing of the manuscript. Tamara Aripova was responsible for the overall supervision of the experiments and writing the manuscript. Javdat Muratkhodjaev was responsible for the overall idea and writing of this manuscript. All authors read and approved the final version of the manuscript and take accountability for all aspects of the work, ensuring that questions related to the accuracy or integrity of any part of the work are appropriately investigated and resolved.

Availability of data and materials

Not applicable.

Financial support and sponsorship

None.

Conflicts of interest

All authors declared that there are no conflicts of interest.

Ethical approval and consent to participate

All animal studies were conducted in accordance with the recommendations of the Ethics Committee of Uzbekistan (2014) and the Ethical Guidelines for the Use of Animals in Scientific Research (2019). The Ethics Committee's permission to work with laboratory animals and the completed ARRIVE checklist are attached.

Consent for publication

The authors declare that they have no competing interest.

1. Yang J, Hayano M, Griffin P, Amorim J, Bonkowski M, Apostolides J, et al. Loss of epigenetic information as a cause of mammalian aging. Cell, 2023, 186(2): 305-326.e327. [Crossref]

2. López-Otín C, Blasco M, Partridge L, Serrano M, & Kroemer G. The hallmarks of aging. Cell, 2013, 153(6): 1194-1217. [Crossref]

3. Pal S, & Tyler J. Epigenetics and aging. Sci Adv, 2016, 2(7): e1600584. [Crossref]

4. Lu Y, Tian X, & Sinclair D. The information theory of aging. Nature Aging, 2023, 3(12): 1486-1499. [Crossref]

5. Guo J, Huang X, Dou L, Yan M, Shen T, Tang W, et al. Aging and aging-related diseases: from molecular mechanisms to interventions and treatments. Signal Transduct Target Ther, 2022, 7(1): 391-405. [Crossref]

6. Diekman B, & Loeser R. Aging and the emerging role of cellular senescence in osteoarthritis. Osteoarthritis Cartilage, 2024, 32(4): 365-371. [Crossref]

7. Loeser R. Aging and osteoarthritis: the role of chondrocyte senescence and aging changes in the cartilage matrix. Osteoarthritis Cartilage, 2009, 17(8): 971-979. [Crossref]

8. Croft A, Campos J, Jansen K, Turner J, Marshall J, Attar M, et al. Distinct fibroblast subsets drive inflammation and damage in arthritis. Nature, 2019, 570(7760): 246-251. [Crossref]

9. Long H, Liu Q, Yin H, Wang K, Diao N, Zhang Y, et al. Prevalence trends of site-specific osteoarthritis from 1990 to 2019: findings from the global burden of disease study 2019. Arthritis Rheumatol, 2022, 74(7): 1172-1183. [Crossref]

10. Global, regional, and national burden of osteoarthritis, 1990-2020 and projections to 2050: a systematic analysis for the global burden of disease study 2021. Lancet Rheumatol, 2023, 5(9): e508-e522. [Crossref]

11. Ushiyama T, Ueyama H, Inoue K, Ohkubo I, & Hukuda S. Expression of genes for estrogen receptors alpha and beta in human articular chondrocytes. Osteoarthritis Cartilage, 1999, 7(6): 560-566. [Crossref]

12. Nüesch E, Dieppe P, Reichenbach S, Williams S, Iff S, & Jüni P. All cause and disease specific mortality in patients with knee or hip osteoarthritis: population based cohort study. Bmj, 2011, 342: d1165. [Crossref]

13. Wehling P, Evans C, Wehling J, & Maixner W. Effectiveness of intra-articular therapies in osteoarthritis: a literature review. Ther Adv Musculoskelet Dis, 2017, 9(8): 183-196. [Crossref]

14. Berenbaum F. Annals of the Rheumatic Diseases collection on osteoarthritis (2018-2023): hopes and disappointments. Ann Rheum Dis, 2024, 83(2): 133-135. [Crossref]

15. Ball H, Alejo A, Kronk T, Alejo A, & Safadi F. Epigenetic regulation of chondrocytes and subchondral bone in osteoarthritis. Life (Basel), 2022, 12(4)： 582-593. [Crossref]

16. Yang J, Petty C, Dixon-McDougall T, Lopez M, Tyshkovskiy A, Maybury-Lewis S, et al. Chemically induced reprogramming to reverse cellular aging. Aging (Albany NY), 2023, 15(13): 5966-5989. [Crossref]

17. Pereira B, Correia F, Alves I, Costa M, Gameiro M, Martins A, et al. Epigenetic reprogramming as a key to reverse ageing and increase longevity. Ageing research reviews, 2024, 95: 102204. [Crossref]

18. Knyazer A, Bunu G, Toren D, Mracica T, Segev Y, Wolfson M, et al. Small molecules for cell reprogramming: a systems biology analysis. Aging, 2021, 13(24): 25739-25762. [Crossref]

19. Nie B, Nie T, Hui X, Gu P, Mao L, Li K, et al. Brown adipogenic reprogramming induced by a small molecule. Cell Rep, 2017, 18(3): 624-635. [Crossref]

20. Kalashnikova S, & V.V N. Estrogens and modulation of the hypothalamic-pituitary-thyroid axis during chronic endogenous intoxication in rats. Bulletin of the Volgograd Scientific Center of the Russian Academy of Medical Sciences, 2009, 1(21): 49-53.

21. Corciulo C, Scheffler J, Gustafsson K, Drevinge C, Humeniuk P, Del Carpio Pons A, et al. Pulsed administration for physiological estrogen replacement in mice. F1000Res, 2021, 10: 809-812. [Crossref]

22. Ge Y, Zhou S, Li Y, Wang Z, Chen S, Xia T, et al. Estrogen prevents articular cartilage destruction in a mouse model of AMPK deficiency via ERK-mTOR pathway. Ann Transl Med, 2019, 7(14): 336-345. [Crossref]

23. Horkeby K, Farman H, Movérare-Skrtic S, Lionikaite V, Wu J, Henning P, et al. Phosphorylation of S122 in ERα is important for the skeletal response to estrogen treatment in male mice. Sci Rep, 2022, 12(1): 22449. [Crossref]

24. Gyarmati J, Földes I, Kern M, & Kiss I. Morphological studies on the articular cartilage of old rats. Acta Morphol Hung, 1987, 35(3-4): 111-124.

25. Mankin H, Dorfman H, Lippiello L, & Zarins A. Biochemical and metabolic abnormalities in articular cartilage from osteo-arthritic human hips. II. Correlation of morphology with biochemical and metabolic data. J Bone Joint Surg Am, 1971, 53(3): 523-537.

26. Moody H, Heard B, Frank C, Shrive N, & Oloyede A. Investigating the potential value of individual parameters of histological grading systems in a sheep model of cartilage damage: the Modified Mankin method. J Anat, 2012, 221(1): 47-54. [Crossref]

27. Kalbhen D. Chemical model of osteoarthritis--a pharmacological evaluation. J Rheumatol, 1987, 14 Spec No: 130-131.

28. Yeh T, Wen Z, Lee H, Lee C, Yang Z, Jean Y, et al. Intra-articular injection of collagenase induced experimental osteoarthritis of the lumbar facet joint in rats. Eur Spine J, 2008, 17(5): 734-742. [Crossref]

29. Salo P, Hogervorst T, Seerattan R, Rucker D, & Bray R. Selective joint denervation promotes knee osteoarthritis in the aging rat. J Orthop Res, 2002, 20(6): 1256-1264. [Crossref]

30. Muzhikyan A, Shekunova E, Kashkin V, Makarova M, & Makarov V. Histological evaluation of joint pathology in various models of chronic arthritis in rats. Laboratornye Zhivotnye dlya nauchnych issledovanii (Laboratory Animals for Science), 2018, 1: 04. [Crossref]

31. Korochina K, Chernysheva T, Polyakova VS, & Korochina I. Features of morphology of articular cartilage in patients with different phenotypes of knee osteoarthritis. Arkhiv patologii, 2020, 82: 13-25. [Crossref]

32. Perepelkina D, Nasibova S, Bakhtin V, & N.V I (2019). Histological technique for preparing articular cartilage preparations from laboratory animals. Proceedings of V International (75th All-Russian) Scientific and Practical Conference "Topical Issues of Modern Medical Science and Healthcare.

33. Sanchez-Lopez E, Coras R, Torres A, Lane N, & Guma M. Synovial inflammation in osteoarthritis progression. Nat Rev Rheumatol, 2022, 18(5): 258-275. [Crossref]

34. Loeser R, Collins J, & Diekman B. Ageing and the pathogenesis of osteoarthritis. Nat Rev Rheumatol, 2016, 12(7): 412-420. [Crossref]

35. Cuollo L, Antonangeli F, Santoni A, & Soriani A. The senescence-associated secretory phenotype (SASP) in the challenging future of cancer therapy and age-related diseases. Biology, 2020, 9(12): 485-496. [Crossref]

36. Zhu X, Chen Z, Shen W, Huang G, Sedivy J, Wang H, et al. Inflammation, epigenetics, and metabolism converge to cell senescence and ageing: the regulation and intervention. Signal Transduct Target Ther, 2021, 6(1): 245-255. [Crossref]

37. Pritzker K, Gay S, Jimenez S, Ostergaard K, Pelletier J, Revell P, et al. Osteoarthritis cartilage histopathology: grading and staging. Osteoarthritis Cartilage, 2006, 14(1): 13-29. [Crossref]

38. Zhang X, Xiang S, Zhang Y, Liu S, Lei G, Hines S, et al. In vitro study to identify ligand-independent function of estrogen receptor-α in suppressing DNA damage-induced chondrocyte senescence. Faseb j, 2023, 37(2): e22746. [Crossref]

39. Wang N, Zhang X, Rothrauff B, Fritch M, Chang A, He Y, et al. Novel role of estrogen receptor-α on regulating chondrocyte phenotype and response to mechanical loading. Osteoarthritis Cartilage, 2022, 30(2): 302-314. [Crossref]