Skip to main content
NCBI home page
Scientific Reports logoLink to Scientific Reports
. 2025 Nov 24;15:41582. doi: 10.1038/s41598-025-25432-3

Investigating inflammation of adipose tissue in a diet induced obesity model of OA in female sprague dawley rats

Nada Abughazaleh 1,2,3,✉, Hannah Smith 1,2, Hijab Chaudhri 4, Ruth-Anne Seerattan 5, David A Hart 2,5,6, Raylene A Reimer 5,7, Walter Herzog 1,2,3,5,6
PMCID: PMC12644849  PMID: 41285874

Abstract

This study was aimed at evaluating the disturbance of mesenteric fat, and the infrapatellar fat pad (IFP) of the knee, and their potential contribution to systemic inflammation and local inflammation in the knee, and the development of knee joint OA in female Sprague–Dawley rats.Female rats were randomized into two groups; a Chow-fed group (Chow) and rats fed a high-fat and high-sucrose diet (HFS). Body composition, serum lipid profile, insulin resistance, serum and synovial fluid inflammatory markers, knee joint degeneration, and adipocyte size in the two fat depots were assessed after 12 weeks of the diet interventions. Mesenteric fat macrophage infiltration and infrapatellar fat pad fibrosis were also quantified.Rats from the HFS group had higher body fat, serum triglycerides levels, serum cholesterol levels, insulin resistance, increases in selected serum biomarkers such as TNF-α, leptin, MCP-1, and MIP-1α, and more severe cartilage degeneration and subchondral bone lesions than did the Chow-fed control rats. However, no increase in macrophage infiltration and fibrosis in the fat tissues were observed in the HFS group compared to the chow-fed group.The mesenteric fat and IFP were protected from inflammation associated with the HFS diet in the female rats, indicating a sex-based difference in the response to the HFS diet. However, cartilage degeneration and subchondral bone lesions were increased in the HFS compared to the control group rats, suggesting that the mechanisms resulting in knee OA development and progression in female rats may distinctly differ from that of male rats.

Supplementary Information

The online version contains supplementary material available at 10.1038/s41598-025-25432-3.

Keywords: Female sprague–dawley rats, Mesenteric fat, Infrapatellar fat pad, Osteoarthritis, Diet-induced obesity, Sex differences

Subject terms: Diseases, Rheumatic diseases, Osteoarthritis

Introduction

Osteoarthritis (OA) is a degenerative joint disorder characterized by the degradation of cartilage and the formation of osteophytes, and in humans is prevalent in the hips, knees, feet and hands1. Clinical signs of OA include joint stiffness, swelling, pain, redness, and limitation of joint range of motion2. OA has often been classified as a ‘wear and tear’ disease1. However, recent studies identified the multifactorial nature of OA, including an association of OA with obesity and inflammatory processes3–5. Obesity and associated metabolic syndrome are now considered among the most important risk factors for the onset and progression of knee OA6. Reyes et al. reported that the incidence of knee OA is more than five times greater in subjects with grade II obesity compared to corresponding adults of normal weight7. Moreover, increased body fat has been positively associated with knee OA in premenopausal women8. Despite increasing evidence for obesity as a risk factor for knee joint OA, the potential mechanisms underlying the relationship between obesity and OA are not fully understood. Previous studies suggested that low-level systemic inflammation and local inflammation in the knee associated with obesity may play a crucial role in OA onset and progression9–11.

In people with obesity, white adipose tissue (AT) expands and adipocytes increase in size (hypertrophy) and/or increase in number (hyperplasia)12. Also, adipocyte turnover rates become elevated13 leading to cell hypoxia and adipocyte death14. The dead adipocytes need to be cleared by macrophages, thereby increasing macrophage and immune cell infiltration into the fat tissue15,16. This infiltration has been thought to be particularly pronounced in visceral AT8. Infiltrated macrophages aggregate around the dead adipocytes forming crown-like structures (CLSs)15, which have become a hallmark of the inflammatory state of adipose tissues in obesity.

The inflamed adipose tissue acts like an endocrine organ producing and then secreting proinflammatory cytokines and adipokines into the bloodstream, which then exert potentially damaging effects on many tissues, including muscles and joints17. Several adipokines have been implicated with the onset and progression of OA, including leptin, adiponectin, resistin, IL-6, and TNF-α18. In the knee joint, the infrapatellar fat pad (IFP) is a major source of adipokines that can contribute to synovial inflammation, matrix metalloproteinase (MMP) production, and cartilage degeneration19. The expansion of adipocytes has been shown to increase inflammatory secretions which can contribute to fibrosis of the adipose tissue20. Fibrosis of the IFP can disrupt its function in cushioning the joint and may contribute to limiting joint mobility21.

Diet-induced obesity (DIO) has been used as a powerful preclinical model to study OA in the context of obesity and metabolic syndrome. Collins et al. were among the first to use a high fat/high sucrose (HFS) diet to induce obesity in male Sprague–Dawley rats, and they observed OA-like changes in the knee joints following 12 and 28 weeks of HFS exposure3. Although knee OA has a higher prevalence and severity in women than men2, most preclinical studies exploring risk factors and treatments for OA have focused on male animals, likely due to their comparatively stable hormonal profiles providing a uniform study population22. However, it has been observed that there are differences in systemic inflammation, obesity development, and glucose tolerance between male and female rats23. Therefore, outcomes from male studies on diet-induced obesity should not be generalized to female models without accounting for potential sex-specific differences.

The aims of the present study were to: (i) determine the development of knee joint OA in female Sprague–Dawley rats in a DIO model, and (ii) evaluate the disturbance of two fat depots (mesenteric fat and infrapatellar fat pad) and the development of local and systemic inflammation. We hypothesized that; (i) female rats fed a HFS diet for 12 weeks will have greater knee joint degeneration, greater systemic and local (knee joint) inflammation, and greater metabolic disturbance compared to chow-fed rats, and (ii) that the IFP and the mesenteric fat depots will exhibit macrophage infiltration and fibrosis contributing to systemic and local inflammation. This study forms part of a larger research project, and some of the data presented herein have been utilized in a previously published paper24.

Results

Body composition and body mass

At the end of the 12 week experimental period, animals in the HFS group were heavier, had a higher percentage of body fat, bone mass, and a lower percentage of lean body mass compared to the Chow group control animals (p < 0.001, p = 0.001, p < 0.001 and p < 0.001 respectively, Fig. 1).

Fig. 1.

Fig. 1

Open in a new tab

(A) Body mass (g). (B) Bone mass (g). (C) Body composition presented by the percent of lean and fat mass. * indicates a significant difference (p < 0.05) compared to the HFS group. Chow: female rats fed a chow diet; HFS: female rats fed a HFS diet. Data are presented as mean ± standard deviation.

Serum lipid profile

Serum total cholesterol, triglyceride levels, and HDL levels were significantly higher in HFS group rats compared to the Chow group control rats (p = 0.013, p = 0.001, and p = 0.002, respectively, Table 1).

Table 1.

Serum lipid profile, with total cholesterol, triglyceride and HDL levels for Chow and HFS group rats in mmol/L.

Female groups Chow HFS
Mean ± SD Median (Range) Mean ± SD Median (Range)
Total cholesterol (mmol/L) 107 ± 0.27 1.7–0.89 2.2 ± 0.5* 2.03–1.76
Triglyceride (mmol/L) 0.52 ± 0.23 0.5–0.8 1.26 ± 0.68* 1.13–2.5
HDL (mmol/L) 1.1 ± 0.2 1.2–0.65 1.69 ± 0.43* 1.59–1.37
Open in a new tab

*Denotes a significant difference.

Adipocyte size in mesenteric fat and IFP

Mesenteric fat from the HFS group rats had fewer small cells in the first five cell area ranges (< 500 µm2, 500–999 µm2, 1000–1499 µm2, 1500–1999 µm2, and 2000–2499 µm2) compared to Chow group rats (p = 0.001, p = 0.001, p = 0.001, p = 0.001, and p = 0.029, respectively, Fig. 2A), and more large cells in area bins of 3500–3999 µm2, 4000–4499 µm2, 4500–4999 µm2, and > 5000 µm2 compared to Chow group rats (p = 0.00, p = 0.00, p = 0.00, and p = 0.029, respectively, Fig. 2A). Adipocytes from the IFP in the HFS rats had fewer small cells in areas ranging less than 250 µm2, 250–499 µm2, and 500–749 µm2 (p = 0.001, p = 0.001, and p = 0.015, respectively) and more large cells in areas ranging from 750 to 999 µm2, 1000–1249 µm2, 1250–1500 µm2, 1500–1999 µm2, and more than 2000 µm2 (p = 0.028, p = 0.001, p = 0.001, p = 0.001, and p = 0.001, respectively, Fig. 2B) compared to IFP adipocytes in the Chow group rats. Figure 3 illustrates the difference in adipocyte size within the mesenteric fat between the two groups.

Fig. 2.

Fig. 2

Open in a new tab

(A) Relative frequency (%) of adipocyte size (µm2) presented as the mean and standard error in the Chow and HFS groups for the mesenteric fat. (B) Relative frequency (%) of adipocyte size (µm2) presented as the mean and standard error in the Chow and HFS groups for the infrapatellar fat. * Indicates a significant difference (p < 0.05) compared to the HFS group. Chow: female rats fed a chow diet; HFS: female rats fed a HFS diet.

Fig. 3.

Fig. 3

Open in a new tab

Adipocyte size in mesenteric fat. (A) an example of adipocyte cell size in chow fed rats. (B) an example of adipocyte cell size in HFS fed rats. Note the increased cell size in the HFS compared to the chow fed control group rats.

Macrophage infiltration of mesenteric fat

There were no detectable differences in macrophage infiltration into the mesenteric fat between the HFS and the Chow group rats Fig. 4A. Supplementary Figure S1 shows immunofluorescent stained adipocytes and an exemplar crown-like structure surrounding a single adipocyte.

Fig. 4.

Fig. 4

Open in a new tab

(A) Macrophage infiltration measured as the number of crown-like structures per frame. (B) Fibrosis of the infrapatellar fat pad, measured as the percent area of collagen relative to the total area. Chow: female rats fed a chow diet; HFS: female rats fed a HFS diet. * Indicates a significant difference (p < 0.05) compared to the HFS group.

IFP fibrosis

The collagen content of the IFP from the HFS group rats was lower than that in the Chow group rats (p = 0.021) Fig. 4B.

Insulin sensitivity

Whole-body insulin resistance, quantified using the HOMA-IR index, was increased in HFS compared to Chow group rats (p = 0.001) (Fig. 5).

Fig. 5.

Fig. 5

Open in a new tab

Insulin sensitivity values indicated by the Homeostatic Model Assessment for Insulin Resistance (HOMA-IR) index for the Chow and HFS groups. * indicates a significant difference (p < 0.05) compared to the HFS group. Chow: female rats fed a chow diet; HFS: female rats fed a HFS diet.

Blood and synovial fluid cytokines and adipokines

HFS group rats compared to Chow group control rats had significantly increased serum levels for Eotaxin, Fractalkine, G-CSF, IFNγ, IL-1α, IL-2, IL-4, IL-5, IL-12 p70, IL-13, IL-17A, IL-18, Leptin, MCP-1, MIP-1α, TNFα, and VEGF (p = 0.012, p = 0.001, p = 0.025, p = 0.025, p = 0.005, p = 0.001, p = 0.001, p = 0.001, p = 0.002, p = 0.005, p = 0.002, p = 0.001, p = 0.001, p = 0.016, p = 0.001, p = 0.004, and p = 0.001, respectively. Table 2). HFS compared to Chow control group rats also had increased synovial fluid IL-1α, Leptin, and IP-10/CXCL10 levels (p = 0.003, p = 0.011, and p = 0.003, respectively. Table 3).

Table 2.

Mean (pg/mL) ± SD and median (range) values for blood cytokines and adipokines for chow and HFS group rats.

Chow HFS P-values
Mean (pg/mL) ± SD Median (Range) Mean (pg/mL) ± SD Median (Range)
Eotaxin 7.19 ± 1.8 6.41 (4.9) 10.4 ± 4.52 8.8 (16) 0.012*
Fractalkine 41.68 ± 15.6 39.66 (39.15) 96.4 ± 36.2 83.4 (106) 0.000*
G-CSF 12.2 ± 6.7 9.76 (21.3) 36.58 ± 44.7 24.2 (160.9) 0.025*
IFNy 74.5 ± 30.98 66.36 (101.12) 173.3 ± 134.7 106.4 (459.49) 0.025*
IL-la 26.46 ± 11.69 20.29 (32.48) 75.37 ± 59.98 63.0 (217.11) 0.005*
IL-lp 106.89 ± 49.55 120.89 (130.83) 300.08 ± 299.3 192.5 (1075.89) 0.135
IL-2 79.61 ± 52.6 67.64 (152.65) 334.17 ± 207.85 282.24 (714.82) 0.000*
IL-4 29.04 ± 17.08 21.6 (48.47) 87.48 ± 34.02 88.76 (125.2) 0.000*
II-5 51.36 ± 23.56 46.97 (75.55) 125.98 ± 38.67 122.28 (116.55) 0.000*
IL-6 673.06 ± 420.07 636.96 (1058.74) 1093.15 ± 708.22 902.36 (2454.84) 0.082
IL-10 138.26 ± 75.08 171.89 (176.72) 272.72 ± 182.3 236.97 (586.55) 0.427
IL-12p70 139.09 ± 123.1 109.37 (360.61) 399.79 ± 198.37 437.95 (611.52) 0.002*
IL-13 20.91 ± 8.80 17.53 (23.32) 35.49 ± 17.21 31.84 (60.53) 0.005*
IL-17A 15.43 ± 11.66 14.91 (34.17) 53.71 ± 51.47 44.43 (189.26) 0.002*
IL-18 99.59 ± 61.27 80.25 (166.85) 401.9 ± 251.25 289.81 (708.74) 0.000*
IP-10 204.3 ± 67.95 179.98 (179.05) 252.07 ± 105.85 253.31 (378.86) 0.678
Leptin 4899.76 ± 2396.73 5014.89 (6663.88) 15,740.72 ± 5357.99 14,257.43 (19,769.02) 0.000*
LIX 2920.99 ± 1006.81 2957.33 (3174.21) 3023.09 ± 709.79 3199.93 (2426.33) 0.91
MCP-1 334.81 ± 123.32 312.81 (317.5) 576.14 ± 318.5 461.69 (1170.38) 0.016*
MlP-la 13.6 ± 5.81 12.98 (15.09) 27.03 ± 7.36 25.77 (20.75) 0.001*
RANTES 896.67 ± 398.9 895.21 (1185.96) 1007.96 ± 609.99 850.22 (2164.27) 0.851
TNFa 1.5 ± 1.54 0.68 (3.93) 6.68 ± 6.08 5.1 (21.67) 0.004*
VEGF 66.26 ± 22.11 64.98 (62.85) 113.67 ± 34.42 104.68 (126.74) 0.001*
Open in a new tab

*Denotes a significant difference.

Table 3.

Synovial fluid cytokines and adipokines analysis. Data are shown as means ± 1 standard deviation and as medians and ranges.

Chow HFS
Mean (pg/mL) ± SD Median (Range) Mean (pg/mL) ± SD Median (Range) P-values
Eotaxin 27.5 ± 17.9 27.5 (25.3) 41.4 ± 16.8 34.5 (39.8) 0.73
Fractalkine 124.1 ± 59.0 115.1 (183.4) 50.4 ± 74.4 68.39 (270.3) 0.228
G-CSF 4329.7 ± 1646.2 4889.4 (5661.6) 5190.9 ± 1928.7 4965.3 (6251.9) 0.771
IFNγ 1347.9 ± 253.7 1291.4 (756.1) 1493.6 ± 629.5 1682.26 (1677.8) 0.974
IL-1α 142.6 ± 58.9 162.6 (196.4) 440.6 ± 407.9 326.30 (1361.5) .003*
IL-1β 165.4 ± 55.4 147.6 (191.6) 188.4 ± 83.3 136.22 (254.4) 0.539
IL-2 79.5 ± 59.5 50.5 (144.1) 134.9 ± 54.6 122.6 (144.8) 0.177
IL-4 54.3 ± 33.3 59.4 (111.8) 85.9 ± 80.7 68.74 (291.9) 0.346
IL-5 67.1 ± 24.8 67.1 (35.1) 139.1 ± 53.5 155.2 (158.4) 0.181
IL-6 1043.7 ± 751.2 595.2 (2309.7) 1434.7 ± 779.9 1748.49 (2312.8) 0.203
IL-10 68.6 ± 40.2 81.4 (124.9) 107.4 ± 44.9 81.43 (124.9) 0.314
IL-12p70 69.1 ± 88.9 69.1 (125.7) 53.3 ± 47.7 53.2 (125.7) 0.529
IL-13 70.3 ± 40.7 64.6 (139.2) 67.9 ± 29.3 64.15 (97.5) 0.862
IL-18 20,220.7 ± 4338.5 21,745.9 (14,606.5) 19,699.3 ± 5369.2 18,530. (19,406.9) 0.628
IP-10 80.7 ± 65.4 58.5 (207.2) 142.9 ± 98.2 98.1 (310.9) .003*
Leptin 4193.7 ± 781.8 4271.9 (2739.9) 11,462.5 ± 7890.9 10,029.5(26,092.1) .011*
LIX 90.9 ± 119.8 90.9 (169) 196.9 ± 85.7 171.4 (232.7) 0.867
MCP-1 1669.5 ± 490.7 1875.0 (1585.2) 2571.9 ± 1277.9 2404.66 (4510.0) 0.059
RANTES 98.1 ± 48.9 104.7 (150.9) 96.9 ± 63.6 77.58 (214.6) 0.314
TNFα 90.8 ± 24.4 90.6(76.6) 104.5 ± 24.8 100.95 (82.9) 0.314
VEGF 205.6 ± 102.2 200.8(325.5) 263.8 ± 134.4 276.04 (483.8) 0.254
GRO/KC/CINC-1 5540.2 ± 2829.6 5766.0 (8211.7) 7625.2 ± 4408.9 8768.0 (12,356.4) 0.349
Open in a new tab

*Indicates a significant difference (p < 0.05) compared to the Chow and HFS female group rats. Chow: rats fed chow diet; HFS: rats fed a high fat/high sucrose diet.

Knee joint degeneration

HFS group rats had greater knee cartilage degeneration (p = 0.016, Fig. 6A), bone lesions (p = 0.00, Fig. 6B), and total knee joint OA scores than did the Chow group rats (p = 0.00, Fig. 6C). Figure 7 represents typical and non typical lesions seen in the tibia and femur of femal rats.

Fig. 6.

Fig. 6

Open in a new tab

Knee joint degeneration represented by OA score Chow and HFS. (A): Typical joint score. (B): Non-typical joint score. C: Total OA scores. * indicates a significant difference (p < 0.05) compared to the Chow group. Chow: female rats fed a chow diet; HFS: female rats fed a HFS diet.

Fig. 7.

Fig. 7

Open in a new tab

(A) Representative joint showing Tibial plateau and Femoral condyle of a female rat fed a chow diet (B): Representative illustration of a typical lesion seen in the tibial cartilage of rat fed the HFS diet. The black arrows show a cartilage cyst and sub- surface fibrillation of the cartilage. (C): Representative illustration of a non-typical lesion in the tibial plateau. The dashed line shows the collapse of cartilage into the subchondral bone region. (D): Representative illustration of a non-typical lesion in the femoral condyle. The dashed line shows local thickening of proteoglycan-rich cells below the tidemark.

Relationship between knee joint degeneration and other outcome variables

The cartilage OA score and bone mass were negatively correlated in the HFS group rats (r = 0.62, p = 0.032, Fig. 8A).The bone lesion scores and leptin serum levels were positively correlated (r = 0.68, p = 0.014, Fig. 8B), and so were the fasting insulin levels and bone mass (r = 0.85, p = 0.001, Fig. 8C) in rats from the HFS group animals.

Fig. 8.

Fig. 8

Open in a new tab

(A) Relationship between typical cartilage OA scores and bone mass for Chow and HFS groups. (B) Relationship between non-typical bone lesion scores and leptin serum levels for the HFS group. (C) Relationship between bone mass and fasting insulin levels for Chow and HFS groups.

Discussion

We investigated the effects of consuming a HFS diet on mesenteric fat, the infrapatellar fat pad, metabolic disturbance, and knee joint degeneration in female Sprague–Dawley rats. The primary results indicate that female rats fed a HFS diet for 12 weeks exhibited greater knee joint degeneration with no indication of elevated mesenteric fat inflammation, IFP fibrosis, or local inflammation in the synovial fluid of the knee joint, relative to rats from the Chow group. An increase in selected serum biomarkers suggests a low level systemic inflammation in the HFS group rats consistent with markers of metabolic syndrome. Feeding female rats a HFS diet for 12 weeks also resulted in an increase in body mass and body fat percentage, dyslipidemia, and insulin resistance. These results were similar to results reported in previous studies using male Sprague–Dawley rats3,25. Thus, exposure to the HFS diet similarly led to obesity in the female rats.

In the present studies, the development of systemic inflammation, was confirmed by an increase in selected serum cytokines and adipokines, including TNF-a, leptin, MCP-1, and MIP-1α in HFS-fed rats. However, no differences were observed in IL-1β and IL-6 levels and in macrophage infiltration into the mesenteric fat between the HFS and Chow control group animals. Visceral adipose tissue has been considered to play a crucial role in metabolic disturbance associated with obesity, and the development of type 2 diabetes in humans and mice26. However, in the present diet-induced obesity model using female animals, the HFS diet did not result in detectable macrophage infiltration and fibrosis of adipose tissue which is considered a hallmark of inflammation. These findings are consistent with a previous study where obesity-prone female rats exposed to a high fat diet for 7 weeks did not experience increased expressions of Macrophage Inflammatory Protein-1 Alpha (MIP-1α ) in mesenteric fat, or increased levels of IL-10 in gonadal fat27. Lam et al. reported that female mice, consuming a high-fat diet for 8–12 weeks, did not exhibit increases in serum concentrations of IL-1β, IL-10, MCP-1, TNF-a, IL-6, and IL-12p4028. Also, female Wistar, in contrast to male Wistar, rats fed a high sucrose diet did not develop sucrose-induced insulin resistance and hypertriglyceridemia29. These results suggest that, in contrast to male rats, factors associated with fat metabolism/accumulation may protect female rats against inflammation and overt metabolic disturbances when exposed to an obesity-inducing diet. The underlying biological processes/mechanisms behind such potential protection are currently not understood. However, female mice fed a high-fat diet had an increase in the number of blood vessels in perigonadal adipose tissue, smaller adipocytes in abdominal fat, and a higher expression of adiponectin, compared to male rats30, all factors that have been reported to protect against adipose tissue hypoxia and fibrosis31. In our study, there was a significant increase in vascular endothelial growth factor (VEGF), a potent angiogenic factor, in the HFS compared to the Chow group rats, a finding which may have contributed to neovascularization of the tissues.

Furthermore, adipose tissue has been shown to respond differently to obesity depending on structure and location32. Female rats tend to store fat preferentially in subcutaneous depots, whereas male rats tend to store fat preferentially in visceral depots33. Typically, visceral adipose tissue becomes more inflamed than subcutaneous adipose tissue as indicated by a higher degree of hypertrophy and greater macrophage infiltration16. This finding may explain the attenuated systemic inflammation indicated by the increase in selected biomarkers in the compared to the male rats. Men with obesity tend to have more visceral adipose tissue than women34 and it has been thought that because of this, men have higher rates of prediabetes/diabetes, cardiovascular disease, and insulin resistance than women27. Thus, location-specific variables may contribute to the observed sex differences in response to the HFS diet.

Furthermore, we did not observe detectable biomarkers of inflammation in the visceral fat of our female rats. Therefore, the increases in serum cytokines observed in our study, and typically associated with systemic inflammation, may be related to dysbiosis of the gut microbiota following exposure to the HFS diet. Although not measured here, female rats consuming a HFS diet for 12 weeks have been shown to have an increased abundance of Clostridium, Dorea and Terrisporobacter, and a decreased abundance of Lactobacillus24 which has been proposed as obesity biomarkers35,36.

Brown adipose tissue, which is considered a lipid-burning tissue, has been reported to have a greater activity and greater mass in female compared to male rodents37,38. Specifically, female rats have larger mitochondria and higher amounts of mitochondrial proteins including mitochondrial uncoupling protein 1(UCP1) than male rats37,39. Furthermore, studies have reported that brown adipose tissue in female rats adapts more than in male rats when fed a high fat or a high fat/high sugar diet because female rats increase their energy expenditure more than male rats in the presence of an obesity-inducing diet37,39,40. Moreover, brown adipose tissue secretes various adipokines, such as adiponectin, that have an anti-inflammatory effect40. These findings suggest that brown adipose tissue may have attenuated some of the metabolic effect of the HFS diet in female rats. However, some consequences of the HFS diet were still evident so any effects will have to be confirmed in future studies.

Synovial fluid from the HFS female rats showed an increase in a few biomarkers and they had greater typical articular cartilage degeneration than the control Chow group rats (Fig. 6). These results suggest that small changes in a specific subset of inflammatory biomarkers may be sufficient to trigger the increased cartilage degeneration in the HFS compared to the control group rats observed in our study.

Leptin, a hormone released by adipose tissues, has been widely explored to explain the link between obesity and OA onset and progression41,42. In contrast to male rats undergoing the same HFS intervention as the female rats in this study5, the female rats exposed to the HFS diet had increased leptin serum and synovial fluid levels compared to the Chow control group rats. Furthermore, there was a trend of greater joint scores (more joint degeneration) with increasing levels of serum leptin levels in the HFS group rats (p = 0.087), suggesting that serum leptin levels might have indirectly affected joint integrity leading to increased joint degeneration. Future studies using both longer and shorter HFS diet intervention periods than used here, and/or using leptin deficient rodent models, may provide clarity about the potential role of leptin in the development and progression of diet-induced knee joint OA.

Interestingly, fasting insulin levels in both the HFS and the chow group rats were associated with an increase in bone mass. These findings are consistent with previous studies where insulin was thought to promote osteoblast differentiation in rats43 and HOMA-IR has been reported to be positively associated with bone mineral density in postmenopausal, non-diabetic women44. In the present study, the HFS group rats had increased bone mass that was associated with decreases in typical cartilage OA scores (less OA), indicating a potential protective effect of leptin on joint integrity. However, this finding is in contrast to previous studies performed by Hardcastle et al. who reported that a high bone mass was associated with an increased risk for radiographic hip and knee joint OA in humans45,46. Similarly, increased bone mineral density has been associated with OA of the hip and knee in humans47,48. These differences in our results compared to some reports in the literature may be due to differences between our preclinical model and OA development in humans, the age of the study participants (i.e. young animals versus menopausal females), or dietary considerations.

Interestingly, in the present study, serum leptin levels were positively correlated with increases in subchondral bone lesions in the HFS-fed rats. The female Chow control group rats did not develop detectable bone lesions. Bone lesions have been reported to occur naturally in Sprague Dawley rats49,50. However, in our female Sprague–Dawley rats, bone lesions only occurred in the HFS group rats, and the severity of these lesions was associated with increasing levels of serum leptin. The detailed mechanism(s) underlying this association remains unknown. Previous studies suggested that estrogen affects long bone development in humans and animals and high doses of estrogen promote epiphysial bone closure and cell proliferation51 Masarwi et al. showed that leptin stimulates the activity of aromatase, an enzyme involved in the conversion of androgen to estrogen, thereby reducing the proliferative capacity of chondrocytes and accelerating growth plate fusion in male Sprague Dawley rats52. Nilsson et al. reported that estrogen treatment of juvenile ovariectomized rabbits accelerated the depletion of resting zone and hypertrophic chondrocytes responsible for remodelling newly formed cartilage into bone53. These findings suggest that an interaction between leptin and sex hormones may accelerate the aging process in female rats, thereby increasing the size and number of age-related subchondral bone lesions in female Sprague–Dawley rats subjected to an obesity-inducing diet. This speculation will need to be confirmed by future investigations.

A limitation of this study is the absence of sex hormone analysis in both the HFS and control groups. While we hypothesized that brown adipose tissue may play a role in attenuating metabolic syndrome, no direct analysis was conducted to confirm this hypothesis. Additionally, the study did not include an analysis of subcutaneous fat or the changes associated with the high-fat diet (HFS), which could provide further insights into the mechanisms underlying metabolic alterations.

Conclosions

In contrast to previous results in male rats, the mesenteric fat and IPF were protected from inflammation associated with the HFS diet in the female rats, indicating a sex-based difference in the response to the HFS diet. However, there was a distinct hypertrophy of the adipocytes following exposure to the HFS diet. Despite the protection of the adipose tissues from inflammation, cartilage degeneration and subchondral bone lesions were increased in the HFS compared to the control group rats, suggesting that the mechanisms resulting in knee OA development and progression in female rats may distinctly differ from that of male rats.

Methods

Twenty-three, twelve-week-old, female Sprague–Dawley rats were randomized into two diet groups: control diet (chow, 5% of total weight as fat, 47.5% carbohydrates (only 4% from sucrose), 25% protein, 12.5% from fiber and micronutrients, and 10% moisture; Lab Diet 5001, Minneapolis, MN, United States, n = 11) and high-fat and high-sucrose diet (HFS, 20% of total weight as fat, 50% sucrose, 20% protein, and 10% fiber and micronutrients; custom Diet #102,412, Dyets, Inc., Bethlehem, PA, United States, n = 12). Animals had access to food and water ad libitum. The diet intervention lasted for 12 weeks, after which animals were euthanized at 24 weeks of age and seven outcomes were measured. All animal experiment were carried out in accordance with institutional regulations, with ethics approval provided by the Life and Environmental Sciences Animal Care Committee (Protocol #AC24-0060). Rats were obtained from the Life and Environmental Sciences Animal Resource Centre at the University of Calgary, where they were bred and housed in compliance with the guidelines set by the Canadian Council on Animal Care (CCAC): Guide to the Care and Use of Experimental Animals.The study is reported in accordance with ARRIVE guidelines. The sample size was determined based on previous studies with HFS and control rats.

Body composition

Body mass, body fat percentage, and bone mass were determined at the end of the 12 week diet intervention period using Dual Energy X-ray Absorptiometry (GE Medical Systems Lunar, Madison, WI, USA), with software for small animals.

Blood lipid profile

Prior to sacrifice, cardiac blood samples were collected following a 16-h fasting period. Serum samples were tested using a colorimetric assay for lipid profile components (total cholesterol, HDL cholesterol, and triglycerides). All blood sample analyses were performed by the Alberta Precision Laboratories (Calgary, AB, Canada).

Histology

Knee joint histology

Following the 12-week diet intervention, rats were anesthetized with 5% isoflurane, bled, and then sacrificed by severing the aorta and vena cava. Knee joints were harvested using cuts approximately 2 cm above and below the joint line, cleared of excess muscle, fixed in 10% neutral buffered formalin for four-weeks, and decalcified in Cal-X II (Fischer Scientific, Canada) for 3 weeks with daily changes. The endpoint for decalcification was determined using a 5% ammonium oxalate solution. Joints were then dehydrated through a graded series of alcohols, cleared in xylene, and infiltrated with paraffin wax, using a Leica TP1020 automated processor (Leica Biosystems, Ontario, Canada). The joints were embedded in a mix of Paraffin Plus and Paraffin Xtra wax (Fisher Scientific, Canada). Serial sagittal sections of 10 µm thickness were cut using a Leica microtome, sections were mounted onto Super-frost Plus slides (Fisher Brand) and alternate slides were stained sequentially with hematoxylin, fast green, and Safranin-O, using a Leica Autostainer XL (Leica Biosystems, Germany). Knee joint OA was assessed by two independent and blinded reviewers using a previously described histological scoring system54. The histological OA scores were a composite of three sub-scores: (i) a score assessing OA histopathological changes of the cartilage not associated with a bone lesion; (ii) a score assessing bone degeneration and the cartilage above bone lesions; and (iii) a total knee joint OA score obtained as the sum of scores (i) and (ii)54.

Infrapatellar fat pad analysis

The paraffin sections of the infrapatellar fat pad (IFP) were stained with Picro Sirius Red to determine collagen content and fat cell size distribution. Four slides from the mid region of the IFP, which were at least 200 um apart and which showed a fully intact sagittal view of the IFP, were used for analysis. Images were captured with a 10X objective on an Olympus BX53 microscope, using an Olympus DP73 camera and the Olympus CellSens Standard imaging system. Collagen content and cell size (n = 2,000 cells) were determined using Image J software (Rasband, W.S., ImageJ, U. S. National Institutes of Health, Bethesda, Maryland, USA).

Mesenteric fat

Mesenteric fat samples were collected, fixed, cut into 10 μm serial paraffin sections, and adhered to superfrost plus slides. Alternate slides were stained with hematoxylin and eosin (H&E) and imaged using the Olympus CellSens Standard imaging system. Cell sizes were determined from 2000 randomly selected cells using Image J software (Rasband, W.S., ImageJ, U. S. National Institutes of Health, Bethesda, Maryland, USA). The alternate, unstained slides were washed in Phosphate Buffered Saline (PBS), blocked with 2.5% Normal Horse serum, and macrophages were identified using CD68-specific antibodies (mouse anti-rat CD68 (BioRad), followed by ImmPRESS anti-mouse IgG Peroxidase (BioLynx. Inc), and a final Tyramide Signal Amplification system: TSA plus-CY3, Akoya Biosciences). DAPI (4′-6-diamidino-2-phenylindole) was applied to the stained sections to visualize cell nuclei. Sections were mounted with Vectashield fluorescence mounting medium (Vector Labs inc.), and images were captured with an Olympus DP28 camera, on an Olympus BX51 fluorescent microscope, using filters appropriate for CY3 and DAPI. Fluorescently-labeled profiles and DAPI images were overlayed with the aid of ImageJ software and profiles containing both CY3 and DAPI were counted to determine macrophage counts. Macrophage infiltration, represented by fluorescently labelled crown-like structures around fat cells, was quantified manually.

Glucose tolerance test

Oral glucose tolerance testing was performed on the day prior to sacrifice. Following 16 h of food deprivation, rats were given an oral gavage of 2 g/kg glucose. Blood glucose was measured with a glucose meter (OneTouch Verio and Blood Glucose Monitoring System, Lifescan, Switzerland) at 0, 15, 30, 60, and 120 min following the gavage. Blood was collected at the same time points via tail nick to measure insulin and estimate insulin resistance using the Homeostatic Model Assessment of Insulin Resistance (HOMA-IR).

Blood and synovial fluid cytokines and adipokines

Blood was drawn from the heart, allowed to clot at room temperature, subjected to centrifugation, and serum was collected. The right knee joints were opened, and synovial fluid was collected using the Whatman chromatography paper method55. Synovial fluid samples were weighed, diluted, and centrifuged. Serum and synovial fluid were analyzed for cytokines and adipokines using Rat 27® xMAP technology, which included Eotaxin, EGF, Fractalkine, IL-1α, IL-1β, IL-2, IL-4, IL-5, IL-6, IL-10, IL-12(p70), IL-13, IL-17A, IL-18, IP-10/CXCL10, GRO/KC, IFN-γ, TNF-α, G-CSF, GM-CSF, MCP-1, leptin, LIX, MIP-1α, MIP-2, RANTES, and VEGF (Eve Technologies, AB, Canada).

Statistical analysis

Mann–Whitney-U tests were used to compare knee joint scores, body composition, serum lipid profiles, glucose tolerance test, blood and synovial fluid cytokines and adipokines, and mesenteric fat and IFP cell size between the chow-fed and the HFS-fed experimental groups. Comparisons were made using IBM SPSS statistic V26 (IBM SPSS, Armonk, NY, USA). Significance was accepted for p < 0.05, two-tailed testing.

Supplementary Information

Supplementary Information. (74.7MB, docx)

Acknowledgements

This work was supported by the Arthritis Society (grant number SOG-10-052), the Canadian Institutes of Health Research FDN-143341, the Canada Research Chair Program 950-200955, the Killam Foundation The Nigg Chair for Mobility and Longevity, and the McCaig Institute for Bone and Joint Health (project number 10010760). The authors thank Andrew Sawatsky, and Timothy Leonard for technical contributions.

Author contributions

N.A. was responsible for data collection, data analysis, interpretation of data, drafting the manuscript, revising the manuscript, and approving the final submitted version of the manuscript under W.H. supervision. H.S. was responsible for animal handling, tissue collection, and revising the manuscript. H.C. was responsible for adipocyte quantification, data analysis and revising the manuscript. R.S. contributed to data analysis, interpretation of data and writing the method section. D.H. and R.R. were responsible for critically revising the manuscript and approving the final submitted version. W.H. contributed to study design, interpretation of the data, critically revising the manuscript, and approving the final submission.

Data availability

The data that support the findings of this study are available from the Human Performance lab (HPL) at the University of Calgary, The data are available from the corresponding author (Nada Abughazaleh) upon request.

Declarations

Competing interests

The authors declare no competing interests.

Footnotes

Publisher’s note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

References

  • 1.J. N. Katz, K. R. Arant, and R. F. Loeser, “Diagnosis and Treatment of Hip and Knee Osteoarthritis: A Review,” Feb. 09, 2021, American Medical Association. 10.1001/jama.2020.22171. [DOI] [PMC free article] [PubMed]
  • 2.J. E. Stevens-Lapsley and W. M. Kohrt, Osteoarthritis in women: Effects of estrogen, obesity and physical activity, (2010) 10.2217/whe.10.38. [DOI] [PubMed]
  • 3.Collins, K. H., Hart, D. A., Seerattan, R. A., Reimer, R. A. & Herzog, W. High-fat/high-sucrose diet-induced obesity results in joint-specific development of osteoarthritis-like degeneration in a rat model. Bone Joint. Res.7(4), 274–281. 10.1302/2046-3758.74.BJR-2017-0201.R2 (2018). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 4.Rios, J., Hart, D. A., Reimer, R. A. & Herzog, W. Effects of prebiotics and exercise on the progression of osteoarthritis in a rat-model of metabolic disturbance. Osteoarthritis Cartilage28, S203. 10.1016/j.joca.2020.02.330 (2020). [Google Scholar]
  • 5.Collins, K. H. H. et al. Relationship between inflammation, the gut microbiota, and metabolic osteoarthritis development: Studies in a rat model. Osteoarthritis Cartilage23(11), 1989–1998. 10.1016/j.joca.2015.03.014 (2015). [DOI] [PubMed] [Google Scholar]
  • 6.Raud, B. et al. Level of obesity is directly associated with the clinical and functional consequences of knee osteoarthritis. Sci. Rep.10.1038/s41598-020-60587-1 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 7.Reyes, C. et al. Association between overweight and obesity and risk of clinically diagnosed knee, hip, and hand osteoarthritis: A population-based cohort study. Arthritis Rheumatol.68(8), 1869–1875. 10.1002/art.39707 (2016). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 8.Nur, H. & Tuncer, T. The relationship between body composition and knee osteoarthritis in postmenopausal women. Turk. J. Phys. Med. Rehabil.64(2), 121–125. 10.5606/tftrd.2018.1496 (2018). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 9.Collins, K. H., Hart, D. A., Reimer, R. A., Seerattan, R. A. & Herzog, W. Response to diet-induced obesity produces time-dependent induction and progression of metabolic osteoarthritis in rat knees. J. Orthop. Res.34(6), 1010–1018. 10.1002/jor.23103 (2016). [DOI] [PubMed] [Google Scholar]
  • 10.Collins, K. H., Reimer, R. A., Seerattan, R. A., Leonard, T. R. & Herzog, W. Using diet-induced obesity to understand a metabolic subtype of osteoarthritis in rats. Osteoarthritis Cartilage23(6), 957–965. 10.1016/j.joca.2015.01.015 (2015). [DOI] [PubMed] [Google Scholar]
  • 11.Sun, A. R. J. et al. Obesity-associated metabolic syndrome spontaneously induces infiltration of pro-inflammatory macrophage in synovium and promotes osteoarthritis. PLoS ONE10.1371/journal.pone.0183693 (2017). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 12.U. White, Adipose tissue expansion in obesity, health, and disease, (2023), Frontiers Media S.A. [DOI] [PMC free article] [PubMed]
  • 13.Lindhorst, A. et al. Adipocyte death triggers a pro-inflammatory response and induces metabolic activation of resident macrophages. Cell Death Dis.10.1038/s41419-021-03872-9 (2021). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 14.M. Kuroda and H. Sakaue, Adipocyte Death and Chronic Inflammation in Obesity, J. Med. Investigat. (2017) [DOI] [PubMed]
  • 15.L. Boutens and R. Stienstra, Adipose tissue macrophages: going off track during obesity, (2016), Springer Verlag. [DOI] [PMC free article] [PubMed]
  • 16.Michaud, A., Drolet, R., Noël, S., Paris, G. & Tchernof, A. Visceral fat accumulation is an indicator of adipose tissue macrophage infiltration in women. Metabolism61(5), 689–698. 10.1016/j.metabol.2011.10.004 (2012). [DOI] [PubMed] [Google Scholar]
  • 17.Courties, A., Berenbaum, F. & Sellam, J. The phenotypic approach to osteoarthritis: a look at metabolic syndrome-associated osteoarthritis. Joint Bone Spine86(6), 725–730. 10.1016/j.jbspin.2018.12.005 (2019). [DOI] [PubMed] [Google Scholar]
  • 18.Collins, K. H. et al. Adipose tissue is a critical regulator of osteoarthritis. Proceed. Nat. Acad. Sci.10.1073/PNAS.2021096118 (2021). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 19.C. Xie and Q. Chen, Adipokines: New Therapeutic Target for Osteoarthritis?, (2019), Springer. [DOI] [PMC free article] [PubMed]
  • 20.Jääskeläinen, I. et al. Collagens regulating adipose tissue formation and functions. MDPI10.3390/biomedicines11051412 (2023). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 21.Belluzzi, E. et al. Contribution of infrapatellar fat pad and synovial membrane to knee osteoarthritis pain. Hindawi Limited.10.1155/2019/6390182 (2019). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 22.A. K. Beery and I. Zucker, Sex bias in neuroscience and biomedical research, (2011). 10.1016/j.neubiorev.2010.07.002. [DOI] [PMC free article] [PubMed]
  • 23.Maric, I. et al. Sex and species differences in the development of diet-induced obesity and metabolic disturbances in rodents. Front. Nutrition9, 1–14. 10.3389/fnut.2022.828522 (2022). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 24.Abughazaleh, N. et al. Development of shoulder osteoarthritis and bone lesions in female and male rats subjected to a high fat/sucrose diet. Sci. Rep.14(1), 25871. 10.1038/s41598-024-76703-4 (2024). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 25.Rios, J. L. et al. Protective effect of prebiotic and exercise intervention on knee health in a rat model of diet-induced obesity. Sci. Rep.9(1), 3893. 10.1038/s41598-019-40601-x (2019). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 26.Kawai, T., Autieri, M. V. & Scalia, R. Adipose tissue inflammation and metabolic dysfunction in obesity. Am. J. Physiol. Cell Physiol.320(3), C375–C391. 10.1152/ajpcell.00379.2020 (2021). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 27.Poret, J. M., Gaudet, D. A., Braymer, H. D. & Primeaux, S. D. Sex differences in markers of metabolic syndrome and adipose tissue inflammation in obesity-prone, Osborne-Mendel and obesity-resistant, S5B/Pl rats. Life Sci.10.1016/j.lfs.2021.119290 (2021). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 28.Lam, Y. Y. et al. Increased gut permeability and microbiota change associate with mesenteric fat inflammation and metabolic dysfunction in diet-induced obese mice. PLoS ONE10.1371/journal.pone.0034233 (2012). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 29.Horton, T. J., Gayles, E. C., Prach, P. A., Koppenhafer, T. A. & Pagliassotti, M. J. Female rats do not develop sucrose-induced insulin resistance. American J. Physiol. Regul. Integrative Comparative Physiol.272(5), R1571–R1576 (1997). [DOI] [PubMed] [Google Scholar]
  • 30.Rudnicki, M. et al. Female mice have higher angiogenesis in perigonadal adipose tissue than males in response to high-fat diet. Front. Physiol.10.3389/fphys.2018.01452 (2018). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 31.Michailidou, Z. et al. Increased angiogenesis protects against adipose hypoxia and fibrosis in metabolic disease-resistant 11β-hydroxysteroid dehydrogenase type 1 (HSD1)-deficient mice. J. Biol. Chem.287(6), 4188–4197. 10.1074/jbc.M111.259325 (2012). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 32.Chait, A. & den Hartigh, L. J. Adipose tissue distribution, inflammation and its metabolic consequences, including diabetes and cardiovascular disease. Front. Med. S.A10.3389/fcvm.2020.00022 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 33.D. J. Clegg, L. M. Brown, S. C. Woods, and S. C. Benoit, Gonadal Hormones Determine Sensitivity to Central Leptin and Insulin, (2006).[Online]. Available: http://diabetesjournals.org/diabetes/article-pdf/55/4/978/384244/zdb00406000978.pdf [DOI] [PubMed]
  • 34.Rytka, J. M., Wueest, S., Schoenle, E. J. & Konrad, D. The portal theory supported by venous drainage-selective fat transplantation. Diabetes60(1), 56–63. 10.2337/db10-0697 (2011). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 35.Studies in the Area of Lactobacillus Reported from Jiangnan University (Lactobacillus Strains Derived From Human Gut Ameliorate Metabolic Disorders Via Modulation of Gut Microbiota Composition and Short-chain Fatty Acids Metabolism), (2021).[Online]. Available: http://www.Newsrx.com [DOI] [PubMed]
  • 36.Companys, J. et al. Gut microbiota profile and its association with clinical variables and dietary intake in overweight/obese and lean subjects: A cross-sectional study. Nutrients10.3390/nu13062032 (2021). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 37.Rodríguez, A. M., Quevedo-Coli, S., Roca, P. & Palou, A. Sex-dependent dietary obesity, induction of UCPs, and leptin expression in rat adipose tissues. Obes Res9(9), 579–588. 10.1038/oby.2001.75 (2001). [DOI] [PubMed] [Google Scholar]
  • 38.Kim, S. N. et al. Sex differences in sympathetic innervation and browning of white adipose tissue of mice. Biol. Sex Differ.10.1186/s13293-016-0121-7 (2016). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 39.Rodríguez-Cuenca, S. et al. Sex-dependent thermogenesis, differences in mitochondrial morphology and function, and adrenergic response in brown adipose tissue. J. Biol. Chem.277(45), 42958–42963. 10.1074/jbc.M207229200 (2002). [DOI] [PubMed] [Google Scholar]
  • 40.Choi, D. K. et al. gender difference in proteome of brown adipose tissues between male and female rats exposed to a high fat diet. Cell. Physiol. Biochem.28(5), 933–948 (2011). [DOI] [PubMed] [Google Scholar]
  • 41.Y. H. Gao et al., An update on the association between metabolic syndrome and osteoarthritis and on the potential role of leptin in osteoarthritis, (2020), Academic Press. 10.1016/j.cyto.2020.155043. [DOI] [PubMed]
  • 42.Z. Liu et al., Novel perspectives on leptin in osteoarthritis: Focus on aging, KeAi Communications Co. (2024) 10.1016/j.gendis.2023.101159. [DOI] [PMC free article] [PubMed]
  • 43.Frost, M. et al. The relationship between bone turnover and insulin sensitivity and secretion: Cross-sectional and prospective data from the RISC cohort study. Bone108, 98–105. 10.1016/j.bone.2017.12.029 (2018). [DOI] [PubMed] [Google Scholar]
  • 44.Shanbhogue, V. V., Finkelstein, J. S., Bouxsein, M. L. & Yu, E. W. Association between insulin resistance and bone structure in nondiabetic postmenopausal women. J. Clin. Endocrinol. Metab.101(8), 3114–3122. 10.1210/jc.2016-1726 (2016). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 45.Hardcastle, S. A. et al. Prevalence of radiographic hip osteoarthritis is increased in high bone mass. Osteoarthritis Cartilage22(8), 1120–1128. 10.1016/j.joca.2014.06.007 (2014). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 46.Hardcastle, S. A. et al. Individuals with high bone mass have an increased prevalence of radiographic knee osteoarthritis. Bone71, 171–179. 10.1016/j.bone.2014.10.015 (2015). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 47.Nevitt, M. C. et al. Radiographic osteoarthritis of the hip and bone mineral density. Arthritis Rheum.38(7), 907–916. 10.1002/art.1780380706 (1995). [DOI] [PubMed] [Google Scholar]
  • 48.M. Sowers, M. Hochberg, J. P. Crabbe, A. Muhich, M. Crutchfield, and S. Updike, Association of Bone Mineral Density and Sex Hormone Levels with Osteoarthritis of the Hand and Knee in Premenopausal Women, 1996.[Online]. Available: https://academic.oup.com/aje/article/143/1/38/63192 [DOI] [PubMed]
  • 49.Kato, M. & Onodera, T. Spontaneous osteochondrosis in rats. Lab Anim.18, 179–187 (1984). [DOI] [PubMed] [Google Scholar]
  • 50.Kato, M. & Onodera, T. Morphological investigation of osteochondrosis induced by ofloxacin in rats. Fundament. Appl. Toxicol.11, 120–133 (1988). [DOI] [PubMed] [Google Scholar]
  • 51.Emmanuelle, N. E. et al. Critical role of estrogens on bone homeostasis in both male and female: From physiology to medical implications. MDPI AG10.3390/ijms22041568 (2021). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 52.Masarwi, M., Shamir, R., Phillip, M. & Gat-Yablonski, G. Leptin stimulates aromatase in the growth plate: Limiting catch-up growth efficiency. J. Endocrinol.237(3), 229–242. 10.1530/JOE-18-0028 (2018). [DOI] [PubMed] [Google Scholar]
  • 53.Nilsson, O. et al. Evidence that estrogen hastens epiphyseal fusion and cessation of longitudinal bone growth by irreversibly depleting the number of resting zone progenitor cells in female rabbits. Endocrinology155(8), 2892–2899. 10.1210/en.2013-2175 (2014). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 54.Abughazaleh, N., Seerattan, R. A., Hart, D. A., Reimer, R. A. & Herzog, W. A novel Osteoarthritis scoring system to separate typical OA joint degeneration from non-typical lesions in male Sprague Dawley rats. Osteoarthr. Cartil. Open10.1016/j.ocarto.2024.100521 (2024). [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 55.Seifer, D. R. et al. Novel synovial fluid recovery method allows for quantification of a marker of arthritis in mice. Osteoarthritis Cartilage16(12), 1532–1538. 10.1016/j.joca.2008.04.013 (2008). [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Supplementary Information. (74.7MB, docx)

Data Availability Statement

The data that support the findings of this study are available from the Human Performance lab (HPL) at the University of Calgary, The data are available from the corresponding author (Nada Abughazaleh) upon request.

Articles from Scientific Reports are provided here courtesy of Nature Publishing Group

RESOURCES