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Local biochemical and morphological differences in human Achilles tendinopathy: a case control study
© J et al; licensee BioMed Central Ltd. 2012
Received: 5 December 2011
Accepted: 5 April 2012
Published: 5 April 2012
The incidence of Achilles tendinopathy is high and underlying etiology as well as biochemical and morphological pathology associated with the disease is largely unknown. The aim of the present study was to describe biochemical and morphological differences in chronic Achilles tendinopathy. The expressions of growth factors, inflammatory mediators and tendon morphology were determined in both chronically diseased and healthy tendon parts.
Thirty Achilles tendinopathy patients were randomized to an expression-study (n = 16) or a structural-study (n = 14). Biopsies from two areas in the Achilles tendon were taken and structural parameters: fibril density, fibril size, volume fraction of cells and the nucleus/cytoplasm ratio of cells were determined. Further gene expressions of various genes were analyzed.
Significantly smaller collagen fibrils and a higher volume fraction of cells were observed in the tendinopathic region of the tendon. Markers for collagen and its synthesis collagen 1, collagen 3, fibronectin, tenascin-c, transforming growth factor-β fibromodulin, and markers of collagen breakdown matrix metalloproteinase-2, matrix metalloproteinase-9 and metallopeptidase inhibitor-2 were significantly increased in the tendinopathic region. No altered expressions of markers for fibrillogenesis, inflammation or wound healing were observed.
The present study indicates that an increased expression of factors stimulating the turnover of connective tissue is present in the diseased part of tendinopathic tendons, associated with an increased number of cells in the injured area as well as an increased number of smaller and thinner fibrils in the diseased tendon region. As no fibrillogenesis, inflammation or wound healing could be detected, the present data supports the notion that tendinopathy is an ongoing degenerative process.
Current Controlled Trials ISRCTN20896880
Tendons connect muscle to bone and enable transmission of forces from contracting muscle to bone, resulting in joint movement. They possess the ability to adapt to changes in loading  and studies have shown that collagen synthesis is increased as a result of both acute exercise [2, 3] and prolonged physical training . The adaptation to loading can ultimately lead to increases in CSA and collagen content in chronically loaded tendons . Despite this physiological ability to adapt, tendinopatic tendons represents a large and constantly growing clinical problem affecting both recreational and professional athletes as well as people involved in repetitive labour [6, 7]. Years of research have unfortunately not provided much insight into the pathogenesis of chronic tendinopathy . Indeed, the etiology of tendinopathy has been related to repeated micro strain below the failure threshold as an initiating stimulus for degenerative processes [9, 10]. Other authors, however, have proposed that mechano-biological under-stimulation results in a degenerative cascade, through the production of a pattern of catabolic gene expression that leads ultimately to extracellular matrix degeneration . Tendinopathy is characterized by activity-related pain, focal tendon tenderness, and decreased local movement in the affected area [12, 13]. The general opinion is that no inflammatory cells are present in the tendinopathic tissue  and that tendinopathy is the result of a degenerative process with collagen disorganization, collagen fibre separation, increased cellularity, neovascularization and focal necrosis .
Previous studies have shown an altered concentration of certain matrix metalloproteinases MMPs, AdAMt's and TIMP's in normal and degenerate human Achilles tendon . Additionally several cytokines [9, 10] can be found in tendons and fibroblasts after cyclic mechanical stretching in healthy tendon tissue. However, the published data arises from the comparison of tendinopathic tissue with either control tissue from different anatomical tendons  or with tissues from identical anatomical tendons but from different subjects . Since considerable microscopic structure differences have been demonstrated in anatomically different tendons , this limits the conclusions that may be drawn from these studies. Taking the aforementioned limitations into account, current data concerning local biochemical differences within tendinopathic tendons, seem to indicate that an altered expression of collagen , proteoglycans  and matrix metalloproteinases [16, 22] exists in tendinopathic tendons. In addition the level of cytokines  VEGF and fibronectin  has been shown to be significantly different in the tendinopathic area. However analyses of local biochemical differences together with morphological differences are lacking.
The aim of the present study was to elucidate if any local structural differences are present in tendinopathic areas of human Achilles tendons compared to healthy areas in the same tendon. Furthermore, we wanted to investigate which proteoglycans, growth factors and cytokines that were involved in the local structural differences observed.
We hypothesize that several markers such as collagen 3 would be locally up regulated indicating formation of scar tissue with in the tendon  and higher concentrations of MMP-2 and MMP-9 indicating an enhanced degradation of collagen structures in the tendinopathic area (t-area) when compared with the healthy area (h-area) of the same tendon. Furthermore it is hypothesized that certain proteoglycans would have altered expression in the two tendon regions, e.g. an increased expression of decorin which might cause the collagen turnover to be increased also in chronic tendinopathic tendons. Additionally we hypothesize that growth factors like fibroblast growth factor (bFGF) are decreased causing a reduced healing capacity in the injured area of the Achilles tendons.
Thirty patients with chronic Achilles tendon pain were included in this study approved by the local Ethical Committee of the Capital Region Copenhagen (H-1-2009-114) and in compliance with the Helsinki Declaration. Additionally the study was registered at Current Controlled Trials (ISRCTN20896880). Due to limitations in the amount of tissue gained from the tendon biopsies patients were randomly assigned to either a Structural study (n = 14) or a Biochemical study (n = 16) by the envelope method. All subjects were recreational athletes or workers with a long-term history of chronic Achilles tendon pain (> 1/2 year) (Table 4) and conventional conservative treatments (eccentric rehabilitation, NSAIDs and corticosteroid injections) had been tried in all individuals with no effect. Intake of NSAID or corticosteroid injection was not allowed 6 months prior to inclusion in the present study. All subjects were recruited from the Rheumatology Department, Silkeborg Hospital, Denmark, and the biopsies from the Achilles tendons were taken as part of a standard procedure in order to examine for deposits of cholesterol, uric acid, and amyloid in the injured Achilles tendons.
The subjects were locally anesthetized, in the peritendinous space from both the medial and lateral side of the tendon with injections of 2 × 10 ml 1% Lidocain, using ultrasound guidance. Biopsies were taken with a semi-automatic biopsy needle (14 GA, 9 cm; Angiotech) also using ultrasound (US) guidance. An initial tendon biopsy was taken in the maximally sick area evaluated using US (defined as the area with maximal increased tendon thickness, neovascularisation, hypoeccogenicity). This area was usually 3-5 cm above the attachment of the Achilles tendon to the calcanaeus bone. A second biopsy was taken from the same tendon 4 cm proximal to the first biopsy in a region of the tendon tissue that was deemed normal using US.
Biopsy samples intended for analysis using Transmission Electron Microscopy were immersed in 2% glutaraldehyde in 0.05 M sodium buffer (pH 7.2), and the samples for gene expression were snap-frozen and stored at -80°C until analysis.
Transmission electron microscopy of tendon biopsies
Fourteen tendon biopsy pairs were cut into small pieces and were immersion-fixed in 2% glutaraldehyde for 24 hours. Following three rinses in 0.15 M sodium phosphate buffer (pH 7.2) the specimens were post-fixed in 1% OsO4 in 0.12 M sodium cacodylic buffer for 2 hours. The specimens were dehydrated in a graded series of ethanol (70%, 96% and 100%), transferred to propylene oxide and embedded in Epon (VWR Bie&Berntsen) in three steps according to standard procedures. For each biopsy one ultra thin section was cut approximately perpendicular to the length axis of the tendon with a Reichert-Jung ultracut E microtome. The section was collected on a one-hole copper grid with a Formvar supporting membrane and stained with uranyl acetate and lead citrate. The sections were examined using a Phillips CM 100 transmission electron microscope operated at an accelerating voltage of 80 kV. Digital images were obtained with a MegaView II camera and an analysis software package. From each ultra thin section the intercellular tissue was examined by taking a simple, random sample of ten digitized TEM images of the intercellular tissue. The cellular component of the tendon was examined in eleven biopsy pairs by taking 6 times 6 images in three randomly positioned regions of the section. The 6 times 6 images were spliced into one image using multiple image alignment (MIA) tools, so for each examined biopsy a total of three MIA images were obtained.
The Stereological analyses of the images were carried out on a computer monitor onto which the digitized EM image was merged with a graphic representation of the stereological test systems for just 12 of the 14 biopsy pairs (2 biopsies was unfortunately not useable for stereology analyses) (C.A.S.T.-grid software, The International Stereology Center at Olympus). The intercellular tissue was analyzed at a final magnification of 210.000 in the ten ordinary TEM images. The volume fraction (Vv) of collagen fibrils per intercellular tissue volume was estimated with the point counting technique as the number of points hitting collagen fibres divided by the number of points hitting the intercellular tissue (including collagen fibrils) using a point grid of 36 points. The number of collagen fibrils per cross sectional area of intercellular tissue (NA) was counted in 16 uniformly positioned, unbiased counting frames, each with an area of 0.0426 mm2 (42.6 μm2), and the individual diameters (d) of the sampled collagen fibrils were measured as the largest diameter perpendicular to the longest axis (i.e. the length of the minor axis of the ellipse) using the "measure-length" feature of the CAST-grid system. The unbiased counting frame ensures that all profiles, regardless of shape, size or orientation, have an equal probability of being sampled within area probe. The MIA images were analyzed at a final magnification of 115,000. The point counting technique, using a point grid with approximately 1000 points, estimated the volume fractions of the cellular component of the tendon tissue. The estimated parameters were: the volume fraction of cells within the tendon, the volume fraction of the nucleus within the cell, and the volume fraction of cytoplasm within the cell. A single experienced investigator performed all stereological analyses in a blinded fashion. The investigator was blinded for all subject characteristics, and whether the sample was obtained from the tendinopathic or the healthy region of the tendon.
RNA extraction and real time-PCR analysis
Total RNA isolation: Total RNA was extracted from frozen tendon samples from 16 subjects (sample weight: mean 23.2 ± 6.4 mg) by using 1 ml of TRI Reagent (Molecular Research Centre, Cincinnati, OH) 5 steel beads (2.3 mm) and 4 silica beads (1.0 mm Silicon Carbide Beads (454 grams) BioSpec Products Inc.). Glycogen was added (120 μg per ml of TriReagent) to the tendon samples to improve RNA precipitation.
Extracted RNA was precipitated from the aqueous phase with isopropanol and was washed with ethanol [75%], dried and suspended in 10 μl of nuclease-free water. The RNA concentration was determined using a RiboGreen RNA Quantitation kit 200-2000 Assays, Molecular Probes USA. RNA quality was determined on the basis of a RNA 6000 nano Chip assay kit, Agilent Technologies, Germany. The RNA samples were stored frozen at -20°C until subsequent use in real-time RT-PCR procedures.
cDNA synthesis: 100 ng RNA was reverse transcribed for each tendon sample in a total volume of 20 μl by using the QiagenOmniscript RT Kit at 37°C for 1 hour followed by 70°C for 15 minutes. The resulting cDNA was diluted twenty times in dilution buffer (10 mMTris EDTA buffer: Sigma Germany) + Salmon Testes DNA (1 ng/μl; Sigma Germany), and samples were stored at -20°C until used in the PCR reactions for specific mRNA analysis.
Polymerase Chain Reaction: The Real-time PCR-method using Glyceraldehyde 3-phosphate dehydrogenase (GAPDH) and 60S acidic ribosomal protein P0 (RPLP0) as reference genes to study specific mRNA's of interest was applied. The primers were purchased from MWG Biotech. For each target cDNA the PCR reactions were carried out under identical conditions by using 5 μl diluted cDNA in a total volume of 25 μlQuantiTect SYBR Green PCR Mix (Qiagen) and 100 nM of each primer (Table 2). The amplification was monitored in real-time using a MX3005P real-time PCR machine (Stratagene, CA). The threshold cycle (Ct) values were related to a standard curve made with cloned PCR products to determine the relative difference between the unknown samples, accounting for the PCR efficiency. The specificity of the PCR reaction was confirmed by melting curve analysis after amplification. The real-time PCR conditions were as follows: to denaturate the DNA strands the reaction mix was heated above the melting temperature of DNA (95°C) for 10 minutes, followed by 50 cycles each of 15 seconds at 95°C, followed by the annealing step where optimal primer hybridization conditions were obtained by lowering the temperature to 58°C for 30 seconds, and the extension step, where the reaction mix was heated to 63°C for 90 seconds. Two housekeeping genes GAPDH and RPLP0 were used as reference genes. The RPLP0 gene had been chosen as an internal control, assuming RPLP0 to be constitutively expressed. To validate this assumption GAPDH mRNA was measured as another unrelated "constitutive" and normalized with RPLP0, showing no difference between the healthy and the tendinopathic region of the tendon (Figure 3).
The PCR data were log transformed and a Paired Students t-test was performed to compare the results from the healthy area of the tendon with the tendinopathic area of the tendon, with exception of the results from IL-6, IL-1b, ki67 and HGF-1. These gene targets could not be detected in all samples. In these cases Chi2 tests were performed. All PCR data are presented as the geo mean ± backtransformed SEM. The collagen fibril data were divided into area and diameter fractions, and a paired Students t-test was performed to compare each fraction between the healthy and the tendinopathic area of the tendon. Likewise, the volume fraction of cells and the volume fraction of the nucleus within the cell were compared using a Paired Student t-test comparing the two areas of the tendon. A P-value < 0.05 was considered to be significant and all data despite of the subject characteristics are shown as Mean ± SEM.
Structural composition of the tendon
Tendon fibril characteristics
Sick Tendon Tissue
Healthy Tendon Tissue
Gene expression analysis
Degradation factors: Expression of MMP-2, MMP-9 and TIMP-2 was significantly increased in the t-areas compared to that of the h-areas with no difference in expression of TIMP-1 (Figure 3).
History of symptoms [Y] (range)
Structural study (n = 14)
48 ± 12
86 ± 17
182 ± 8
26 ± 4
3 ± 2.5
Biochemical study (n = 16)
49 ± 10
85 ± 18
175 ± 10
28 ± 5
2 ± 1
The present study examined the differences in structural proteins, cellular volume densities and expression levels of various genes involved in regulation of matrix proteins in clinically and ultrasonographiclytendinopathic regions of the human Achilles tendon and healthy regions within the same Achilles tendons. The main findings were differences in the composition of collagen structures with the tendinopathic region containing significantly higher number of small size fibrils (diameter 10-40 nm) compared to the healthy region of the tendon. In addition, the tendinopathic region had a significantly higher volume fraction of cells, compatible with a greater number of cells per unit volume. Furthermore, expression of several genes involved in both collagen synthesis and collagen degradation was significantly up-regulated, an observation that is consistent with an increased local turnover of collagen tissue in the affected tendinopathic area of the tendon. Gene expression was also influenced by the disease as several factors involved in wound healing were expressed at a lower number in the tendinopathic area. Lastly no sign of increased inflammation was found in the diseased region. Taken together these data indicate that local morphological and biochemical differences are present within the tendon during Achilles tendinopathy. These findings may have implications in the choice of treatment for these patients.
We are grateful to Dr. A.P. Harrison, Faculty of LIFE Sciences, Copenhagen University, for reading and correcting this manuscript. This study was supported by grants from the Danish Rheumatism Association, The NovoNordic Foundation, the Danish Ministry of Culture Committee for Sports Research, the Danish Medical Research Counsel (22-04-0191) and the Nordea foundation (Healthy Aging grant).
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