Repeated (Weekly) Intra‐Articular Injections of Sulfated Galactans Attenuate Cartilage Degeneration in a Rat Model of Osteoarthritis
Nirada Srianake, Amarin Thongsuk, Scarlett Desclaux, Thitiya Phuagpan, Jiraporn Sriwong, Alita Kongchanagul, Wanwisa Himakhun, Ruedee Hemstapat, Kanokpan Wongprasert
TL;DR
Sulfated galactans from a red alga reduced cartilage degeneration in a rat model of osteoarthritis, suggesting potential for joint protection.
Contribution
Demonstrates sulfated galactans from Gracilaria fisheri can preserve cartilage in osteoarthritis models.
Findings
SG at 500 µg significantly attenuated cartilage degeneration in OA rats.
SG suppressed matrix-degrading enzymes and enhanced cartilage-protective markers in human chondrocytes.
SG treatment improved cartilage structure preservation in histological assessments.
Abstract
Osteoarthritis (OA) is a progressive joint disease characterized primarily by pain, leading to substantial impairment of quality of life. Current treatments primarily alleviate symptoms but have limited efficacy in protecting or repairing articular cartilage. Marine algae have recently gained attention in pharmaceutical research due to their diverse bioactivities. Gracilaria fisheri, a red alga, contains abundant sulfated galactans (SG) that may exert chondroprotective effects. This study investigated the effects of SG on pain‐related behaviors and cartilage degeneration in a Wistar rat OA model induced by anterior cruciate ligament transection combined with medial meniscus removal (ACLT + MMx). Pain‐related behaviors were monitored weekly using a hind limb weight‐bearing distribution test. Four weeks post‐surgery, rats received intra‐articular injections (50 µL) of normal saline (NSS),…
Genes, proteins, chemicals, diseases, species, mutations and cell lines named across the full text — each resolved to its canonical identifier and authoritative record.
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Figure 8| Gene | Primer sequences (5′−3′) |
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Forward: CTGACTTCAACAGCGACACC Reverse: TGCTGTAGCCAAATTCGTTG |
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Forward: GAAGCTGCTTACGAATTTGCCG Reverse: CCAAAGGAGCTGTAGATGTCCT |
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Forward: TGACTGAGAGGCTCCGAGAA Reverse: CATCAGGAACCCCGCATCTT |
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Forward: GTGTCTGCGGATACTTCCACAG Reverse: AGCTAAGCTCAGGCTGTTCCAG |
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Forward: CCTGGTCTTGGTGGAAACTT Reverse: CAGAGACACCAGGTTCACCA |
- —National Science Research and Innovation Fund (NSRF)
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Taxonomy
TopicsSeaweed-derived Bioactive Compounds · Osteoarthritis Treatment and Mechanisms · Phytochemical and Pharmacological Studies
Introduction
1
Osteoarthritis (OA) is a chronic degenerative joint disease that primarily affects weight‐bearing joints, such as the knees. Knee OA ranks among the top ten causes of disability in the elderly and poses a significant public health challenge, particularly in Thailand, where it substantially impairs patients' quality of life. Articular cartilage, which covers the joint surfaces, is avascular and aneural, limiting its intrinsic repair capacity [1]. Cartilage damage, caused by trauma or repetitive mechanical stress, heals slowly and often incompletely; ongoing mechanical loading accelerates cartilage erosion, ultimately leading to OA [2, 3].
Clinically, OA manifests as joint pain, stiffness, functional impairment, and structural degeneration, including cartilage destruction, subchondral bone changes, synovial inflammation, and osteophyte formation. These pathological alterations arise from an imbalance in chondrocyte activity, particularly between the synthesis and degradation of extracellular matrix (ECM) components such as type II collagen, aggrecan, glycosaminoglycans, and hyaluronan [4]. In response to inflammatory and mechanical stimuli, chondrocytes secrete pro‐inflammatory cytokines (e.g., IL‐1β, TNF‐α) and matrix‐degrading enzymes (e.g., matrix metalloproteinases; MMPs), promoting ECM breakdown and impairing cartilage regeneration [5, 6].
In the early stages, OA is treated symptomatically. Moderate cases may be managed with intra‐articular injections, while advanced disease often requires surgical intervention. Conventional therapies, including nonsteroidal anti‐inflammatory drugs (NSAIDs), glucocorticoids, hyaluronic acid injections, and joint replacement can alleviate symptoms but do not restore cartilage, and carry risks of adverse effects [7]. Recent tissue engineering approaches aim to stimulate cartilage regeneration; however, these often fail to achieve effective cell adhesion at the injury site, leading to incomplete tissue repair [8].
Biological therapies have gained interest for cartilage regeneration. Intra‐articular hyaluronic acid improves joint lubrication and promotes ECM precursor synthesis, providing symptomatic relief for moderate OA patients who are not yet candidates for surgery [9, 10]. Platelet‐rich plasma (PRP), rich in growth factors, further enhances natural tissue repair [11]. Cell‐based therapies using in vitro‐expanded chondrocytes also show promise [3], particularly when combined with scaffolds such as hyaluronic acid or sulfated polysaccharides [12]. These scaffolds facilitate chondrocyte adhesion, survival, proliferation, and tissue regeneration. For instance, Wei et al., reported that co‐injecting bone marrow‐derived stromal stem cells with hyaluronic acid significantly reduced cartilage and subchondral bone damage in an OA goat knee model [13]. While synthetic scaffolds have limitations, natural scaffolds, particularly sulfated polysaccharides derived from marine sources, are emerging as attractive alternatives due to their bioactivity, biocompatibility, and ability to enhance cell adhesion, proliferation, and growth factor interactions, thereby improving regenerative capacity [14].
Marine algae, rich in sulfated polysaccharides, have broad biological activities. Sulfated polysaccharides from brown algae reduce OA symptoms, promote type II collagen production, upregulate chondrogenic genes, and protect cartilage in experimental models [15, 16]. In southern Thailand, red algae such as G. fisheri are abundant and contains sulfated galactans (SG), structurally similar to heparan sulfate proteoglycans, essential components for cartilage ECM organization, cell adhesion, and growth factor signaling [17]. SG from G. fisheri enhance collagen synthesis, promote ECM deposition, and are non‐cytotoxic in various cell types [18, 19, 20]. In vitro, they possess antioxidant properties, stimulate collagen synthesis, and inhibit ECM degradation in fibroblasts [18], while in vivo studies of related Gracilaria species (e.g., G. cornea, G. caudata, G. crinale) demonstrate anti‐inflammatory and anti‐nociceptive effects [21, 22, 23, 24].
We hypothesize that SG derived from G. fisheri can alleviate OA symptoms and promote cartilage repair by enhancing ECM synthesis, inhibiting ECM degradation, and supporting chondrocyte function. An anterior cruciate ligament transection combined with medial meniscectomy (ACLT + MMx)‐induced OA rat model was employed, as it is widely used and well‐validated model that reproduces cartilage degeneration and joint structural changes characteristic of human OA [25]. Using this model, we evaluated the therapeutic potential of SG, which may serve as a natural scaffold for cartilage regeneration, either alone or in combination with cell‐based therapies. These findings provide a foundation for developing SG‐based strategies to improve OA treatment and alleviate the healthcare burden in aging populations.
Materials and Methods
2
Sulfated Galactans (SG)
2.1
Sulfated galactans (SG) were extracted from the marine red alga Gracilaria fisheri following established protocols [17]. Briefly, dried algal material was ground into a fine powder and sequentially depigmented with benzene and acetone. SG was extracted using a cold‐water extraction method, precipitated with absolute ethanol, and further purified by DEAE‐Sepharose fast‐flow anion‐exchange chromatography. The purified fraction was dialyzed against distilled water (molecular weight cutoff, 10,000 Da) and freeze‐dried. Structural characterization of SG was performed using nuclear magnetic resonance (NMR) and Fourier‐transform infrared (FT‐IR) spectroscopy, revealing a linear backbone composed of 3‐linked‐β‐d‐galactopyranose (G) and 4‐linked 3,6‐anhydro‐α‐l‐galactopyranose (LA) or α‐l‐galactose‐6‐sulfate (L6S), with partial methylation at C‐2 of LA and C‐6 of G, and sulfation at C‐4 and C‐6 of d‐galactopyranose units (G4S and G6S). The final product exhibited approximately 90% purity, as determined by HPLC analysis (Waters, Milford, MA, USA). The average molecular weight of SG was approximately 228 kDa, with a total carbohydrate content of 67%, sulfate content of 10%, and solution pH of 6 [26]. Purified SG was stored as a freeze‐dried powder, aliquoted, and kept at −20°C until use. For experimental applications, SG working solutions were freshly prepared at 10 mg/mL in Milli‐Q water and fully dissolved at 37°C until a homogeneous solution was obtained. SG was readily soluble in aqueous solutions, including normal saline solution and cell culture media. All working concentrations were prepared immediately before use.
Animals and Experimental Design
2.2
All animal procedures were approved by the Institutional Animal Care and Use Committee (IACUC) of the Faculty of Science, Mahidol University (Approved No. MUSC66‐061‐691) and conducted in accordance with the ARRIVE guidelines [27]. Forty‐two male Wistar rats (8–9 weeks old, 250–280 g) were obtained from Nomura Siam International Co Ltd., (Bangkok, Thailand). Animals were housed in pairs in an Association for the Assessment and Accreditation of Laboratory Animal Care (AAALAC)‐accredited facility and acclimatized for 5–7 days before study initiation. Housing conditions were maintained at 22 ± 1°C with 30%–70% relative humidity under a 12‐h light/dark cycle. Standard rat chow and water were provided ad libitum.
This study was designed to evaluate the potential of early SG treatment in preventing cartilage degeneration in a surgically‐induced knee OA model. Forty‐two male Wistar rats were randomly assigned to six groups (n = 7/group) as illustrated in Figure 1. The sample sizes were determined based on our previous study [28]. To confirm OA development, osteoarthritic features, including joint pain and cartilage degeneration were assessed using behavioral and histopathological assessments following OA induction. Knee joint pain progression was monitored weekly using the weight bearing distribution test, while the cartilage damage severity was assessed by histopathological examinations. Four weeks after OA induction, each animal received an intra‐articular injection (50 μL) of normal saline solution (NSS), a low‐to‐intermediate molecular weight hyaluronic acid (HA; Ostenil®; molecular weight: 1.2–1.4 × 10^6^ Da, TRB Chemedica, Geneva, Switzerland), a clinically approved and widely used intra‐articular HA formulation for the treatment of knee OA, or SG (2.5, 50 or 500 μg) into the right knee joint once weekly for four consecutive weeks (Figure 1) [29]. Injections were administered through the patellar ligament using a 26 G needle attached to a Hamilton microsyringe. Each injection was delivered over 10 s, after which the knee was gently flexed and extended three times to facilitate sample distribution within the joint cavity.
Schematic diagram summarizing the experimental design (Created with Biorender.com/Mahidol University).
Induction of Surgically‐Induced Knee OA
2.3
Knee OA was induced in 10‐week‐old male Wistar rats (332.5 ± 7.5 g) by anterior cruciate ligament transection combined with medial meniscectomy (ACLT + MMx). Animals were anaesthetized with inhaled isoflurane (Attane™, Piramal Critical Care. Inc., USA) in oxygen (5% for induction and 1.5%–3% for maintenance). To prevent infection, cefazolin (20 mg/kg; Cefaben; L.B.S laboratory Ltd, Part., Bangkok, Thailand) was administered subcutaneously 30 min before surgery and once daily for five consecutive days postoperatively. The ACLT + MMx procedure was performed as previously described [30, 31]. Briefly, the right knee was shaved and disinfected with 70% ethanol, followed by povidone‐iodine (Figure 2A). A skin incision was made over the knee joint, and the soft tissue was bluntly dissected using small scissors (Figure 2B). A longitudinal incision was made on the medial side of the joint capsule using a No. 21 scalpel blade (Figure 2C), and the patella was dislocated laterally to expose the femoral condyles and anterior cruciate ligament (ACL) (Figure 2D). The ACL was transected (Figure 2E,F), followed by transection of the medial collateral ligament (MCL) with a No. 11 surgical blade (Figure 2G,H). Successful ACL transection was verified by the anterior drawer test, after which the medial meniscus (C‐shaped cartilage) was excised using Vannas spring scissors (Figure 2I,J). The joint capsule was closed with 4‐0 Vicryl absorbable sutures (Ethicon® Inc., Livingston, UK) (Figure 2K), and the skin was sutured with 4‐0 braided non‐absorbable silk sutures (Mersilk 4‐0 braided silk non‐absorbable suture; Ethicon® Inc., Livingston, UK) (Figure 2L). The wound was cleaned with 0.9% normal saline and disinfected with povidone‐iodine. Sham‐operated animals underwent identical surgical procedures, except that the ACL and MCL were not transected and the medial meniscus was preserved. Postoperative care and welfare monitoring were carried out following recovery from anesthesia.
Induction of surgically induced knee osteoarthritis (OA) in a rat model by an anterior cruciate ligament transection (ACLT) and meniscectomy (MMx) (ACLT + MMx). (A) The surgical area on the right knee was shaved and disinfected. (B–D) Incisions were made in the skin and knee joint capsule, and the patella was dislocated laterally to expose the femoral condyles and ACL. (E–J) The ACL and MCL were transected using a No. 11 surgical blade, followed by excision of the medial meniscus with Vannas spring scissors. (K–L) The joint capsule and skin were closed with absorbable and braided non‐absorbable sutures, respectively.
Assessment of Pain‐Like Behavior
2.4
In this study, the term “pain‐like behavior” refers specifically to spontaneous, joint‐related pain and functional impairment assessed by hind limb weight‐bearing distribution asymmetry. Pain‐like behavior was assessed weekly for up to 8 weeks post‐OA induction using an incapacitance meter (Columbus Instruments, Columbus, OH, USA) to determine weight distribution asymmetry between the ipsilateral (injured limb) and contralateral hind limb. The investigators who performed the behavior tests were not informed of the experimental condition of the group being test. Data were expressed as the % hind limb weight distribution (% HLWD), as previously described [32, 33]. Both pain assessment and body weights were assessed prior to OA induction and monitored weekly thereafter. Following the final pain assessment at week eight post‐OA induction, animals were euthanized via intraperitoneal injection of thiopental (150 mg/kg; Anesthal®; Scott‐Edil Pharmacia Ltd., Solan, India) and xylazine (10 mg/kg; L.B.S. Laboratory Ltd. Part., Bangkok, Thailand) at an overdose level. The knee joint tissues from the operated hind limb (right knee) of each animal were collected for further histological evaluation.
Specimen Preparation and Histological Evaluation
2.5
In brief, knee joint tissues were fixed in 10% neutral buffered formalin for 48 h, followed by decalcification in 10% ethylenediaminetetraacetic acid (EDTA) (Elago Enterprises Pty Ltd., New South Wales, Australia) at 4°C for 5 weeks, with the solution replaced twice weekly. After decalcification, each knee joint was bisected in the sagittal plane to obtain two approximately equal halves (medial and lateral aspects). The medial portion was embedded in paraffin blocks with the medial aspect oriented downward. The paraffin blocks were trimmed approximately 150 μm from the medial margin of the joint and then serially sectioned at 4 μm thickness along the sagittal plane to obtain three sets of 20 sections, collected at 80 μm intervals (total step size of 240 μm). This sectioning protocol ensured a comprehensive assessment of lesions of varying severity across the tibial plateau. Tissue sections were stained with hematoxylin and eosin (H&E; Cat# MHS16; Merck Millipore Darmstadt, Germany) for morphological evaluation and with Safranin‐O/Fast green (Panreac, Darmstadt, Germany) to visualize matrix proteoglycans in the articular cartilage. Sections were imaged in the tibial plateau cartilage using a Nikon light microscope with NIS‐Elements D software (version 5.21) at 4 × and 10 × magnification. The severity of cartilage lesion was independently assessed by two observers (N.S. and W.H.), who were unaware of the treatment allocation, using the Osteoarthritis Research Society International (OARSI) scoring system, which evaluates both the severity and the extent of cartilage damage [34] (see Supporting Information). The specimen preparation procedures are illustrated in Figure 3.
Schematic diagram summarizing the stepwise procedures for specimen preparation, including fixation, decalcification, and sectioning (Created with Biorender.com/Mahidol University).
Immunohistochemical Analysis
2.6
Sections were deparaffinized, rehydrated, and subjected to antigen retrieval in sodium citrate buffer (10 mM sodium citrate, 0.05% Tween 20, pH 6.0) at 70° C for 20 min. Endogenous peroxidase activity was blocked by incubation in 0.3% hydrogen peroxide (Chem‐Supply Pty. Ltd., Gillman, Australia) in methanol for 15 min at room temperatures, followed by blocking of non‐specific binding with 1% normal goat serum (Invitrogen Corporation, MD, USA) and 1% bovine serum albumin (Sigma‐Aldrich, MO, USA) for 1 h at room temperature. Sections were then incubated with anti‐collagen II antibody (1:200, Cat# ab34712; Abcam, Cambridge, UK) on a rocker shaker at 4° C overnight. For secondary antibody detection, biotin‐conjugated antibody (Cat# ab64256; Abcam, Cambridge, UK) and streptavidin (Cat# ab64269; Abcam, Cambridge, UK) were applied for 10 min each. Antibody binding was visualized using 3, 3′‐diaminobenzidine (DAB; Cat# ab64238, Abcam, Cambridge, USA) chromogenic kit, followed by counterstaining with Mayer's hematoxylin solution (Cat# 05‐06002, Bio‐optica, Milano, Italy).
Image Analysis
2.7
All sections were imaged in the tibial plateau cartilage using a Nikon light microscope with NIS‐Elements D software (version 5.21) at 4 × and 10 × magnification, and regions of interest (ROIs; 75 × 50 μm), primarily located in the posteromedial tibial plateau, were analyzed using NIS‐Elements BR software (version 5.41) with background subtraction. Data were expressed as the percentage of the ROI area showing positive staining.
Cell Culture of Human Chondrocytes and RT‐qPCR Analysis
2.8
C28/I2 human chondrocytes were obtained from Merck Millipore (Temecula, California, USA). Cells were cultured in Dulbecco's Modified Eagle Medium (DMEM; Gibco, Life Technologies, USA) supplemented with 10% fetal bovine serum (FBS; Gibco Life Technologies, USA) and 1% antibiotic‐antimycotics (Gibco, USA). Cultures were maintained at 37°C in a humidified atmosphere containing 5% CO₂.
To investigate the effects of SG on the expression of cartilage matrix‐related genes under inflammatory conditions, C28/I2 cells were stimulated with interleukin‐1 beta (IL‐1β, 10 ng/mL) for 12 h, followed by treatment with SG at concentrations of 1, 10, or 20 µg/mL for 24 h. The SC concentrations were selected based on our previous study [35]. Total RNA was extracted using the RiboEx™ kit (GeneAll Biotechnology, Korea), and RNA concentrations were quantified with a NanoDrop™ ND‐2000C spectrophotometer (Thermo Scientific, USA). Complementary DNA (cDNA) was synthesized from 1 µg of total RNA using the RevertAid First Strand cDNA Synthesis Kit (Thermo Scientific, USA). Quantitative real‐time polymerase chain reaction (RT‐qPCR) was carried out using iTaq Universal SYBR® Green Supermix (Bio‐Rad Laboratories, USA) and gene‐specific primers (Table 1) on a Bio‐Rad CFX real‐time PCR system. The target genes included COL2A1, TIMP‐1, MMP‐1, and MMP‐13. Gene expression was normalized to GAPDH, and relative expression levels were calculated using the 2^–ΔΔCt method [36].
Statistical Analysis
2.9
All statistical analyses were performed using GraphPad Prism version 10 (GraphPad Software, La Jolla, CA, USA). Data are presented as mean ± standard error of the mean (SEM). The Shapiro‐Wilks test was used to assess the normality of the data. Pain‐related behavior data were analyzed by one‐way ANOVA, followed by Dunnett's multiple comparisons test. Histological data were analyzed using the Kruskal‐Wallis test followed by Dunn's post hoc test. All in vitro experiments were independently conducted in triplicate and analyzed by one‐way ANOVA followed by Tukey's post hoc test. A p‐value < 0.05 was considered statistically significant.
Result
3
General Animal Health
3.1
All animals maintained good general health throughout the study period. The mean (± SEM) body weight gain in all experimental groups was comparable to that of the sham + NSS group during the 4 weeks of intra‐articular (i.a.) SG injections. No signs of adverse reactions were detected at the injection sites in any animals receiving i.a. SG injections at doses up to 500 μg for four consecutive weeks (Figure 4).
Mean (± SEM) body weights were not significantly different (p > 0.05) among different experimental groups.
Repeated (Weekly) Intra‐Articular SG Injections Did Not Attenuate Pain‐Related Behavior in ACLT + MMx Rats
3.2
One week after OA induction, all experimental groups exhibited a marked reduction in the mean ± SEM percentage of hind limb weight distribution (% HLWD), reflecting postoperative pain likely caused by the surgical procedure (Figure 5A). In the sham + NSS group, %HLWD returned to baseline levels by 3 weeks post‐surgery. In contrast, the OA group displayed significant and persistent pain from week 5 onward. Furthermore, no significant differences were observed in the mean ± SEM% HLWD among OA groups receiving HA or SG injections at doses up to 500 μg compared with the vehicle‐treated OA group at any time point. Similarly, analysis of the extent and duration of weight bearing on the operated hind limb, expressed as the area under the curve (AUC), revealed no significant differences in the % HLWD AUC values among the treatment groups relative to the vehicle‐treated OA group (Figure 5B).
*The pain‐like behavioral was expressed as the mean ± SEM percentage (% mean ± SEM) of weight‐bearing distribution on the operated hind limb. (A) The effect of intra‐articular injection of SG on changes in the % mean ± SEM of weight bearing on the operated hind limb was evaluated. (B) The corresponding AUC values of % mean of weight bearing on the operated hind limb in in each group of animals. Each point represents the mean ± SEM of each group (7 rats/group). Statistical differences among experimental groups were analyzed using a one‐way ANOVA, followed by Dunnett's multiple comparisons test. ***p < 0.0001 vs. the OA + NSS, indicating statistical significance.
Repeated (Weekly) Intra‐Articular SG Injections Attenuate Cartilage Degeneration in ACLT + MMx Rats. Histological Assessment of Cartilage Degeneration
3.3
At 8 weeks post‐surgery, H&E and Safranin O/Fast Green staining revealed pronounced cartilage degeneration in the OA + NSS group, including surface fibrillation and denudation, chondrocyte death, cartilage matrix proteoglycan loss, reparative tissue and osteophyte formation compared with the Sham group (Figure 6E–H vs. A–D). In contrast, HA or SG treatment at all doses partially preserved cartilage integrity, with the most evident protection observed in the SG 500 µg group (Figure 6U–Y). Safranin O staining demonstrated substantial retention of proteoglycan in the SG 500 µg group, comparable to HA treatment. Quantitative OARSI scoring confirmed these observations, showing significantly lower scores in the SG 500 µg group compared with OA + NSS (*p *< 0.05), indicating attenuation of cartilage degeneration (Figure 6Z).
Histological evaluation of cartilage degeneration after SG treatment. Representative images of articular cartilage from the tibial plateau at 8 weeks after ACLT + MMx surgery in rats. Hematoxylin and eosin (H&E) and Safranin O/Fast Green staining are shown for the following groups, n = 7 each: (A–D) Sham + NSS (OARSI grade 1, stage 1): (A–B) H&E shows intact cartilage surface. (C) Safranin O shows uniform matrix staining. (D) Mild chondrocyte clustering (white arrow); (E–H) OA + NSS (OARSI grade 5, stage 3): H&E and Safranin O show surface denudation with subchondral bone sclerosis (arrowheads) and fibrocartilage formation (). (G–H) 25%–50% of surface with irregularity and complete loss of Safranin O staining (black arrow); (I–L) OA + HA (OARSI grade 4–5, stage 3): Cartilage erosion to mid–deep zones (arrowheads), involving 25%–50% of surface. (L) Loss of Safranin O staining throughout the cartilage depth (black arrow); (M–P) OA + 2.5 µg SG (OARSI grade 4–5, stage 3): Surface delamination, fibrocartilage formation (), and erosions (arrowheads) affecting 25%–50% of surface. (N, P) Hypocellularity (white arrow); (P) diffuse Safranin O loss (black arrow); (Q–T) OA + 50 µg SG (OARSI grade 4–5, stage 3): Delamination, fibrocartilage (), and hypocellularity (white arrow). (T) Chondrocyte hypertrophy and clustering (circle). Surface defect 25%–50% with partial Safranin O loss in upper one‐third (black arrow); (U–Y) OA + 500 µg SG (OARSI grade 4–5, stage 2): Minimal fibrocartilage (*), localized surface discontinuity (arrowheads), and hypocellularity (white arrow). (Y) Chondrocyte clustering (circle). Defect 10%–25% with Safranin O loss in upper one‐third (black arrow); (Z) OARSI scoring of cartilage degeneration. Scale bars: 1000 µm (A–E), 100 µm (F–J). Data are presented as mean ± SEM. Statistical analysis was performed using the Kruskal–Wallis test followed by Dunn's post hoc test, *p < 0.05, ***p < 0.00001 compared to OA + NSS.
Type II Collagen Immunohistochemistry
3.4
Immunohistochemical analysis further evaluated cartilage matrix composition. Sham‐operated rats exhibited abundant type II collagen in the articular cartilage (Figure 7A,F). In contrast, OA + NSS animals showed a marked reduction in staining (Figure 7B,G). HA and SG‐treated groups displayed partial restoration of type II collagen distribution, although the difference did not reach statistical significance compared with OA controls (Figure 7K). The SG 500 µg group showed a trend toward higher type II collagen staining than OA + NSS.
*Immunohistochemical staining of type II collagen in articular cartilage of the tibial plateau at 8 weeks after ACLT + MMx surgery in rats. (A, F) Sham + NSS (n = 6); (B, G) OA + NSS (n = 7); (C, H) OA + HA (n = 7); (D, I) OA + SG 50 µg (n = 7); (E, J) OA + SG 500 µg (n = 7). Brown staining indicates type II collagen distribution. Scale bars: 1000 µm (A–E), 100 µm (F–J). (K) Quantification of type II collagen–positive area. Data are present as mean ± SEM. Statistical significance was assessed using the Kruskal‐Wallis test followed by Dunn's post‐hoc test, *p < 0.01 compared to Sham + NSS.
SG Reduces IL‐1β‐Induced Extracellular Matrix Degradation and Promotes Matrix Synthesis in C28/I2 Chondrocytes
3.5
C28/I2 human chondrocytes were stimulated with IL‐1β (10 ng/mL) for 12 h to induce an inflammatory response. IL‐1β significantly upregulated the mRNA levels of matrix metalloproteinases MMP‐1 and MMP‐13 compared to untreated control cells (p < 0.001) (Figure 8A–B). Concurrently, IL‐1β suppressed the expression of the endogenous MMP inhibitor, TIMP‐1 (p < 0.01), and the key cartilage matrix component, COL2A1 (p < 0.05) (Figure 8C–D). Post‐treatment with SG at concentrations of 1, 10, and 20 µg/mL substantially reversed these effects. SG significantly reduced IL‐1β–induced MMP‐1 and MMP‐13 expression toward baseline levels (p < 0.01 and p < 0.001, respectively). Moreover, SG markedly upregulated TIMP‐1 and COL2A1 expression, exceeding levels of untreated controls (p < 0.001 and p < 0.01, respectively). These findings demonstrate that SG exhibits potential as a chondroprotective agent under inflammatory conditions.
*Effect of SG on IL‐1β‐induced expression of genes related to cartilage matrix degradation and synthesis in C28/I2 chondrocytes. Cells were pre‐treated with IL‐1β (10 ng/mL) for 12 h, followed by post‐treatment with SG (1, 10, and 20 µg/mL) for 24 h. IL‐1β significantly upregulated the expression of MMP‐1 (A) and MMP‐13 (B) and downregulated TIMP‐1 (C) and COL2A1 (D), compared to untreated controls. Post‐treatment with SG reversed these effects of IL‐1β. Gene expression levels were quantified by RT‐qPCR and normalized to GAPDH using the 2^–ΔΔCt method. Data are presented as mean ± SEM (n = 3). p < 0.05 compared to IL‐1β‐treated group, # p < 0.05 compared to untreated control.
Discussion
4
Despite extensive research efforts, no effective therapy currently exits that can halt or reverse the underlying pathology of OA. Long‐term use of conventional analgesics is associated with adverse effects and an increased risk of complications. This has driven growing interest in natural bioactive compounds derived from marine sources, such as sulfated polysaccharides [14]. These polysaccharides are natural polymers with promising potential in cartilage regeneration due to their structural resemblance to the physicochemical properties of the cartilage extracellular matrix (ECM) [37]. G. fisheri contains SG, a class of sulfated polysaccharides structurally similar to heparan sulfate proteoglycans, key ECM molecules involved in cell adhesion, growth factor signaling, and matrix organization [17]. SG has demonstrated beneficial bioactivities, including stimulation of collagen production in fibroblast cells through upregulation of COL1A1 expression at both the mRNA and protein levels, without cytotoxicity [18]. More recently, our study in C28/I2 human chondrocytes showed that SG from G. fisheri promotes ECM synthesis via activation of the integrin‐β1/FAK/Akt signaling pathway [35]. However, its potential role in regulating cartilage matrix homeostasis has not yet been examined in experimental OA models.
The present study provides the first in vivo evidence that SG exerts chondroprotective effects in the ACLT + MMx rat model of OA. Weekly intra‐articular injections of SG attenuated cartilage degeneration, particularly at the highest dose (500 µg), as evidenced by preservation of cartilage structure and proteoglycan content. Importantly, these structural effects parallel the in vitro findings, where SG suppressed IL‐1β–induced expression of catabolic enzymes (MMP‐1 and MMP‐13) and enhanced anabolic and protective markers (TIMP‐1 and COL2A1) in human chondrocytes. Together, these effects suggest that SG shifts the balance of ECM regulation toward cartilage preservation by inhibiting degradation and promoting anabolic processes. This dual anti‐catabolic and pro‐anabolic action provides a plausible mechanism for the cartilage‐preserving effects of SG observed in vivo.
Previous reports indicate that repeated intra‐articular delivery of high doses of candidate compounds exert stronger protective effects than single administrations [38, 39]. For instance, intra‐articular delivery of a high concentration of glucosamine hydrochloride (350 μg) for three consecutive weeks significantly suppressed OA progression in a surgery‐induced OA model, suggesting disease‐modifying potential [39]. Similarly, Chen et al demonstrated that repeated weekly intra‐articular injections of HA conferred chondroprotection in ACLT + MMx‐induced OA rats [38]. Consistent with these findings, our study showed that repeated intra‐articular SG injections (500 μg) over four consecutive weeks exerted a potential disease‐modifying effect. In contrast, we and did not observe a significant reduction in mean OARSI score following HA treatment. This discrepancy may reflect differences in treatment timing and regimen. While our study initiated intra‐articular injections of HA at 4 weeks post‐surgery, Chen et al. [38] administered HA earlier (2 weeks post‐surgery) and employed two distinct treatment periods (weeks 2–4 and 9–11).
Interestingly, despite attenuating cartilage degeneration, SG treatment did not significantly restore type II collagen levels in vivo. This may reflect the complexity of the joint environment, where persistent mechanical stress, inflammatory cytokines, or rapid matrix turnover could limit the translation of in vitro anabolic effects. Furthermore, the 4‐week duration of SG treatment may have been insufficient to induce detectable increases in mature type II collagen within the cartilage matrix, whereas in vitro conditions provide a controlled environment conducive to collagen synthesis [40]. Future studies examining longer treatment durations, optimized dosing, or combination strategies with hyaluronic acid (HA) or platelet‐rich plasma (PRP) may better enhance collagen restoration.
Notably, SG treatment did not produce measurable improvements in pain‐related behavior, as assessed by weight‐bearing asymmetry, despite protecting cartilage structure. This structural symptom disconnect between structural preservation and persistent pain is well recognized in OA research and likely reflects the multifactorial nature of OA pathophysiology, where nociceptive pathways may remain active despite partial tissue protection [39, 40]. Similar dissociation has been reported with other intra‐articular interventions; for example, fucoidan and polysaccharides from the brown seaweed Sargassum, when administered orally, demonstrated cartilage‐protective and anti‐inflammatory effects without consistent analgesic benefit in OA models [15, 16]. In our study, both SG and HA were administered intra‐articularly, and neither produced significant analgesic effects, consistent with previous reports showing limited antinociceptive effects of intra‐articular HA in rat OA pain models [38, 41, 42].
The study has several limitations concerning the treatment duration, the assessment of symptomatic relief, and the potential oversight of sex differences. First, the 4‐week treatment duration may be insufficient to fully capture the SG' anabolic potential; this is evidenced by the failure to observe a significant in vivo restoration of type II collagen despite strong in vitro results, suggesting that a longer period is necessary for the deposition and maturation of new collagen within the complex cartilage matrix. This likely masked a true, slower‐acting anabolic effect. Second, a significant structural‐symptom disconnect was observed, where cartilage preservation occurred without corresponding pain relief. This limitation is compounded by the reliance on weight‐bearing asymmetry, a standard but limited measure, which may not fully capture the multifactorial nature of chronic OA pain, thus restricting the study's translational relevance. Future studies should, therefore, incorporate a variety of pain assessments to better represent the true symptomatic burden of the disease. Finally, the potential oversight of sex as a biological variable is a critical constraint, as OA prevalence and severity differ between sexes in humans; without incorporating both male and female animals and conducting sex‐disaggregated analysis, the generalizability of the results and the potential for sex‐specific therapeutic effects are limited.
Collectively, these findings demonstrate that the SG primarily act as a cartilage‐preserving scaffold rather than a direct analgesic. While SG effectively attenuated structural degeneration, the lack of corresponding pain relief suggests that its potential for symptomatic benefit is complex and likely depends on factors such as the specific animal model types, route of administration, and dosage regimen. Our results thus underscore the need for further optimization of the therapeutic approach to achieve both the observed structural benefits and symptomatic relief in OA.
Conclusion
5
This study provides the first evidence that SG from Gracilaria fisheri exerts chondroprotective effects in an experimental OA model by attenuating cartilage degeneration and modulating key catabolic and anabolic genes. Although SG treatment did not alleviate pain‐related behaviors within the timeframe of this study, its ability to preserve cartilage structure highlights its potential as a candidate for disease‐modifying interventions. Optimizing treatment parameters, including dose, frequency, and duration, will be essential to enhance type II collagen restoration in vivo and to translate the in vitro protective effects into therapeutic outcomes. Overall, our findings support further investigation of SG as a natural, locally abundant cartilage‐preserving candidate in the spectrum of disease‐modifying strategies for OA.
Author Contributions
Nirada Srianake: investigation, formal analysis, visualization, writing – original draft preparation. Amarin Thongsuk: investigation, formal analysis, visualization, writing – original draft preparation. Scarlett Desclaux, Thitiya Phuagpan and Jiraporn Sriwong: investigation, methodology. Alita Kongchanagul: supervision, conceptualization, research designs, methodology, formal analysis, and writing – review and editing of the manuscript. Wanwisa Himakhun: supervision, conceptualization, research designs, methodology, formal analysis, writing – review and editing of the manuscript. Ruedee Hemstapat: supervision, conceptualization, research designs, methodology, formal analysis, validation, writing – original draft, and writing – review and editing of the manuscript. Kanokpan Wongprasert: supervision, conceptualization, research designs, methodology, funding acquisition, validation, writing – original draft, and writing – review and editing of the manuscript. All authors have read, and approved the final submitted manuscript.
Conflicts of Interest
The authors declare no conflicts of interest.
Supporting information
Supplementary_material_5Oct225_Final.
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