Recombinant human IL-1β was obtained from Genzyme (Cambridge, MA, USA). Cycloheximide (CHX) was purchased from Sigma-Aldrich Canada (Oakville, ON, Canada). SB203580, SP600125, PD98059, SN-50, and N-[N-(3,5-diflurophenylacetate)-L-alanyl]-(S)-phenylglycine t-butyl ester (DAPT) were from Calbiochem (now part of EMD Biosciences, Inc., San Diego, CA, USA). PGD₂ was from Cayman Chemical Company (Ann Arbor, MI, USA). Dulbecco's modified Eagle's medium (DMEM), penicillin and streptomycin, foetal calf serum (FCS), and TRIzol^® reagent were from Invitrogen (Burlington, ON, Canada). All other chemicals were purchased from either Bio-Rad Laboratories (Mississauga, ON, Canada) or Sigma-Aldrich Canada.

Healthy cartilage and synovial fluids were obtained at necropsy, within 12 hours of death, from donors with no history of arthritic diseases (n = 13, mean ± standard deviation [SD] age of 64 ± 17 years). To ensure that only healthy tissue was used, cartilage specimens were thoroughly examined both macroscopically and microscopically. OA cartilage and synovial fluids were obtained from patients undergoing total knee replacement (n = 32, mean ± SD age of 67 ± 16 years). All OA patients were diagnosed on criteria developed by the American College of Rheumatology Diagnostic Subcommittee for OA 28. At the time of surgery, the patients had symptomatic disease requiring medical treatment in the form of nonsteroidal anti-inflammatory drugs or selective COX-2 inhibitors. Patients who had received intra-articular injections of steroids were excluded. The Clinical Research Ethics Committee of Notre-Dame Hospital (Montreal, QC, Canada) approved the study protocol and the informed consent form.

Chondrocytes were released from cartilage by sequential enzymatic digestion as previously described 29. Briefly, this consisted of 2 mg/mL pronase for 1 hour followed by 1 mg/mL collagenase for 6 hours (type IV; Sigma-Aldrich Canada) at 37°C in DMEM and antibiotics (100 U/mL penicillin and 100 μg/mL streptomycin). The digested tissue was briefly centrifuged and the pellet was washed. The isolated chondrocytes were seeded at high density in tissue culture flasks and cultured in DMEM supplemented with 10% heat-inactivated FCS. At confluence, the chondrocytes were detached, seeded at high density, and allowed to grow in DMEM, supplemented as above. The culture medium was changed every second day, and 24 hours before the experiment, the cells were incubated in fresh medium containing 0.5% FCS. Only first-passaged chondrocytes were used.

Total RNA from homogenized cartilage or stimulated chondrocytes was isolated using the TRIzol^® reagent (Invitrogen) in accordance with the manufacturer's instructions. To remove contaminating DNA, isolated RNA was treated with RNase-free DNase I (Ambion, Inc., Austin, TX, USA). The RNA was quantitated using the RiboGreen RNA quantitation kit (Molecular Probes, Inc., now part of Invitrogen Corporation, Carlsbad, CA, USA), dissolved in diethylpyrocarbonate (DEPC)-treated H₂O, and stored at -80°C until use. One microgram of total RNA was reverse-transcribed using Moloney murine leukemia virus reverse transcriptase (RT) (Fermentas, Burlington, ON, Canada), as detailed in the manufacturer's guidelines. One fiftieth of the RT reaction was analyzed by real-time quantitative polymerase chain reaction (PCR) as described below. The following primers were used: L-PGDS [GeneBank: NM000954], sense 5'-AACCAGTGTGAGACCCGAAC-3', antisense 5'-AGGCGGTGAATTTCTCCTTT-3'; H-PGDS [GeneBank: NM014485], sense 5'-CCCCATTTTGGAAGTTGATG-3', antisense 5'-TGAGGCGCATTATACGTGAG-3; and glyceraldehyde-3-phosphate dehydrogenase (GAPDH) [GeneBank: NM002046], sense 5'-CAGAACATCATCCCTGCCTCT-3', antisense 5'-GCTTGACAAAGTGGTCGTTGAG-3'.

Quantitative PCR analysis was performed in a total volume of 50 μL containing template DNA, 200 nM of sense and antisense primers, 25 μL of SYBR^® Green master mix (Qiagen, Mississauga, ON, Canada), and uracil-N-glycosylase (UNG) (0.5 units; Epicentre Biotechnologies, Madison, WI, USA). After incubation at 50°C for 2 minutes (UNG reaction) and at 95°C for 10 minutes (UNG inactivation and activation of the AmpliTaq Gold enzyme; Qiagen), the mixtures were subjected to 40 amplification cycles (15 seconds at 95°C for denaturation and 1 minute for annealing and extension at 60°C). Incorporation of SYBR^® Green dye into PCR products was monitored in real time using a GeneAmp 5700 Sequence detection system (Applied Biosystems, Foster City, CA, USA), allowing the determination of the threshold cycle (C_T) at which exponential amplification of PCR products begins. After PCR, dissociation curves were generated with one peak, indicating the specificity of the amplification. A C_T value was obtained from each amplification curve using the software provided by the manufacturer (Applied Biosystems).

Relative amounts of mRNA in healthy and OA cartilage were determined using the standard curve method. Serial dilutions of internal standards (plasmids containing cDNA of target genes) were included in each PCR run, and standard curves for the target gene and for GAPDH were generated by linear regression using log (C_T) versus log (cDNA relative dilution). The C_T values were then converted to number of molecules. Relative mRNA expression in cultured chondrocytes was determined using the ΔΔC_T method, as detailed in the guidelines of the manufacturer (Applied Biosystems). A ΔC_T value was first calculated by subtracting the C_T value for the housekeeping gene GAPDH from the C_T value for each sample. A ΔΔC_T value was then calculated by subtracting the ΔC_T value of the control (unstimulated cells) from the ΔC_T value of each treatment. Fold changes compared with the control were then determined by raising 2 to the -ΔΔC_T power. Each PCR generated only the expected specific amplicon as shown by the melting-temperature profiles of the final product and by gel electrophoresis of test PCRs. Each PCR was performed in triplicate on two separate occasions for each independent experiment.

Cartilage specimens were processed for immunohistochemistry as previously described 29. The specimens were fixed in 4% paraformaldehyde and embedded in paraffin. Sections (5 μm) of paraffin-embedded specimens were deparaffinized in toluene and were dehydrated in a graded series of ethanol. The specimens were then preincubated with chondroitinase ABC (0.25 U/mL in phosphate-buffered saline [PBS] pH 8.0) for 60 minutes at 37°C, followed by a 30-minute incubation with Triton X-100 (0.3%) at room temperature. Slides were then washed in PBS followed by 2% hydrogen peroxide/methanol for 15 minutes. They were further incubated for 60 minutes with 2% healthy serum (Vector Laboratories, Burlingame, CA, USA) and overlaid with primary antibody for 18 hours at 4°C in a humidified chamber. The antibody was a rabbit polyclonal anti-human L-PGDS (United States Biological Inc., Swampscott, MA, USA), used at 10 μg/mL. Each slide was washed three times in PBS pH 7.4 and stained using the avidin-biotin complex method (Vectastain ABC kit; Vector Laboratories). The colour was developed with 3,3'-diaminobenzidine (DAB) (Vector Laboratories) containing hydrogen peroxide. The slides were counterstained with eosin. The specificity of staining was evaluated by using antibody that had been preadsorbed (1 hour at 37°C) with a 20-fold molar excess of recombinant human L-PGDS (Cayman Chemical Company) and by substituting the primary antibody with nonimmune rabbit IgG (Chemicon International, Temecula, CA, USA), used at the same concentration as the primary antibody. The evaluation of positive-staining chondrocytes was performed using our previously published method 29. For each specimen, six microscopic fields were examined under × 40 magnification. The total number of chondrocytes and the number of chondrocytes staining positive were evaluated, and the results were expressed as the percentage of chondrocytes staining positive (cell score).

Chondrocytes were lysed in ice-cold lysis buffer (50 mM Tris-HCl, pH 7.4, 150 mM NaCl, 2 mM EDTA [ethylenediaminetetraacetic acid], 1 mM PMSF [phenylmethylsulphonyl fluoride], 10 μg/mL each of aprotinin, leupeptin, and pepstatin, 1% NP-40, 1 mM Na₃VO₄, and 1 mM NaF). Lysates were sonicated on ice and centrifuged at 12,000 revolutions per minute for 15 minutes. The protein concentration of the supernatant was determined using the bicinchoninic acid method (Pierce, Rockford, IL, USA). Twenty micrograms of total cell lysate was subjected to SDS-PAGE and electrotransferred to a nitrocellulose membrane (Bio-Rad Laboratories). After blocking in 20 mM Tris-HCl pH 7.5 containing 150 mM NaCl, 0.1% Tween 20, and 5% (wt/vol) nonfat dry milk, blots were incubated overnight at 4°C with the primary antibody and washed with a Tris buffer (Tris-buffered saline pH 7.5 with 0.1% Tween 20). The blots were then incubated with horseradish peroxidase-conjugated secondary antibody (Pierce), washed again, incubated with SuperSignal Ultra Chemiluminescent reagent (Pierce), and, finally, exposed to Kodak X-Omat film (Eastman Kodak Company, Rochester, NY, USA). Bands on the films were scanned using the imaging system Chemilmager 4000 (Alpha Innotech Corporation, San Leandro, CA, USA), and the intensity of the L-PGDS bands was normalized by dividing them by the intensity of the β-actin band of the corresponding sample.

The levels of 11β-PGF2α in hyaluronidase-treated synovial fluids and of PGD₂ in chondrocyte supernatants were determined using competitive enzyme immunoassays from Cayman Chemical Company. Assays were performed according to the manufacturer's recommendation.

We first analyzed the levels of L-PGDS and H-PGDS mRNAs in healthy and OA cartilage using real-time quantitative RT-PCR. As shown in Figure 1, cartilage predominantly expresses L-PGDS mRNA, and its levels of expression were approximately threefold higher in OA cartilage compared with healthy cartilage. In contrast to L-PGDS, there was no statistically significant difference in the levels of H-PGDS mRNA between OA and healthy cartilage (Figure 1). In preliminary experiments, we showed that the amplification efficiencies of tested genes and GAPDH were similar. The efficiencies for the amplification of each gene and the reference were approximately equal, ranging between 1.95 and 2.

Next, we used immunohistohemistry to analyze the localization and the expression level of L-PGDS and H-PGDS proteins in healthy and OA cartilage. As shown in Figures 2a and 2b, the immunostaining for L-PGDS was located in the superficial and upper intermediate layers of cartilage. Statistical evaluation for the cell score revealed a clear and significant increase in the number of chondrocytes staining positive for L-PGDS in OA cartilage (43% ± 6%, mean ± SEM) compared with healthy cartilage (20% ± 4%, mean ± SEM). The specificity of the staining was confirmed using antibody that had been preadsorbed (1 hour at 37°C) with a 20-fold molar excess of the recombinant protein (Figure 2c) or nonimmune control IgG (data not shown). Using several commercially available antibodies directed against human H-PGDS, we were unable to detect H-PGDS protein expression in OA or healthy cartilage. Together, these data indicate that the expression level of L-PGDS is increased in OA cartilage.

To assess the level of PGD₂ in synovial fluids from OA and healthy donors, we quantified its major stable metabolite, 11β-PGF₂α. We measured this metabolite because PGD₂ is unstable in vivo 30 and quantification of PGD₂ in synovial fluid can be unreliable. We found a higher level of 11β-PGF₂α in OA synovial fluid when compared with healthy synovial fluid (Figure 3), indicating that the production of PGD₂ is higher in OA synovial fluids. Together, these data indicate increased expression and activity of L-PGDS in OA tissues.

IL-1β plays a major role in the cartilage physiology and in the pathogenesis of OA 2; therefore, we examined its effects on the expression of L-PGDS in cultured OA chondrocytes. Cells were treated with IL-1β (100 pg/mL) for different time periods, and the levels of L-PGDS mRNA were quantified using real-time RT-PCR. IL-1β-induced changes in gene expression were evaluated as fold over control (untreated cells) after normalization to the internal control gene, GAPDH. As shown in Figure 4a, treatment with IL-1β (100 pg/mL) enhanced L-PGDS mRNA expression in a time-dependent manner. L-PGDS mRNA expression started to gradually increase 24 hours post-stimulation with IL-1β and remained elevated until 72 hours. The induction of L-PGDS mRNA by IL-1β was also dose-dependent. A significant increase at concentrations as low as 10 pg/mL was observed and the maximal effect was reached at 100 pg/mL (Figure 4b). To determine whether changes in mRNA levels were paralleled by changes in L-PGDS protein levels, we performed Western blot analysis. Consistent with its effects on L-PGDS mRNA, treatment with IL-1β led to a dose- and time-dependent increase in the L-PGDS protein expression (Figure 4c, d). To establish whether the IL-1β-induced increase in L-PGDS expression corresponded with an increase in PGDS activity, we measured PGD₂ levels in conditioned media. As shown in Figures 4e and 4f, the increased expression of L-PGDS protein was accompanied by a time- and dose-dependent increase in PGD₂ production.

The lag period required for IL-1β to induce L-PGDS mRNA in chondrocytes contrasts with those required for other IL-1β-inducible genes, the expression of which starts as early as 2 to 6 hours and reaches a maximum at 8 to 18 hours. This suggests that de novo protein synthesis is required for IL-1β-induced L-PGDS expression. To evaluate this possibility, we examined the impact of the protein synthesis inhibitor CHX. Chondrocytes were stimulated with IL-1β in the absence or presence of CHX, and the levels of L-PGDS mRNA were analyzed by real-time PCR. As shown in Figure 5, treatment with CHX prevented IL-1β-mediated upregulation of L-PGDS mRNA expression. This suggests that, to upregulate L-PGDS expression in chondrocytes, IL-1β must induce the synthesis of one or more proteins.

IL-1β exerts its effects acting through activation of the mitogen-activated protein kinase (MAPK) (extracellular signal-regulated kinase [ERK], c-jun N-terminal kinase [JNK], and p38) and nuclear factor-kappa-B (NF-κB) signalling cascades 3132333435. To evaluate the potential contribution of these pathways in IL-1β-induced L-PGDS expression, we used specific pharmacological inhibitors. Chondrocytes were pretreated for 30 minutes with selective inhibitors for the above pathways and then stimulated or not with IL-1β for 48 hours. As shown in Figure 6a, pretreatment with the p38 MAPK inhibitor SB203580 (1 μM), the JNK MAPK inhibitor SP600125 (10 μM), or the NF-κB inhibitor SN-50 (1 μM) suppressed IL-1β-induced upregulation of L-PGDS expression. In contrast, pretreatment with the p42/44 MAPK inhibitor PD98059 (10 μM) had no effect on IL-1β-induced upregulation of L-PGDS. The concentration of the MAPK and NF-κB inhibitors used for these experiments had no significant effect on cell viability as indicated by the results of the MTT (3- [4,5-dimethylthiazol-2-yl]-2,5-diphenyltetrazolium bromide) assay (data not shown). These results suggest that the activation of JNK and p38 MAPK as well as NF-κB is essential to the induction of L-PGDS by IL-1β in chondrocytes.

The Notch signalling pathway regulates diverse cellular processes, including proliferation, differentiation, and apoptosis 36, and was reported to contribute to the regulation of L-PGDS expression 37. To determine whether this pathway participates in IL-1β-induced L-PGDS expression in human chondrocytes, we assessed the effect of DAPT. DAPT is a γ-secretase inhibitor, which blocks cleavage of the intracellular domain of all Notch proteins, and is widely used to evaluate the effect of Notch inhibition 36. As shown in Figure 6b, pretreatment with DAPT dose-dependently prevented IL-1β-induced L-PGDS protein expression, indicating the involvement of Notch signalling in this process. Notch inhibition was confirmed by transcriptional inhibition of its direct target gene, Hes1 (data not shown).

To further characterize the regulation of L-PGDS expression in cartilage, we examined the effect of PGD₂, the end product of L-PGDS. Chondrocytes were stimulated with IL-1β in the absence or presence of increasing concentrations of PGD₂ for 48 hours, and the expression of L-PGDS was evaluated by Western blotting. As shown in Figure 7, treatment with PGD₂ dose-dependently reduced IL-1β-induced L-PGDS expression.