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Title: Platelet Rich Plasma enhances theimmunohistochemical expression of platelet derived growthfactor and vascular endothelial growth factor in the synoviumof the meniscectomized rat models of osteoarthritis
Author: Shaima M. Almasry Hala M. Soliman Shereen A.El-Tarhouny Sami A. Algaidi Ehab M. Ragab
PII: S0940-9602(14)00179-4DOI: http://dx.doi.org/doi:10.1016/j.aanat.2014.10.006Reference: AANAT 50903
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Received date: 20-3-2014Revised date: 5-9-2014Accepted date: 14-10-2014
Please cite this article as: Almasry, S.M., Soliman, H.M., El-Tarhouny, S.A., Algaidi,S.A., Ragab, E.M.,Platelet Rich Plasma enhances the immunohistochemical expressionof platelet derived growth factor and vascular endothelial growth factor in the synoviumof the meniscectomized rat models of osteoarthritis, Annals of Anatomy (2014),http://dx.doi.org/10.1016/j.aanat.2014.10.006
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Platelet Rich Plasma enhances the immunohistochemical expression of platelet derived growth factor and vascular endothelial growth factor in the synovium of the meniscectomized
rat models of osteoarthritis
Shaima M Almasry,1, 2 Hala M.Soliman,1, 3 Shereen A. El‐Tarhouny,4, 5 Sami A. Algaidi,1 Ehab M.Ragab6
1Department of Anatomy, Taibah University, Saudi Arabia 2Department of Anatomy and Embryology, Almansoura University, Egypt 3Department of Histology and Cell Biology, Zagazig University, Egypt. 4Department of biochemistry and Molecular Medicine Taibah University, Saudi Arabia 5Department of Medical Biochemistry Zagazig University, Egypt 6Department of Orthopedic Surgery, Al Azhar University, Assuit branch, Egypt. Address correspondence to: Shaima Mohamad Almasry, Anatomy and Embryology Department, Taibah University, Almadinah Almunawarah, Saudi Arabia. Tel: 00966594380057. E‐mail: [email protected] Running head: Effect of PRP on osteoarthritic synovium
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SUMMARY
This study was carried out on a rat model of surgically‐induced osteoarthritis (OA) to assess the histological and immunohistochemical changes in the synovial membrane and to evaluate the effects of intra‐articular injection of Platelet Rich Plasma (PRP) in such cases. Forty five male albino rats were divided into 3 equal groups; control, surgically‐induced OA and surgically‐induced OA followed by intra‐articular injection of PRP. Knee joints were processed for histological and immunohistochemical staining with anti‐ platelet derived growth factor (PDGF‐A) and anti‐vascular endothelial growth factor (VEGF) and the area percentages of immunostaining were measured by digital image analysis. Serum levels of PDGF‐A and VEGF were analyzed by ELISA. The osteoarthritis research society international (OARSI) score was significantly higher in OA (2433.8±254) than in control (230.4±37.8; p<0.001) and in PRP‐treated tissues (759.7±45.8; p<0.001). The immunostained area percentages for PDGF‐A was significantly higher in PRP‐treated tissues (20.6±2.4) than in OA (11.06±1.3; p=0.007) and in control tissues (4.1±0.78; p<0.001). Likewise, the immunostained area percentage for VEGF was significantly higher in PRP‐treated tissues (22.5±1.6) than in OA (14.9±1; p=0. 002) and in control tissues (6.5±0.7; p<0.001). ELISA analysis revealed a significant increase in serum levels of the PDGF‐A and VEGF after intraarticular PRP injection when compared to the other groups (p<0.000). The present study concluded that intra‐articular injection of PRP could produce optimizing effects in surgically induced OA in the form of; decreasing the OARSI score, improving the inflammatory events in synovium and modulating the PDGF‐A and VEGF serum levels and synovial tissue immunoexpression. These effects could be reflected positively on the associated chondral defect.
Key words: PRP; rat model; osteoarthritis; synovium; PDGF; VEGF
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INTRODUCTION
Osteoarthritis (OA) is a group of diseases that involve the degeneration of articular cartilage and subchondral bone. Articular cartilage is a highly specialized load‐bearing tissue that provides low‐friction joint surfaces. Functions of articular cartilage depend on the precise interplay of the molecules that compose its extracellular matrix (ECM). The unique biomechanical properties of articular cartilage depend on two principal supramolecular matrix components; proteoglycans that are embedded in a mesh‐work of different types of collagen fibrils and aggrecan, the major proteoglycan in cartilage that has a high density of fixed negative charges owing to its anionic glycosaminoglycan side chains. This in turn pulls in water into the tissue and generates an osmotic swelling pressure that is counterbalanced by the tense collagen mesh‐work. Thus, collagens and proteoglycans create a hydrated tissue that resists compressive loads (Boschetti and Peretti, 2008; Bougault et al., 2013).
The chondrocyte in normal articular cartilage exists in a quiescent state in an avascular ECM nourished by diffusion from the synovial fluid and the vasculature of the subchondral bone. The term synovium or synovial membrane (SM) refers to the delicate tissue lining the spaces of diarthrodial joints, tendon sheaths and bursae. It includes the continuous surface layer of cells (intima) and the underlying tissue (subintima). The intima consists of macrophages and fibroblasts while the subintima includes blood and lymphatic vessels, a cellular content of both resident fibroblasts and infiltrating cells in a collagenous extracellular matrix. Between the intimal surfaces, there is a small quantity of fluid, usually rich in hyaluronic acid. In concert, this structure provides a non‐adherent surface between tissue elements (Smith, 2011; Smith and Walker, 2011; Siebelt et al., 2014). The regeneration capacity of cartilage is limited because of its isolation from systemic regulation and its lack of vessels and nerve supply (Buckwalter and Brown, 2004). Biomechanical, metabolism and biological changes may contribute to the loss of tissue homeostasis, resulting in an accelerated loss of the articular surface and followed by end‐stage arthritis (Hayami et al., 2008).
Clinically relevant OA joint disease is invariably associated with some sort of synovial pathology. This contemplates the feeling that there is a direct relationship between clinical symptoms and the synovial reaction in OA and most likely these changes in SM are at least partly involved in the advancement of the disease (Uchida et al., 2013). Local therapy such as intra‐articular injection of anti‐inflammatory agents is projected to be an effective measure to alleviate the symptoms and prevent the progression of OA (Palmieri et al., 2013). The rationale for intraarticular therapy is the delivery of the drug to the site of action, therefore minimizing the systemic toxic effect of the drug.
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Platelet-rich plasma (PRP) is a simple, low cost, and minimally invasive method that permits one to extract a natural concentrate of autologous growth factors (GFs) from the blood (Anitua et al., 2004; Say et al., 2013). PRP is composed of 3-8-fold greater concentration of platelets, as compared to whole blood (Qureshi et al., 2009). The use of animal models of OA has been fairly extensive for testing of potential anti‐arthritic agents and disease modifying effects for agents currently used to treat patients with OA (Kobayashi et al., 2000). Animals that predominantly load the medial aspect of the joint like rodents will develop more severe lesions on the medial side after a medial meniscectomy than on the lateral side after a similar insult and vice versa (Bendele, 2001).
Considering the possible regenerative effects of PRP on cartilage damage and SM changes, this study, for the first time, aimed to evaluate the effects of intra‐articular PPR injection on the histopathological changes and the immunoexpressive behavior of PDGF‐A/VEGF in the SM of a rat model of OA.
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MATERIALS AND METHODS
Experimental animals
Forty five adult male healthy Wistar albino rats, 250‐350 gms were obtained from Jeddah and kept under constant conditions; 12:12 light/dark cycle and a room temperature of 28 0C. They were allowed free access to standard rat chow diet and water ad libitum. Animals were divided randomly into 3 equal groups (n=15). The groups included control, surgically‐induced OA and surgically‐induced OA followed by intra‐articular injection of PRP.
Prior to dosing, animals were acclimated to the environment for 7 days. An experiment was executed in conformity with National Institutes of Health (NIH) guidelines for the maintenance and use of science lab animals; NIH Publication 1986 (86/609/EEC) and in accordance with local laws and ordinances.
Preparation of PRP
Rats were anesthetized with a subcutaneous injection of Ketamine (0.15 ml/kg) and Medetomidine (0.1 ml/kg) and then submitted to cannulation via the jugular vein. Two milliliters of each animal blood was collected using a disposable syringe containing 0.5 ml of 0.1 Mol sodium citrate as an anticoagulant. The blood was preserved in silicone vacuum tubes. The same amount of withdrawn blood was immediately replaced through injection of sterile saline according to the method adopted by Messora et al. (2001).
Then animals were left for a 3 week recovery phase before the surgical intervention (for OA and PRP‐treated groups) during which, the PRP preparation was prepared by adapting the protocol proposed by Sonnleitner et al. (2000), using a cold centrifuge and a vertical laminar flow cabinet for manipulating the biological samples. The collected blood was firstly centrifuged at 160 G, for 20 minutes, at room temperature (22oC). The whole plasma above the buffy coat was collected, separating platelets from red blood cells and leukocytes and transferred to other 5 ml vacuum tube. The sample was then submitted to a new centrifugation at 400 G, for 15 minutes, resulting in two components; the upper half is platelet‐poor plasma (PPP) and the lower half is the platelet rich plasma (PRP). PRP were pipetted and transferred to sterile tubes stored at ‐30°C. Immediately before intraarticular injection, PRP were activated by using 0.05 ml of 0.2 Mol calcium chloride solution to each 1 ml of PRP. 150‐200 µl of PRP could be obtained from each ml whole blood.
Surgical induction of osteoarthritis and dosing protocol
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Thirty rats were anaesthetized and subjected to complete medial meniscal tear procedure under complete aseptic technique using Chlorhexidine spray (ICI Company, London, UK). A skin incision was made over the medial aspect of the knee, and the medial collateral ligament was exposed by blunt dissection and then transected just below its attachment to the meniscus so that when the joint space opens, the meniscus is reflected toward the femur. The meniscus is cut at its narrowest point (away from the ossicles) taking care not to damage the tibial surface. Skin and subcutis were closed in 4‐0 Vicryl suture (Bendele, 2001; Flannery et al., 2009). Postoperative daily use of the Gentamicin Topical Spray (Pharmex Comp., Chicago, Illinois, USA) was applied to the site of surgery for one week.
Rats of the PRP‐treated group were subjected to surgical intervention followed by intra‐articular injection of 85 µl of activated PRP in the operated knee by using an insulin syringe, one week after surgery, with dosing performed under local anesthesia three times per week (every other day) for two weeks. At the end of the procedure, the rat’s knees were bent and extended for a few times to allow the PRP to spread throughout the joint before becoming a gel.
The second blood sample (1 ml) was drawn from tail veins in plain tubes, 24 hours after the last injection of PRP in the PRP‐treated group and for the other 2 groups at the same time and left for 30‐60 minutes for spontaneous clotting at room temperature then centrifuged at 3000 rpm for 10 minutes. Serum samples were separated into another set of tubes and kept frozen at ‐80°C. All rats from control, OA and PRP treated groups were sacrificed 6 weeks after induction of surgery.
Enzyme‐Linked Immunosorbent Assay
Total serum levels of PDGF‐A and VEGF in samples were determined by the ELISA technique, using an available commercial kit (Quantikine R&D Systems, CAT DAA00B and RRV00, Minneapolis, USA) with horseradish peroxide detection in accordance with the manufacturer’s instructions. We used 50 μL to sample for both PDGF‐A and VEGF. Samples were assayed as duplicates.
Tissue sampling
A sample of synovium from the suprapatellar fold was removed from the operated knee joint and placed in 10% neutral buffered formalin. Then, the muscle was trimmed from the femur and tibia and the patella was reflected distally and removed to allow fixative (10% neutral buffered formalin) to enter the joint space. The femur and tibia were transected some distance from the joint to avoid fragmentation of bone into the joint area. The joint was dropped into 4% paraformaldehyde, pH 7.4 for 3 days. Then, the ends of the bones were trimmed and a forceps was placed in the patellar groove and posterior aspect of the joint. The
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joint was cut (using the collateral ligaments as a land mark) into approximately equal halves (frontal plane), and were decalcified in Ethylene diamine tetraacetic acid for one month. The trimmed halves were washed extensively, processed for paraffin embedding. Coronal tissue blocks were cut across the medial tibial plateaus.
Measuring the tibial cartilage only may be adequate for tibiofemoral disease in cross sectional studies. The use of the tibial cartilage has an advantage in that it is a clearly defined, anatomical structure and thus less susceptible to error in its identification across diferent study sites and in different operator’s hands (Cicuttini et al., 2001).
For histological analysis, synovium specimens were stained with HE, Alcian blue with PAS, Masson’s Trichrome and immunohistochemistry for PDGF‐A and VEGF. Chondral specimens were stained with HE, Alcian blue with PAS only. Alcian blue with PAS is a combination used to evaluate the distribution of the various mucins, glycogen and other PAS‐positive tissue elements. PAS stain demonstrates certain polysaccharides, specifically glycogen and mucoproteins as reddish structures. Whole sections were studied microscopically and scored by a histopathologist who was blinded with respect to treatment group. SM was examined generally, despite their location in the areas of chondral defects.
Immunohistochemical assessment
For immunohistochemical analysis, we used primary goat polyclonal anti‐PDGF‐AA antibody (R&D Systems Inc., Minneapolis, MN 55413, USA; catalog # AF1560) and primary rabbit polyclonal anti‐VEGF antibody (Spring bioscience, 6920 Koll Center Parkway, Pleasanton, CA 94566 USA; catalog # M3281). Staining procedure used an immunoperoxidase technique as previously described (Bancroft and Gamble, 2002). Paraffin sections were deparaffinized, rehydrated then treated in 3% H2O2 for 5 min and washed with phosphate buffer solution (PBS) for 15 minutes. The sections were blocked with 1.5% normal goat serum in PBS, and were then incubated for 45 min at room temperature with the primary antibody. Sections were subsequently incubated with a second‐stage biotinylated antibodies (Biotinylated goat anti‐rabbit IgG, Spring Bioscience, catalog # DGR‐125) at room temperature. After rinsing in PBS, the reaction products were visualized by immersing the section into diaminobenzidine. Finally, the sections were counterstained with hematoxylin, dehydrated, and covered. For negative controls, incubation was carried out by a non‐specific IgG antibody at the same concentration as the primary antibody (Taylor, 1993).
Interpretation of staining
Tissue sections were examined using an optical microscope at 4x and 10x magnification for the initial screening. Measurements were performed at 10x magnification. Images were
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digitized in a 512 × 512‐pixel matrix, using a color video camera (digital camera CH‐9435 DFC 290). Digitized pictures were visualized on a high‐resolution color display. The true color image analysis software package using an image analysis system (Leica Imaging System, Switzerland & Germany) was run for manipulation, quantification of the images and data collected. Computerized imaging analysis systems have been introduced in order to minimize subjectivity in evaluation the percentage of area occupied by the brownish immunohistochemical staining.
A positive reaction to the studied proteins was distinct as a brownish staining in the cytoplasm as referenced in the Catalog. Microanatomic‐distribution of the staining and staining intensity were evaluated as previously described (Jackson et al., 2011), weak (light yellow); moderate (yellow–brown); and strong (brown). Strong intensity was defined as the strongest intensity relative to weaker staining intensity within the examined sections.
Histologic analysis and OARSI (osteoarthritis research society international) scoring
Most of the microscopic scoring systems described for rat OA restrict evaluation of OA to cartilage changes, while a few include OA‐associated changes in bone and synovial membrane. In some of the recently published rat OA studies, the OARSI score was used to grade the degenerative status of the repaired tissue (Appleton et al., 2007; Yorimitsu et al., 2008). It multiplies the cartilage damage score by a factor to reflect the extent of the involved tibial plateau. OARSI was assessed by evaluating the following nine points as modified by Gerwin et al. (2010).
1. Cartilage matrix loss width
The width of the area of collagen matrix loss was measured along the surface (0% depth), as well as at the level of the mid‐zone (50% depth) and tidemark (100% depth) (Fig. 1A).
2. Cartilage degeneration score
This score is an evaluation of overall cartilage pathology and includes the important pathology parameters of collagen matrix fibrillation/loss and chondrocyte death/loss with chondrocyte loss being the primary determinant of the score. For the cartilage degeneration score, the MTP was divided into three zones. The % area of each zone that contains a cartilage exhibiting loss of chondrocytes (50% or greater loss of normal cellularity) or loss of matrix was estimated and a score was assigned to that zone based on that percentage. A 3‐zone‐sum for cartilage degeneration was also estimated by totaling the values obtained for each zone. The maximum 3‐zone sum for the medial tibia was 15 (Fig. 1B).
3. Zonal depth ratio of lesions
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This is a measurement of the depth of cartilage degeneration (e.g., including areas of chondrocyte and proteoglycan loss, which may have good retention of collagenous matrix and no fibrillation) that was taken at the midpoint in each of the three zones across the tibial surface. The lesion depth ratio was calculated by dividing the micrometer depth of the area of degeneration by the thickness of the cartilage (both in micrometers), from projected cartilage surface to tidemark (Fig. 1B).
4, 5. Total and significant cartilage degeneration width
The total width of the area of articular cartilage affected by any type of degenerative change (matrix fibrillation/loss, PG loss with or without chondrocyte death) was measured in micrometers. Significant cartilage degeneration width is a measurement of the width of the tibial cartilage in which 50% or greater of the thickness (from surface to tidemark) is seriously compromised (Fig. 1C).
6. Osteophyte score
The largest osteophyte in each section was measured (from the deepest point of its base at the chondro‐osseous junction to the surface of the overlying cartilage at its thickest point) (Fig. 1D). This categorizes osteophyte measurements on a scale of 0‐4, where 0=marginal zone proliferative changes (≤200 µm), 1=small (200‐299 µm), 2=moderate (300–399 µm), 3=large (400–499 µm), and 4=very large (≥500 µm).
7. Calcified cartilage and subchondral bone damage score
This parameter was used to quantify effects of agents on OA‐associated changes in subchondral bone and calcified cartilage (Fig. 1D). Damage to the calcified cartilage layer and subchondral bone was scored using a numerical scale of 0‐5, where 0= no changes and 5= increased basophilia at the tidemark, increased fragmentation at the tidemark and/or of the calcified cartilage, increased area of marrow mesenchymal changes, and collapse of articular cartilage into the epiphysis.
8. Synovial reaction
Inflammation of SM (if present) was scored using a numerical scale of 0‐4, where 0= no changes (1‐2 layers of synovial lining), 4= increased number of lining cell layers (>4 layers) and/or proliferation of subsynovial tissue and infiltration of inflammatory cells.
9. Medial capsule width
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Micrometer measurements of the medial capsule width were obtained for each joint, as an assessment of soft tissue fibroplasia.
Statistical data analysis
Statistical analysis was done using the SPSS V. 13.0 software (SPSS, Chicago, USA). The mean±standard error (SE) for each parameter or measurement is defined in order to get a total joint score. Statistical analysis was performed using One‐way ANOVA with Dunnett’s test for analysis of the measured parameters. Scored parameters were analyzed using Kruskale Wallis and Man‐Whitney tests. P‐value<0.05 was considered statistically significant.
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RESULTS
Survival rate of animals
No death occurred in the rats of the control group, whereas 3 rats in the OA group had died; one arrested on the table from an overdose of anesthesia, and the remaining two died within two days after operation. In the PRP‐treated group, 2 instances of deaths were reported by the end of the first week after surgery. Rats of the OA/PRP groups which had died in the days following the operative day showed temperature elevations before death.
Assessment of OARSI score
OARSI score is found to be significantly lower in control animals (230.4±37.8) compared to OA (2433.8±254; p<0.001) and PRP‐treated (759.7±45.8; p<0.001) animals. After PRP‐treatment, the OARSI score is decreased significantly compared to OA specimens (p<0.001) and the control specimens (p<0.001). Representative statistical data among the individual histopathological scores used to assess OARSI score in the knee sections are indicated in Table (1).
Histological analysis
Light microscopic examination of the SM from the control group reveals a regular structure of the SM in the form of 2 rows of flat synoviocytes with spindle shaped nuclei resting on subintimal adipose and loose areolar connective tissue. Subintimal layer contains fibroblasts, and apparently normal blood vessels. Few thin collagen fibers as well as weak PAS reactions are seen in intimal and subintimal layers and around the blood vessel (Figs. 2A, B & 3A, B).
Arthritis induction produces thick SM which is constituted of multiple layers of synoviocytes and with wide intercellular spaces. Subintima is formed by thick fibrous tissue separated by adipocytes and mononuclear infiltrates. Subintimal blood vessels are numerous, congested with perivascular areas of fibrosis. Macrophages in intima and subintima can be seen. Thick collagen fiber in the intima and subintimal layers and around the blood vessel and strong PAS reaction are present (Figs. 2C, D & 3C, D).
After PRP injection, SM shows a thin intima, compared to osteoarthritis sections, with a single layer of synoviocytes resting on subintimal layer formed mainly of loose areolar tissue. Subintimal adipose connective tissue and regular blood vessels can be viewed. Less collagen fibers in the intimal and subintimal layers and around blood vessels with moderately positive PAS reaction when compared to osteoarthritis group are observed (Figs. 2E, F & 3E, F).
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Thus, the histological results validated the sampling method of tissues and correspond to the grading data of the OARSI score and to the discrimination of the synovial membrane specimens from control, OA and PRP‐treated groups.
Immunohistochemical analysis
Localization of PDGF‐A
PDGF‐A is expressed throughout all layers of the SM of control specimens; mild staining is detected in the synovial lining layer, in the stroma cells as well as in the vascular endothelium (Figs. 4A, B). In conditions of tissue localization, similar results for PDGF‐A are obtained with SM of OA and PRP‐treated specimen; however, the density of staining differs. In SM of OA, moderate staining is observed in the fibroblasts, cellular infiltrates and in the vascular endothelium (Figs. 4C, D). After injection of PRP, specimens reveal strong PDGF‐A expression in the synoviocytes, stroma cells and in the vascular endothelium (Figs. 4E, F). Localization of VEGF
Mild positive immunoreactivity can be seen in the synovial lining and sublining cells and in the vascular endothelial cells (Figs. 5A, B). In OA cases, mild staining is found in the lining cells while strong staining is observed in the perivascular and vascular endothelial cells (Figs. 5C, D). In SM of PRP‐treated cases, moderate staining is observed in the intimal lining and subintimal cells while strong staining is observed around the blood vessels and in the vascular endothelial cells (Figs. 5E, F).
Statistical analysis demonstrates significantly higher immunostained area percentages for PDGF‐A in SM from the knees of the PRP‐treated group (mean±SE, 20.6±2.4) compared to OA (11.06±1.3; p=0. 007) and control groups (4.1±0.78; p<0.001). Likewise, there is a significantly higher immunostained area percentage for VEGF in SM from the knees of the PRP‐treated group (22.5±1.6) than in OA (14.9±1; p=0. 002) and control ones (6.5±0.7; p<0.001) (Fig. 6).
ELISA analysis
PDGF‐A and VEGF serum levels are measured by ELISA assay and confirm the immunohistochemical findings. We observe a significant increase in serum levels of PDGF‐A and VEGF after intraarticular PRP injection (154.2±5.3, 232.1±3.1 pg/mg) compared to OA (128.9±1.3, 185.9±3.07 pg/mg; p<0.001) and control (113±1.4, 89.6±1.6 pg/gm; p<0.001) groups (Table 2).
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DISCUSSION
To date, there are insufficient studies on the histological evidence to sanction the clinical usage of intra‐articular injection of PRP as a treatment modality in OA problems, and to our knowledge, no studies have dealt with the effects of intra‐articular delivery of PRP on the histological picture of SM in these troubles. In this study, we assessed the OA‐related histopathological and growth factor immunoexpressive changes in the synovium and cartilage of rat knee and the possible, improving effects of PRP particularly on the synovial changes which were hypothesized to be reflected positively on the chondral defect.
The overall results showed that the intra‐articular injection of PRP has positive effects as shown by the return of the regular histological features, lessening of the OARSI grading score, and enhancement of expression of PDGF‐A and VEGF in the synovial membrane. These effects are especially noteworthy since they provide the possibility of a new therapeutic measure to attenuate the progression and ease the symptoms of OA.
Examination of the histological sections illustrates the regular structure of SM in the control specimens while OA specimens show proliferative reactions in the synovial intima and subintima in the form of synovial cell hyperplasia, inflammatory cellular infiltrates, fibrosis and increased vascularity. These changes are indicative of synovitis, as the histological examination has previously been the gold standard for the diagnosis of synovitis (Wenham and Conaghan, 2010). Nearly similar results have been previously described; the scores for synovial cellularity, vascularity and inflammation were found to be markedly increased in the OA knees compared with normal ones (Smith et al., 1997). The proliferative synovial cells and infiltrated mononuclear cells may play a major part in the destruction of cartilage and bone in the joints. Interestingly, Blom and colleagues demonstrated that macrophage activation in the synovium is essential for cartilage damage via the production of matrix metalloproteinases, suggesting that inflammation within the synovium may be pivotal to cartilage damage (Blom et al., 2007).
In addition, the synovitis that has been detected in OA knees is found to be not confined to areas of chondral defects as ensured by the examination sites. Similar general localization of synovitis have been lately reported by Wenham and Conaghan (2010), however; earlier studies demonstrated an association between the chondral defects and associated synovitis in the medial tibiofemoral compartment of the knee (Ayral et al., 2005; Loeuille et al., 2005). Whether the synovitis is limited to the areas of chondral defects or not, it has previously been well established that synovial inflammation is directly implicated in the initiation and progression of OA (Suri et al., 1996). Synovial inflammation is an important source of
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proinflammatory mediators produced by synovial cells induces a degradative cascade leading to joint damage (Lambert et al., 2012).
After intra‐articular injection of PRP, there is noticeable improvement in the histological picture with lessening of the proliferative response in the intima and subintima compared to OA specimens, therefore, PRP could attenuate the subsequent inflammatory response in OA. Backing up this histological analysis in SM, the statistical analysis of OARSI score reflecting the articular cartilage changes shows significantly lower scores after injection of PRP than those of OA without treatment. The possible regenerative effects of PRP injection and its role in preserving the histological structure of SM in surgically‐induced knee OA, have been explained by the physiological role of platelets reported previously in wound healing (Nurden et al., 2008) and by the evidence that platelets discharge bioactive proteins responsible for attracting macrophages, mesenchymal stem cells, osteoblasts, facilitating tissue regeneration, healing and promote scavenging of necrotic tissue (Sampson et al., 2008). In summary, it was demonstrated that PRP increased hyaluronic acid concentration, stabilizing angiogenesis in patients with osteoarthritis knees (Anitua et al., 2007; Sampson et al., 2008). Besides, intraarticular delivery of PRP allows its presence in a high concentration in the joint fluid and thus, PRP is allowed to pass through the affected cartilage and reach the SM by diffusion.
Recent animal studies have assessed the PRP effect on chondrogenesis and cartilage healing. In a rabbit model with osteochondral defects in the femoropatellar groove, Sun et al. (2010) demonstrated a higher extent of cartilage regeneration and increased production of glycosaminoglycans in the extracelluar matrix after treatment with autogenous PRP in poly‐lacticglycolic acid. Additionally, Qi et al. (2009) established chondrogenesis with PRP when PRP was used with a scaffold.
In this study, for the first time, we show the regulatory effect of intraarticular PRP on the immunoexpressive manners of PDGF‐A and VEGF in SM of osteoarthritis rat knee. PDGF‐A is immunoexpressed in nearly all the layers of SM of control, OA and PRH‐treated specimens; namely the synovial lining layer, the stroma cells as well as in the vascular endothelium. Nevertheless, the statistical data reveal differential regulation between groups; there are substantial increases in the percentage of PDGF‐A immunostained area in the PRP‐treated group compared to OA and is lowest in the control group. ELISA analysis is consistent with immunoexpressive findings.
The expression pattern of PDGF‐A in the layers of synovial membrane is matched by many previous reports (Sano et al., 1993; Pohlers et al., 2006). Numerous GFs were found to regulate the growth and homeostasis of such tissues throughout life (Goldring et al., 2006). PDGF‐A and VEGF receptors were expressed in capillary endothelial cells and have been
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demonstrated to induce angiogenic effects to varying degrees (Marx et al., 1994, Nicosia et al., 1994).
The present results indicating the upregulation of PDGF‐A after induction of osteoarthrities is matched by a previous studied reporting the potential role of PDGF‐A in synovial hyperplasia and in augmenting the response of fibroblast like synoviocytes to selected inflammatory mediators (Chen et al., 2005; Pohlers et al., 2006). Early observations in wound healing revealed that PDGF‐A is released by platelets and by activated macrophages (Shimokado et al., 1985), yet the detailed functional roles of PDGF‐A and its importance in the pathogenesis of arthritic diseases remain speculative.
The substantial increase in PDGF‐A expression after injection of PRP could be explained by the findings that platelets contain an abundant amount of GFs that are crucial in soft tissue healing and bone mineralization (Anitua et al., 2006) and gives an evidence of the role of PDGF‐A in cartilage repair. Recently, Amable et al., (2013) stated that high concentrations of PDGF and other GFs are secreted from platelet α‐granules after activation and that GFs are highly retained inside clots and would be slowly released in vivo. The slow release of the GFs could guard against formation of pannus tissue during OA or after PRP injection.
The possible regenerative role of PDGF‐A is postulated either to its mitogenic activity to the mesoderm‐derived cells, such as fibroblasts and chondrocytes or to its specific activities in angiogenesis and chemotaxis for fibroblasts and collagen synthesis (Fortier et al., 2011; Civinini et al., 2013). Also, PDGF‐A could have a stimulatory effect on matrix synthesis in growth plate chondrocytes (Schmidt et al., 2006). Thus, the upregulation of PDGF‐A in the OA and in the PRP‐treated groups could suggest a multifunctional role of this GF in the process of cartilage destruction and healing.
VEGF immunoexpressive analysis reveals positive staining in the synovial lining and sublining cells and in the vascular endothelial cells of all studied specimens and in the perivascular tissue of the OA group. Statistical data showed variability in the expression levels in the studied groups. VEGF‐immunoexpressed area percentage is significantly higher in PRP‐treated specimens compared to the control and OA ones. These data are consistent with the serum findings.
VEGF has been extensively studied with regard to its function in angiogenesis; it acts on endothelial cells to stimulate their proliferation and migration (Neufeld et al., 1999) and found to be associated with synovial angiogenesis (Haywood et al., 2003). Uncontrolled angiogenesis is a decisive event of synovial inflammation and these two interdependent processes, that is, angiogenesis and inflammation, are now seen as an important contributor of OA (Bonnet and
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Walsh, 2005). This is coherent with the present result which reveals upregulation of VEGF expression in OA synovium and is matched with the previous data of Enomoto et al. (2003) and Haywood et al. (2003). They reported a possible role of VEGF in hypervascularization of inflamed tissue. Though they considered the VEGF as an early marker of arthritis and synovitis, the exact mechanism by which this GF affect the pathogenesis of OA is not distinctly seen.
In the present work, PRP treatment causes significant increase in the expression of VEGF in PRP‐treated compared to OA specimens which could be attriputed to its stimulating effect on mitogenesis and regeneration of the endothelial cells in this tissue (Hoeben et al., 2004; Leitner et al., 2006). Nevertheless, dissimilar to our results, other investigators have shown no change in VEGF gene expression in a porcine model of immunologic arthritis (Lippross et al., 2011).
An asymmetry of these regulatory GFs, which may result from disease, or injury, may hinder tissue maintenance and repair, resulting in deleterious changes in gene expression, altered extracellular matrix, tissue degeneration and consequently an accelerated erosion of the articular surface, leading to end‐stage arthritis (Alves et al., 2010).
More significantly, our study related the synovial GF expression with the synovial and chondral histopathological changes in the path of the OA and suggested that PDGF‐A and VEGF may play an important part in joint destruction and repair. Although functional collaboration of different GFs exists in articular inflammation, specific GFs may be responsible for a specific aspect of the disease process. PRP treatment could modulate the expression of a specific GF at an appropriate stage and thus could effectively regulate the disease process.
CONCLUSION
The histological and biochemical results of our study are encouraging and suggest that intraarticular injected platelet rich plasma may be successfully used for the handling of surgically‐induced osteoarthritis in the rat knee. PRP may act at different levels; reducing inflammatory events in synovium and modulating PDGF‐A and VEGF serum levels and synovial tissue immunoexpression. These effects could reflect positively on the associated chondral defect.
RECOMMENDATION
Large animal model and longer duration postoperative follow up should be performed in order to confirm the findings from this study and provide evidence for further clinical investigation.
ACKNOWLEDGMENTS
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This study was supported by grant No. 929⁄1433 from the Deanship of Taibah University, Almadinah Almonawarah, Kingdom of Saudi Arabia. The authors thank Mr. Mohammed Farook Ahmed, a technologist in Almadinah Maternity and children Hospital for his assistance during handling the samples.
CONFLICT OF INTEREST
The authors declare that there is no conflict of interest related to this study.
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TABLE
Table 1. Histopathological changes in the knee of the three studied groups.
Histopathological changes (mean±SE)
Control (n=15)
Osteoarthritis (n=12)
Platelet rich plasma‐treated
(n=13)
P value
1. Cartilage matrix loss (in µm) (mean±SE)
42.9±14.2 823.8±105.9 267.3±20.8 P1<0.001* P2<0.001* P3<0.001* P4=0.001*
2. Cartilage degeneration score (mean rank)
8.9 33.96 21.46 P1<0.001* P2<0.001* P3<0.001* P4<0.001*
3. Zonal depth ratio of lesions (mean±SE)
0.2±0.05 1.5±0.1 0.6±0.1 P1<0.001* P2<0.001* P3=0.013* P4<0.001*
4. Total cartilage degeneration width (in µm) (mean±SE)
71.3±22.5 776.3±102.8 161.8±12.8 P1<0.001* P2<0.001* P3=0.006* P4<0.001*
5. Significant cartilage degeneration width (in µm)
(mean±SE)
24.2±6.9 547.3±81.4 134.3±13.3 P1<0.001* P2<0.001* P3<0.001* P4=0.001*
6. Osteophyte score (mean rank) 11.6 21.96 29.42 P1<0.001* P2=0.004* P3<0.001* P4=0.041*
7. Calcified cartilage and subchondral bone damage score
(mean rank)
11.6 17.67 33.38 P1<0.001* P2=0.025* P3<0.001* P4<0.001*
8. Synovial reaction (mean rank) 9.5 32.38 22.23 P1<0.001* P2<0.001* P3<0.001* P4<0.003*
9. Medial capsule width (in µm) 90.2±2.7 272±16.1 177±8.3 P1<0.001*
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(mean±SE) P2<0.001* P3<0.001* P4<0.001*
Total OARSI score (mean±SE) 230.4±37.8 2433.8±254 759.7±45.8 P1<0.001* P2<0.001* P3<0.001* P4<0.001*
P1: significance in differences of variables between the 3 studied groups; P2 between the control and osteoarthritis groups; P3 between the control and PRP‐treated groups; P4: between the osteoarthritis and PRP‐treated groups. * Significant p<0.05.
Table 2. Mean levels of PDGF‐A and VEGF in plasma of control, osteoarthritis and Platelet rich plasma‐treated cases.
Control (n=15)
Osteoarthritis (n=12)
Platelet rich plasma‐treated
(n=13)
P value
PDGF‐A (pg/ml) (mean±SE) 113±1.4 128.9±1.3 154.2±5.3 P<0.001*
VEGF (pg/ml) (mean±SE) 89.6±1.6 185.9±3.07 232.1±3.1 P<0.001*
P: significance in differences of variables between the 3 studied groups; * Significant p<0.05.
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FIGURE LEGENDS
Figure (1): Photomicrograph of medial tibial plateau from rat which had medial meniscal tear surgery. A) For measuring the widths of collagen matrix loss*1*at 0% (surface), 100% (tidemark) and 50% (midzone) depths. B) For evaluation of the cartilage degeneration score*2*, the original surface of the tissue must be estimated (red tracing), then the percentage area of each zone that contains cartilage exhibiting loss of chondrocytes (50% or greater loss of normal cellularity) or loss of matrix (yellow tracing) is estimated. The lesion depth ratio*3*is calculated by dividing the depth of the lesion (yellow line) by the thickness of the cartilage from projected articular surface to tidemark (black vertical line). These measurements are taken at the midpoint of each zone. C) The total cartilage degeneration width*4*(black horizontal line) is taken at the projected cartilage surface from the outer edge of the tibial plateau, adjacent to the osteophyte to the point at which the cartilage is normal. The significant cartilage degeneration width*5*(red horizontal line) represents the width of tibial cartilage in which 50% or more of the original cartilage thickness is seriously compromised. D) The largest osteophyte*6*in the section is measured from base to edge at the thickest point (green line) and then given a score based on that measurement. Calcified cartilage and subchondral bone damage score*7*. There is increased basophilia at the tidemark (red arrow) with marked multifocal fragmentation of calcified cartilage of tidemark and marrow mesenchymal changes upto ¾ of the total area (A‐D, Alcian blue/PAS X40) (modified from Gerwin et al., 2010).
Figure (2): Photomicrographs of histological sections of synovial membrane from rat knee. (A, B) Control sections show thin layer of synovial intima (arrows) formed of two rows of flat synoviocytes with spindle shaped nuclei resting on subintimal adipose (A) and loose areolar connective tissue (L). Subintima contains fibroblasts (arrow head), few collagen fibers (double arrows) and regular blood vessels (inset, V). (C, D) Osteoarthritis sections show thick layer of intima (I) with synoviocytes arranged in multiple layers and separated by wide intercellular spaces (thin arrow). Subintima is formed of dense fibrous tissue (arrow head),(inset, arrow head) separated by adipocytes (D) and mononuclear infiltrates (double arrows). Subintimal blood vessels are numerous, congested with perivascular areas of fibrosis (V). Note the presence of Macrophages in intima and subintima (thick arrow). (E, F) PRP‐treated sections show thin intima (arrows), compared to osteoarthritis sections, with single layer of synoviocytes resting on subintimal layer formed mainly of loose areolar tissue (L). Subintimal adipose connective tissue (double arrows) and regular blood vessels (V) can be seen. (A, C, E: HE X100, B, D, F, insets; HE X1000).
Figure (3): Photomicrographs of histological sections of synovial membrane from rat knee. (A, B) Control sections show few thin collagen fibers (thin arrows) in the subintimal layer and
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around blood vessels. Weak PAS reactions (thick arrows) are seen in intimal and subintimal layers and around the blood vessel. (C, D) Osteoarthritis sections reveal thick collagen fibers (thin arrows) in the intimal and subintimal layers and around the blood vessel with strong PAS reaction (thick arrows). (E, F) PRP‐treated sections show less collagen fibers (thin arrows) in the intimal and subintimal layers and around blood vessels with moderately positive PAS reaction (thick arrows) when compared to osteoarthritis group. (A, C, E: Masson’s Trichrom X100 & B, D, F: Alcian blue/PAS X100).
Figure (4): Photomicrographs of PDGF‐immunostained sections. (A, B) Control sections reveal mild PDGF expression in the cytoplasm of synoviocytes of intima (thin arrow) and in the subintimal adipose connective tissue (double arrows). Moderate staining are present in the cytoplasm of vascular endothelium (thick arrows). (C, D) Osteoarthritis sections reveal moderate PDGF expression in the cytoplasm of synoviocytes (thin arrow), subintimal adipocytes (double arrows) and around the blood vessels. Note the strong positive staining in the cytoplasm of vascular endothelium (thick arrow). (E, F) PRP‐treated sections reveal strong PDGF expression in the cytoplasm of synoviocytes (thin arrow), adipocytes (double arrows), vascular endothelium (thick arrows) and around the blood vessels. (A, C, E: anti‐PDGF X100 & B, D, F: anti‐PDGF X400).
Figure (5): Photomicrographs of VEGF‐immunostained sections. (A, B) Control sections reveal weak VEGF expression in the cytoplasm of synoviocytes (thin arrow) and subintimal adipocytes (double arrows). Moderate staining is seen in the cytoplasm of vascular endothelium (thick arrows). (C, D) Osteoarthritis sections reveal mild VEGF expression in the cytoplasm of synoviocytes (thin arrow), subintimal connective tissue (double arrows), while strong reactions around the blood vessels (red arrow) and in the cytoplasm of vascular endothelium (thick arrows) can be seen. (E, F) PRP‐treated sections reveal moderate VEGF expression in the cytoplasm of synoviocytes (thin arrow) and subintimal adipocytes (double arrows) while strong reactions are seen in the cytoplasm of vascular endothelium. (A, C, E: anti‐VEGF X100 & B, D, F: anti‐VEGF X400).
Figure (6): Box and whisker plot (median, interquartile range and range) of immunoexpressed area% of PDGF and VEGF in the studied groups.
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