HNE is of specific interest in the present study. Its formation is enhanced in articular tissues, including cartilage and synovial membranes from OA patients ( 13 , 20 ). It has been suggested that HNE generated inside the joints contributes significantly to OA pathogenesis, since inorganic oxidants can modify ECM components and, consequently, accelerate their degradation ( 13 , 14 ). Furthermore, the relevance of HNE to OA pathogenesis has been demonstrated by its ability to activate MMP‐13 via adduct formation without proteolytic cleavage ( 13 ). Oxidative modification of cartilage collagen by HNE in vivo could alter the biochemical and biophysical properties of cartilage collagen fibrils, making them prone to degradation and initiating the changes observed in aging and OA. Interestingly, modification of type II collagen (CII) by HNE affects chondrocyte–matrix interaction and, in turn, evokes multiple abnormalities of chondrocyte phenotype and function, denoting its contribution in OA ( 21 ). Other mediators related to cartilage damage are also up‐regulated by HNE in human OA chondrocytes, including cyclooxygenase 2 (COX‐2) and 5‐lipoxygenase ( 22 ).

Articular cartilage is predominantly made up of molecules with a long biologic half‐life, which, compared to other tissues, makes it more sensitive to increased oxidative stress–related molecules, such as lipid peroxidation products ( 13 , 14 ), advanced glycation end products (AGEs) ( 15 ), and nitrosylated proteins ( 16 ). The oxidative stress–related molecule 4‐hydroxynonenal (HNE) has generated considerable research interest in the last 10 years ( 17 ). It is an aldehyde end product generated by peroxidation of the most abundant class of ω‐3 and ω‐6 polyunsaturated fatty acids ( 18 ). Similar to free radicals, HNE is an electrophile that reacts readily to nucleophilic residues of proteins, nucleic acids, and lipids, but its relatively long half‐life makes it a candidate for damage propagation to neighboring cells. Interest in HNE stems not only from its potential use as a biomarker of oxidative stress–induced lipid peroxidation, but also because of accumulating evidence indicating that it modulates signaling pathways involved in cell proliferation, apoptosis, and inflammation ( 19 ).

Oxidative stress plays a crucial role in maintaining and sustaining cartilage degradation. In articular chondrocytes, evidence implicates reactive oxygen species (ROS) as signaling intermediates of IL‐1β ( 6 , 7 ). It has been suggested that ROS produced inside the joints may contribute significantly to the pathogenesis of OA, since these inorganic oxidants are able to degrade cartilage via oxidation of extracellular matrix (ECM) components or posttranslational modification of matrix metalloproteinases (MMPs) ( 8 , 9 ). The modification of MMPs by ROS results in their activation by the “cysteine switch” mechanism, with or without further proteolysis ( 8 , 10 ). Furthermore, ROS may shift the balance between MMPs and tissue inhibitors of metalloproteinases (TIMPs) by augmenting MMP production or by inactivating TIMPs ( 11 , 12 ).

Osteoarthritis (OA) is an extremely common joint disorder in Western populations. Its high prevalence, especially in the elderly, and the frequency of OA‐related physical damage make it one of the leading causes of disability. This prevalent and crippling chronic condition affecting the elderly poses a significant public health problem. The most prominent feature of OA is the progressive destruction of articular cartilage, resulting in impaired joint motion, severe pain, and, ultimately, disability. One of the foremost characteristics of OA is damage to the collagen network, as reflected by increased tissue swelling and loss of proteoglycans ( 1 , 2 ). Both collagen damage and loss of proteoglycans adversely affect the mechanical properties of cartilage. Chondrocytes respond to tissue injury by increasing proteoglycan and collagen synthesis in an attempt to repair it ( 3 ). If repair fails, damage will progress to articular cartilage degeneration. These processes are thought to be largely elicited through excess production of proinflammatory and catabolic mediators. Among them, interleukin‐1β (IL‐1β) has been demonstrated to be predominantly involved in disease initiation and progression ( 4 , 5 ).

HNE–CII and HNE–MMP‐13 conjugates were analyzed, as described previously ( 13 ). Cartilage explants (∼50 mg) were homogenized on ice in 1 ml of RIPA buffer supplemented with protease inhibitors and 2 M guanidine hydrochloride. Extracts were dialyzed overnight against RIPA buffer at 4°C. One hundred micrograms of tissue protein was subjected to immunoprecipitation with mouse anti‐human CII (1:500 dilution; EMD Millipore) or anti‐human MMP‐13 antibody (1:500 dilution; R&D Systems) in RIPA buffer overnight at 4°C and then for an additional 2 hours with protein G (Sigma‐Aldrich) ( 13 ). After brief centrifugation, the resin was washed with RIPA buffer, and proteins were removed by the addition of 100 μl of undiluted Laemmli sample buffer (Bio‐Rad). The immunoprecipitates were analyzed by Western blotting with rabbit anti‐HNE (1:1,000 dilution; EMD Millipore), mouse anti–MMP‐13 (1:1,000 dilution; R&D Systems), or mouse anti‐CII (1:1,000 dilution; EMD Millipore) as primary antibody.

Twenty micrograms of total proteins from synovium or articular cartilage extracts were loaded for discontinuous 4–12% sodium dodecyl sulfate–polyacrylamide gel electrophoresis. They were then transferred electrophoretically onto nitrocellulose membranes (Bio‐Rad) for protein immunodetection and semiquantitative measurement ( 13 ). The primary antibodies were rabbit anti‐HNE (EMD Millipore), mouse antinitrotyrosine (Santa Cruz Biotechnology), rabbit anti–COX‐2 (Cayman Chemical), mouse anti‐CII (EMD Millipore), mouse anti–MMP‐13 (R&D Systems), and mouse anti–β‐actin (Sigma‐Aldrich). After serial washes, primary antibodies were revealed by goat anti‐mouse or anti‐rabbit IgG conjugated to horseradish peroxidase (Cell Signaling Technology). Immunoreactive proteins were detected with SuperSignal blotting substrate (Pierce) and exposed to X‐Omat film (Eastman Kodak).

Each slide was washed 3 times in PBS (pH 7.4), stained by the avidin–biotin complex method (Vectastain ABC kit), and incubated in the presence of biotin‐conjugated goat anti‐mouse IgG antibody (Cell Signaling Technology) for 45 minutes at room temperature, followed by the addition of avidin–biotin–peroxidase complex for 45 minutes. All incubations were performed in a humidified chamber at room temperature, with subsequent staining by 3,3′‐diaminobenzidine (Dako) containing H 2 O 2 . Slides were counterstained with hematoxylin and eosin. Staining specificity was determined in sham‐operated and OA tissues by substitution of the primary antibody with autologous preimmune sera. Each section was examined and photographed under a light microscope (Leica). Positive staining of chondrocytes was evaluated in the central region of cartilage samples from dogs. Six microscopic fields in each specimen were examined at 40× magnification. The total number of chondrocytes and the number of chondrocytes staining positive were evaluated, and results were expressed as the percentage of chondrocytes staining positive.

Cartilage samples from weight‐bearing areas of the tibial plateaus of dogs were processed for immunohistochemistry. They were fixed in TissuFix #2 for 24 hours, then embedded in paraffin. A total of 3–6 paraffin‐embedded sections (5 μm) were deparaffinized in toluene, rehydrated in reverse‐graded ethanol series, and preincubated with 0.25 units/ml chondroitinase ABC (Sigma‐Aldrich) in phosphate buffered saline (PBS) (pH 8.0) for 60 minutes at 37°C. They were subsequently washed in PBS, incubated in 0.3% Triton X‐100 for 20 minutes, then placed in 3% H 2 O 2 /PBS for 15 minutes. The slides were further incubated with blocking serum (Vectastain ABC kit; Vector) for 60 minutes, after which they were blotted and overlaid with mouse anti–MMP‐13 and anti–ADAMTS‐5 antibody (R&D Systems) for 18 hours at 4°C in a humidified chamber.

Full‐thickness cartilage sections were removed from weight‐bearing lesional areas of the femoral condyles and tibial plateaus, allowing sampling standardization as per OARSI guidelines ( 24 ). Sagittal sections of cartilage from each femoral condyle and tibial plateau specimen were evaluated histologically, as described previously ( 25 ). They were dissected, fixed in TissuFix #2 (Laboratoires Gilles Chaput), and embedded in paraffin. Serial sections (5 μm) were stained with Safranin O. The severity of cartilage pathology was graded by 2 independent observers (JCF, MB) according to the OARSI histopathology scoring system ( 24 ). This scale served to assess OA lesion severity on the basis of cartilage structure (0–12 scale), cellular changes (0–12 scale), proteoglycan staining (0–12 scale), and collagen integrity (0–9 scale).

The right knee of each dog was dissected on ice immediately after the dog was killed. Each knee was examined for gross morphologic changes, as described previously ( 23 ), by 2 independent observers (JCF, MB) who were blinded to the treatment groups. Macroscopic cartilage lesions (mm 2 ) were measured and graded (0–4 scale) according to the Osteoarthritis Research Society International (OARSI) scoring system ( 24 ). Overall scores were obtained for femoral condyles and tibial plateaus separately by calculating the mean cartilage lesion score of lateral and medial regions.

Synovial fluid was collected at the time that dogs were killed by inserting a needle with syringe into the joint space; synovial fluid was then stored at −80°C. Knee synovial tissue was collected and immediately placed on ice and transported to the laboratory. Synovial membrane fragments (4–5 mm 2 ) were excised with a fine scalpel, placed in microcentrifuge tubes, and frozen in liquid nitrogen. Samples were pulverized in a liquid nitrogen–chilled Plattner's mortar (Fisher Scientific) and stored in powder form at −80°C. An aliquot of pulverized tissue was weighed and homogenized in 1 ml of radioimmunoprecipitation assay (RIPA) buffer (150 m M NaCl, 1 m M EDTA, 40 m M Tris, pH 7.6, 1% Triton X‐100) supplemented with protease inhibitors. After incubating the mixture for at least 10 minutes on ice, it was transferred to a microcentrifuge tube and centrifuged at 10,000 g for 10 minutes at 4°C. The resulting supernatant was divided into aliquots and stored at −80°C for later analysis. The total protein concentration in extracts was determined with a bicinchoninic acid protein assay kit (Pierce).

Twelve dogs received a weekly intraarticular injection of 0.3 ml saline solution (group 6, n = 6) or saline solution containing 80 nmoles/ml free HNE (Cayman Chemical) (group 7, n = 6). This concentration corresponds to that found in vivo in synovial fluid from dogs that have undergone ACLT. The dogs were housed for 8 weeks and then euthanized. All dogs (in both protocols) were actively exercised in exterior runs for 1 hour each day 5 days a week, under the supervision of an animal care technician.

Thirty dogs were used in this protocol. OA was induced in 24 dogs by right knee ACLT through a stab wound after joint capsule opening under general sodium pentobarbital anesthesia (25 mg/kg) ( 23 ). All 30 dogs received a preemptive analgesic (buprenorphine, 0.2 mg/kg) subcutaneously. During the postoperative period, they were treated with the same analgesic, if necessary. Dogs were randomly assigned to 5 treatment groups. Group 1 (sham operated, n = 6) received daily placebo (encapsulated methylcellulose) starting on day 1. Group 2 (ACLT surgery, n = 6) received daily placebo starting on day 1. Groups 3 and 4 (ACLT surgery, n = 6) received carnosine (Sigma‐Aldrich) orally at 5 and 20 mg/kg/day, respectively, starting on day 1. Group 5 (ACLT surgery, n = 6) received carnosine orally at 20 mg/kg/day, starting 4 weeks after surgery. Treatment was continued until the dogs were euthanized 8 weeks after surgery.

We studied healthy adult crossbred dogs (ages 2–3 years) weighing 22–25 kg. Two protocols, conducted in accordance with Canadian Council on Animal Care guidelines, were approved by the institutional ethics committee. The dogs were housed individually in galvanized steel cages (1 meter in width × 1.5 meters in length × 2 meters in height), each separated by a panel. They were selected after complete physical and musculoskeletal evaluation by a veterinarian who declared them to be healthy.

To confirm the formation of HNE–MMP‐13 and HNE–CII adducts, cartilage extracts were immunoprecipitated with rabbit anti‐human MMP‐13 and mouse anti‐human CII antibody, then subjected to Western blotting. With anti‐HNE antibody, we found signals corresponding to HNE–MMP‐13 and HNE–CII conjugates (Figure 5 D). Treatment with 20 mg/kg/day carnosine prevented the formation of these conjugates, reduced MMP‐13 levels, and normalized CII levels in articular cartilage from dogs with OA. These data confirm our expectations, namely, that MMP‐13 and CII are targets of HNE binding in OA cartilage and that carnosine prevents their modification by HNE.

To verify that articular cartilage is a target of HNE binding and nitrosylation, we first analyzed HNE–protein adducts and nitrosylated proteins extracted by Western blotting. Corresponding profiles of HNE–protein adducts and nitrosylated proteins extracted from cartilage explants are illustrated in Figure 5 C. The intensity of immunoreactive bands decreased in cartilage from dogs with OA treated with carnosine compared to placebo‐treated dogs with OA, and the molecular weight of most bands was ≥40 kd. Tissue extracts from sham‐operated placebo‐treated dogs showed bands with very low immunoreactivity corresponding to HNE–protein adducts and nitrosylated proteins. To ensure that anti‐HNE antibody recognized the targeted proteins, HNE–BSA adducts served as positive controls (Figure 5 C).

Carnosine administration suppresses the lipid peroxidation products HNE, pentosidine, and nitrosylated proteins as well as the formation of HNE–MMP‐13 and HNE–type II collagen (CII [Col II]) adducts. A, HNE–protein adducts were assessed in synovial membrane explants using an in‐house enzyme‐linked immunosorbent assay (ELISA). B, HNE–protein adducts and pentosidine levels were assessed in synovial fluid using an in‐house ELISA and a commercial kit, respectively. C, Cartilage explants from placebo‐ and carnosine‐treated dogs that had undergone anterior cruciate ligament transection were homogenized as described in Materials and Methods , and total HNE–protein adducts were analyzed by Western blotting using anti‐HNE and antinitrotyrosine antibodies. D, Cartilage extracts (100 μg) were immunoprecipitated using anti–MMP‐13 and anti‐CII antibodies and subjected to Western blotting using an anti‐HNE antibody. Total MMP‐13 and CII levels were revealed by Western blotting using anti–MMP‐13 and anti‐CII antibodies. In C and D, samples are from sham‐operated placebo‐treated dogs (lane 1), dogs with OA treated with placebo (lane 2), dogs with OA treated with 5 mg/kg/day carnosine (lane 3), and dogs with OA treated with 20 mg/kg/day carnosine (lane 4). Values are the mean ± SEM of 6 experiments. ## = P < 0.01; ### = P < 0.001 versus sham‐operated dogs, by Student's unpaired t ‐test. BSA = bovine serum albumin; COX‐2 = cyclooxygenase 2 (see Figure 4 for other definitions).

This part of our study was designed to investigate the protective effect of carnosine against oxidative stress–induced protein modifications in articular tissues from dogs with OA. Our findings, which are shown in Figure 5 A, revealed that oral administration of 20 mg/kg/day carnosine reduced HNE–protein adducts in synovial membrane 5.8‐fold ( P < 0.01). In synovial fluid from carnosine‐treated dogs, we showed that concentrations of HNE–protein adducts and pentosidine (an AGE) were decreased 3.2‐fold ( P < 0.01) and 1.8‐fold ( P < 0.05), respectively, compared to concentrations in synovial fluid from placebo‐treated dogs with OA (Figure 5 B). Very low levels of both HNE and pentosidine were detected in synovial fluid from sham‐operated placebo‐treated dogs.

A and B, Representative sections of superficial zones of articular cartilage from placebo (P)– and carnosine (CAR)–treated dogs 8 weeks after sham surgery or anterior cruciate ligament transection ( A ) and after 8 weeks of intraarticular injections of vehicle or 4‐hydroxynonenal (HNE) ( B ), showing immunostaining for matrix metalloproteinase 13 (MMP‐13) and ADAMTS‐5. Original magnification × 40. Immunohistochemistry data are presented as the mean ± SEM. ## = P < 0.01 versus sham‐operated dogs; ∗ = P < 0.05; ∗∗ = P < 0.01 versus vehicle, by Mann‐Whitney 2‐tailed U test. OA = osteoarthritis.

In the next set of experiments, we investigated the articular cartilage expression of MMP‐13 and ADAMTS‐5, principal proteases involved in CII and aggrecan degradation, respectively. Cartilage immunohistochemistry showed a significant decrease in the percentage of positively stained cells in dogs with OA treated with 20 mg/kg/day carnosine compared to untreated controls for MMP‐13 (mean ± SEM 23 ± 4.6% versus 72.5 ± 15.2%; P < 0.05) and ADAMTS‐5 (mean ± SEM 29.5 ± 6.1% versus 82.1 ± 18%; P < 0.05) (Figure 4 A). Furthermore, as illustrated in Figure 4 B, immunohistochemical analysis of cartilage disclosed a significant increase in MMP‐13 and ADAMTS‐5 levels in dogs receiving intraarticular HNE injections as compared to controls (for MMP‐13, 52 ± 10.3% versus 11.7 ± 1.4%; P < 0.05) (for ADAMTS‐5, 84 ± 17.3% versus 7.2 ± 0.1%; P < 0.01). Negative controls for MMP‐13 and ADAMTS‐5 presented only background staining.

Histologic assessment of osteoarthritic (OA) articular cartilage from tibial plateaus. A and B, Representative Safranin O–stained sections of cartilage from placebo (P)– and carnosine (CAR)–treated dogs 8 weeks after sham surgery or anterior cruciate ligament transection ( A ) and after 8 weeks of intraarticular injections of vehicle or 4‐hydroxynonenal (HNE) ( B ). Original magnification × 40. Histologic OA scores of the medial tibial plateaus are presented as the mean ± SEM. ∗ = P < 0.05 versus vehicle, by Mann‐Whitney 2‐tailed U test.

Figure 3 depicts representative sections of cartilage from the experimental ACLT dog model of OA with or without carnosine treatment 8 weeks after surgery. Histologically, articular cartilage from control (sham‐operated) dogs appeared to be normal (Figure 3 A). Eight weeks after surgery, histologic changes were evident in loss of surface integrity, hypocellularity, and fibrillations (Figure 3 A). These alterations were significantly reduced in animals treated with carnosine for 8 weeks ( P < 0.05). On the other hand, the severity of cartilage lesions on tibial plateaus (Figure 3 B) was also found to be significantly greater in the group receiving intraarticular injections of HNE for 8 weeks compared to vehicle ( P < 0.05).

This part of our experiments was performed to determine whether reduction of cartilage lesions correlated with the presence of carnosine in synovial fluid. Our findings revealed that the concentration of carnosine reached 23 ± 6.4 ng/ml (mean ± SEM) in dogs with OA treated with 20 mg/kg/day. However, synovial fluid from placebo‐treated sham‐operated dogs and placebo‐treated dogs with OA contained undetectable concentrations of carnosine. In addition, there were no significant differences in serum pH between all animal groups (data not shown).

A, Shown is macroscopic appearance of osteoarthritic articular cartilage from femoral condyles and tibial plateaus. Representative photographs of cartilage from dogs receiving intraarticular injections of vehicle or 4‐hydroxynonenal (HNE) for 8 weeks show erosion and pitting (circled areas). B, Synovial membrane explants from vehicle‐ and HNE‐treated dogs were homogenized as described in Materials and Methods , and cyclooxygenase 2 (COX‐2) protein expression was analyzed by Western blotting using anti–COX‐2 antibody. C, Prostaglandin E 2 (PGE 2 ) in synovial fluid was measured with a commercial kit. Values are the mean ± SEM of 6 experiments. ∗∗ = P < 0.01 versus vehicle, by Student's unpaired t ‐test.

We conducted additional experiments to obtain direct evidence of HNE involvement in the OA process. A pathophysiologic dose of HNE (80 nmoles/ml), corresponding to that found in vivo in synovial fluid from dogs that have undergone ACLT, was injected weekly for 8 weeks into knee joints. As illustrated in Figure 2 and Table 1 , compared to the vehicle‐treated group, significant increases in lesion surface and depth on femoral condyles and tibial plateaus were seen when HNE was injected into knee joints ( P < 0.01). Moreover, the macroscopic lesions for tibial plateaus in the ACLT model were more severe and significantly different from those in the HNE model (Table 1 ). In synovial membrane and synovial fluid, the expression levels of COX‐2 and PGE 2 , respectively, were higher in HNE‐treated animals than in vehicle‐treated animals (Figures 2 B and C). To our knowledge, these data are the first in the literature to show that HNE is involved in the OA process in vivo.

To test the efficacy of carnosine in a model of preexisting OA, 20 mg/kg/day carnosine was administered orally to dogs with OA starting 4 weeks after surgery. The severity of macroscopic lesions on femoral condyles and tibial plateaus of these carnosine‐treated dogs was significantly different from that observed in placebo‐treated controls with OA ( P < 0.05). Taken together, these findings disclosed, in an in vivo animal model, that carnosine reduces the progression and initiation of structural OA changes.

Lesions were assessed for size and depth in all groups. Minimal macroscopic cartilage lesions were found in the control (sham‐operated, placebo‐treated) group (Figure 1 and Table 1 ). Macroscopic lesions in all carnosine‐treated groups with OA were decreased in size and depth on both femoral condyles and tibial plateaus compared to the placebo‐treated group with OA (Figure 1 and Table 1 ). A significant reduction in lesion surface was noted on tibial plateaus ( P < 0.05) when dogs were treated with 5 mg/kg/day of carnosine and on both femoral condyles and tibial plateaus ( P < 0.01) when animals were treated with 20 mg/kg/day of carnosine. A significant decrease in lesion depth was noted on femoral condyles at 5 and 20 mg/kg/day of carnosine ( P < 0.05 and P < 0.01, respectively). However, macroscopic cartilage analysis demonstrated no difference between dogs with OA treated with 20 mg/kg/day carnosine and placebo‐treated sham‐operated controls.

DISCUSSION

The principal finding of the present study was that the lipid peroxidation end product HNE could be involved in OA progression. First, we demonstrated increased HNE in synovial fluid, and we showed that the initiation and progression of cartilage lesions in the experimental dog model of OA induced by ACLT were reduced with HNE‐trapping carnosine. Second, we observed that intraarticular injection of HNE into the knee joints of dogs induced cartilage lesions and expression of MMP‐13, ADAMTS‐5, and COX‐2. The injected concentration of HNE corresponded to that observed in synovial fluid in the dog model of ACLT‐induced OA. Our macroscopic and histologic findings were informative with respect to the effects of carnosine and HNE on cartilage structure in OA. In addition, carnosine treatment was found to reduce levels of HNE, AGEs, nitrosylated proteins, MMP‐13, ADAMTS‐5, and COX‐2. The strength of our study is that, to our knowledge, it is the first to research the involvement of HNE in the context of OA pathophysiology. In addition, our observations support the notion that dietary carnosine intake could decrease the formation of cartilage lesions.

In the present investigation, we confirmed a significant increase in endogenous HNE as well as in AGEs and nitrosylated proteins in articular tissues, including synovial membrane, synovial fluid, and articular cartilage, from dogs with ACLT‐induced OA compared to controls. In our previous work, we reported increased HNE in synovial fluid from OA patients and in isolated OA chondrocytes incubated with tumor necrosis factor α (TNFα) or free radical donors (13). Our data directly implicate these mediators in HNE production in articular tissues, most likely via ROS generation. Increased HNE in OA was also discovered by Grigolo et al (20) and Shah et al (14), who found higher HNE and malondialdehyde (MDA) levels in human OA synovium and cartilage tissues than in normal control tissues.

Accumulation of HNE implicates its involvement in OA, but factors promoting increased HNE–protein adducts in OA remain to be clarified. They include the inactivation of antioxidant enzymes and alteration of glutathione (GSH) redox status. In addition to increased ROS production, the intrinsic capacity of cells to metabolize HNE, whether free or bound to nontoxic products, is another factor that should be considered. This can occur through reduction by aldose reductase to 1,4‐dihydroxynonenol, oxidation to 1,4‐dihydroxynonenoic acid by aldehyde dehydrogenase, and conjugation to GSH by glutathione S‐transferase A4 (26). Our early data showed decreased glutathione S‐transferase A4 expression in human and canine OA cartilage (Abusarah J, Fernandes JC, Fahmi H, Benderdour M: unpublished observations). One could speculate that if glutathione S‐transferase A4 down‐regulation is involved in HNE accumulation in articular cartilage, then it is likely to be crucial in OA pathogenesis. In cells, HNE is predominantly metabolized by this enzyme via its conjugation to GSH (27).

HNE is considered to be the most plentiful and reactive byproduct of lipid peroxidation, exerting powerful biologic effects in various cell and tissue systems (28). In the present study, our data indicate a link between HNE burden and OA pathogenesis. At the transcriptional level, exposure of human chondrocytes to nontoxic doses of HNE elicits a series of catabolic and inflammatory genes involved in cartilage degradation (13, 29). With regard to bone metabolism, we found that OA osteoblasts produced high levels of HNE compared to osteoblasts from normal controls (30). We also determined that HNE exerts multiple effects on human OA osteoblasts by selective activation of signal transduction pathways and alteration of osteoblastic phenotype expression through inhibition of alkaline phosphatase activity and expression. Similar to HNE, accumulation of AGEs in cartilage stimulates different kinds of genes, such as COX‐2 and MMPs, via activation of the NF‐κB and p38 MAPK signaling pathways, mediated by specific AGE receptors on the chondrocyte surface.

Macromolecule modification by HNE has received considerable attention. Of particular relevance to this work is adduction of the very enzyme responsible for cartilage degradation, namely, MMP‐13. Our immunoprecipitation data indicated increased HNE–MMP‐13 adducts in OA cartilage. We previously reported the relevance of MMP‐13 modification by HNE when we demonstrated that HNE binding to recombinant human MMP‐13 results in its activation without proteolytic cleavage (13). Given the chemical reactivity of HNE to cysteine residues and the known molecular structure of MMP‐13, we speculate that HNE activates MMP‐13 by disrupting Zn2+–cysteine interactions, a mechanism known as the “cysteine switch” (31). In a similar mechanism, it has been reported that nitric oxide activates MMP‐9 by S‐nitrosylation in mouse neuronal cells (32).

Continuing within the context of macromolecule modification by HNE, we assessed the formation of HNE–CII adducts in OA canine cartilage compared to normal control cartilage. Our results are consistent with findings indicating increased HNE–CII adducts in cartilage in response to TNFα and free radical donors (13). In a study conducted in isolated rabbit chondrocytes, Tiku et al (33) demonstrated that MDA also mediates CII modification. The consequences of CII modification by HNE have been investigated in our laboratory (13). The results showed that HNE‐modified CII is vulnerable to degradation by MMP‐13 action compared to digestion of unmodified CII. Similar to aldehydes, other nucleophilic agents such as ROS mediate cartilage degradation via oxidative damage of cartilage components (34). Collagen modification most likely represents one of the main mechanisms by which ROS can influence physiologic and pathologic processes. On the other hand, the significance of high HNE–CII adduct levels in OA cartilage was clarified by our research group. We established that interactions between OA chondrocytes and HNE‐modified CII affect cell phenotype and induce catabolic and inflammatory factors, indicating their contribution to cartilage damage seen during OA development (21).

In the present study, we tested our hypothesis that carnosine prevents cartilage damage. Carnosine has recently attracted much attention as a naturally occurring antioxidant and antiglycating agent. We demonstrate that carnosine, given prophylactically or for preexisting OA, is able to reduce cartilage damage most likely through its distinctive combination of antioxidant and antiglycating properties. These findings may suggest the involvement of HNE as well as glycation and nitrosylation in the process of OA. In keeping with the findings of our study, several recent studies have indicated that carnosine abolishes the generation of HNE, AGEs, and nitrosylated proteins (35, 36). HNE is of particular interest because of its high chemical reactivity, as evidenced by its production of such varied biologic effects (19). The chemical characteristics of HNE revealed that, compared with other aldehydes, it can also react with reduced GSH at much higher rates that are further accelerated by glutathione S‐transferase A4 (27, 37). Indeed, its reaction with carnosine may be a second line of defense under conditions of GSH depletion. The fact that the administration of exogenous carnosine prevents HNE production as well as inflammatory and catabolic factors suggests that despite its slow reactivity, carnosine offers significant protection against HNE toxicity and its biologic effects, even in GSH‐repleted cells and tissues.

That oral intake of carnosine reduces MMP‐13, ADAMTS‐5, and COX‐2 expression in cartilage and synovium explants from dogs with ACLT‐induced OA is further supported by data showing that this drug inhibits catabolic and inflammatory mediators, such as MMP‐13, COX‐2, PGE 2 , and nitric oxide, in human chondrocytes and macrophages (21, 38, 39). Carnosine also diminished the release of proinflammatory cytokines, including IL‐1β, IL‐6, and TNFα, in a mouse model of spinal cord injury (40). Our observations revealed that carnosine can reduce the initiation and progression of OA structural changes by removing HNE from the knee joint and attenuating the expression of factors known to be involved in cartilage degradation. These findings imply that HNE is a significant contributor to OA cartilage degradation.

Several studies have demonstrated that carnosine is safe, well‐tolerated, and commonly considered as a dietary supplement (41). It is a natural, water‐soluble antioxidant that suppresses ROS generation and scavenges lipid peroxidation products in human HCT116 colon cancer cells and apolipoprotein E–null mice (35, 42). Carnosine occurs naturally in several tissues but decreases with age and oxidative stress load. Its levels in the body decline 63% from age 10 years to age 70 years (43). Considerable evidence indicates that it has protective properties in several diseases, including atherosclerosis (35), cardiomyopathy (44, 45), and osteoporosis (46-48). To date, there are no reported studies concerning the in vivo beneficial effects of carnosine in OA. Indeed, Tallon et al (49) reported that carnosine levels in type II muscle fibers are reduced by a mean of 53.2% in elderly subjects with OA. In another recent investigation, Ohsawa and colleagues (50) provided novel, interesting information on the beneficial effect of carnosine in a mouse model of inflammation‐induced nociceptive pain. Those authors determined that it has antinociceptive actions on inflammatory pain, probably mediated by the attenuation of nociceptive sensitization in the spinal cord. Although the influence of carnosine on pain is unknown, we speculate that its effects may be attributed to HNE removal. This hypothesis is based on findings that HNE activates TRPA1 sensory ion channels on nociceptive neurons, promoting acute pain and neurogenic inflammation (51).

Our study has limitations. Its duration was relatively short, with only 1 dose of HNE being tested. However, care was taken to ensure that the dose selected was that found in synovial fluid in the ACLT model of OA. Another limitation concerned the time course of analysis. Eight weeks after the operation represents a relatively early stage of OA. Long‐term analysis is needed to clarify the role of HNE in more advanced stages of OA. Moreover, in‐depth analysis needs to be performed on the mode of action of carnosine on pathophysiologic OA pathways, with measurements of HNE–carnosine conjugates in dog urine.

In conclusion, this study strengthens the rationale for investigating the role of HNE in OA and for testing whether HNE‐trapping carnosine could attenuate the OA process. We observed promising outcomes with carnosine in the prevention of OA cartilage lesions induced by joint instability. Carnosine treatment seems to affect oxidative stress and the catabolic and inflammatory pathways of OA by inhibiting HNE production.