Mesenchymal stem cell derived exosomes and injectable platelet-rich fibrin enhance SDFT healing in a donkey tendonitis model.

Skip to main content An official website of the United States government Here's how you know Here's how you know Official websites use .gov A .gov website belongs to an official government organization in the United States. Secure .gov websites use HTTPS A lock ( Lock Locked padlock icon ) or https:// means you've safely connected to the .gov website. Share sensitive information only on official, secure websites. Search Log in Dashboard Publications Account settings Log out Search… Search NCBI Primary site navigation Search Logged in as: Dashboard Publications Account settings Log in Search PMC Full-Text Archive Search in PMC Journal List User Guide PERMALINK Copy As a library, NLM provides access to scientific literature. Inclusion in an NLM database does not imply endorsement of, or agreement with, the contents by NLM or the National Institutes of Health. Learn more: PMC Disclaimer | PMC Copyright Notice Sci Rep . 2026 Apr 10;16:12013. doi: 10.1038/s41598-026-44967-7 Search in PMC Search in PubMed View in NLM Catalog Add to search Mesenchymal stem cell derived exosomes and injectable platelet-rich fibrin enhance SDFT healing in a donkey tendonitis model Mahmoud Najeb Mahmoud Najeb 1 Department of Surgery, Anesthesiology, and Radiology, Faculty of Veterinary Medicine, Mansoura University, Mansoura, 35516 Egypt Find articles by Mahmoud Najeb 1 , Alaa Samy Alaa Samy 1 Department of Surgery, Anesthesiology, and Radiology, Faculty of Veterinary Medicine, Mansoura University, Mansoura, 35516 Egypt Find articles by Alaa Samy 1 , Awad Rizk Awad Rizk 1 Department of Surgery, Anesthesiology, and Radiology, Faculty of Veterinary Medicine, Mansoura University, Mansoura, 35516 Egypt Find articles by Awad Rizk 1 , Esam Mosbah Esam Mosbah 1 Department of Surgery, Anesthesiology, and Radiology, Faculty of Veterinary Medicine, Mansoura University, Mansoura, 35516 Egypt Find articles by Esam Mosbah 1 , Iman Ibrahim Iman Ibrahim 2 Department of Pathology, Faculty of Veterinary Medicine, Mansoura University, Mansoura, Egypt Find articles by Iman Ibrahim 2 , Gamal Karrouf Gamal Karrouf , Ph.D 1 Department of Surgery, Anesthesiology, and Radiology, Faculty of Veterinary Medicine, Mansoura University, Mansoura, 35516 Egypt Find articles by Gamal Karrouf 1, ✉ Author information Article notes Copyright and License information 1 Department of Surgery, Anesthesiology, and Radiology, Faculty of Veterinary Medicine, Mansoura University, Mansoura, 35516 Egypt 2 Department of Pathology, Faculty of Veterinary Medicine, Mansoura University, Mansoura, Egypt ✉ Corresponding author. Received 2025 Aug 31; Accepted 2026 Mar 16; Collection date 2026. © The Author(s) 2026 Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if changes were made. The images or other third party material in this article are included in the article’s Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article’s Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit http://creativecommons.org/licenses/by/4.0/ . PMC Copyright notice PMCID: PMC13069002  PMID: 41963425 Abstract Tendon injuries pose a significant challenge in equine medicine due to the inherently limited regenerative capacity of tendon tissue and the high likelihood of reinjury. Recently, injectable platelet-rich fibrin (I-PRF) and exosomes derived from mesenchymal stem cells (MSCs) have shown promise as regenerative treatments, offering potential benefits in matrix restoration and modulation of inflammatory responses. This study investigated the effects of I-PRF alone and in combination with MSC-derived exosomes on collagenase-induced superficial digital flexor (SDF) tendonitis in donkeys. Twenty-seven donkeys were randomly assigned to three treatment groups: saline (placebo), I-PRF, and PRF/exosome. Tendon healing was monitored over a 20-week period using clinical evaluation, ultrasonography, histology, immunohistochemistry, and biomechanical testing. At the study endpoint, the combined PRF/exosome group exhibited the greatest improvement in tendon consistency and weight-bearing ability compared to placebo ( P < 0.001). Ultrasonographic findings were markedly better in the PRF/exosome group, even though tendon cross-sectional area (T-CSA) remained elevated across all treated groups, while it decreased significantly in the placebo group. Immunohistochemical analysis revealed the highest collagen type I and lowest type III levels in tendons treated with PRF/exosome. Biomechanical testing further demonstrated superior failure stress and strain in this group relative to placebo. Overall, these results indicate a potential synergistic effect of combining I-PRF with MSC-derived exosomes in promoting tendon repair. Nevertheless, additional long-term studies in naturally occurring tendon injuries are required to confirm these findings and assess their clinical relevance. Supplementary Information The online version contains supplementary material available at 10.1038/s41598-026-44967-7. Keywords: Injectable platelet-rich fibrin, MSC-derived exosomes, Tendon regeneration, Superficial digital flexor tendonitis, Donkey Subject terms: Diseases, Medical research, Rheumatology, Stem cells Introduction Injuries to tendons, especially the superficial digital flexor tendon (SDFT), are common and often career-limiting in equines. Such injuries can account for as much as 72% of lost training days and contribute significantly to early retirement from athletic activity 1 – 3 . Equine SDF tendonitis not only affects horse performance but also imposes notable economic problems, with reported incidence rates ranging between 11% and 46% of all limb-related injuries 4 , 5 . Management of such injuries remains highly challenging, as tendon healing predominantly occurs through fibrotic scar formation rather than true regenerative 6 . Consequently, tendons seldom regain their native elasticity and mechanical integrity, which explains the high recurrence rates reaching nearly 80% following conventional therapies 7 – 9 . Numerous treatment strategies have been suggested and continue to be developed for managing SDF tendonitis in horses. The ongoing emergence of these treatment approaches highlights both the complex pathophysiology of the condition and the continued difficulty in establishing clear evidence for their effectiveness 10 . Recent progress in regenerative medicine has led to the development of various biologic treatments designed to enhance tendon healing, including mesenchymal stem cells and platelet-derived products (PDPs) 11 , 12 . Among PDPs, platelet-rich plasma (PRP) has been widely studied and has demonstrated promising outcomes in controlled experimental settings 13 , 14 . Yet, its therapeutic effectiveness in equine tendon repair continues to be debated in both experimental studies and clinical trials 10 . Two systematic reviews, including thirty studies, found that PRP improves lameness, tissue healing, and return-to-competition rates 15 , 16 . Conversely, a recent meta-analysis encompassing fifteen studies reported insufficient proof to support a significant contribution of PRP to tendon healing in equines 17 . As a second-generation platelet concentrate, platelet-rich fibrin (PRF) provides distinct advantages compared with PRP. Its preparation is straightforward, requiring only autologous blood and simple stall-side handling. Unlike PRP, which releases growth factors for a limited period of around four days, PRF maintains a higher and more sustained release for up to two weeks 18 , 19 . Additionally, the fibrin-rich structure of PRF supports cell migration and provides a natural scaffold, without containing chemical additives that could potentially compromise its therapeutic effects 20 . Unlike the traditional clotted form, the liquid variant, known as injectable PRF (I-PRF), allows for direct administration into the tendon tissue. Although PRF has not yet been widely investigated in the management of equine SDF tendonitis, its application in other tendinopathies has yielded variable results 21 – 24 . Although these biologics possess regenerative potential, their clinical outcomes in horses remain inconsistent, and complete regenerative repair is frequently not attained 10 . One of the emerging explanations for this inconsistency is inadequate resolution of the inflammatory response during tendon repair 25 – 27 . Prolonged or unresolved inflammation can delay progression to the reparative phase and may also reduce the effectiveness of regenerative treatments 28 , 29 . The initial phase of tendon repair is characterized by inflammation and cellular proliferation that, when inadequately regulated, may hasten tendon fiber deterioration. Concurrently, imbalanced collagen production promotes the predominance of mechanically inferior type III collagen over type I, ultimately giving rise to structurally compromised scar tissue 30 . Understanding and precisely modulating inflammation has become central to improving treatment outcomes and promoting true regeneration 31 . Tendon stem cells and immune cells are essential regulators of tendon healing, particularly through their control of inflammation and coordination of repair 32 , 33 . Tendon stem/progenitor cells respond to inflammatory cytokines like IL-1β by releasing anti-inflammatory factors such as IL-10 and TIMP-3, helping to limit excessive inflammation and support tissue remodeling 34 . In parallel, immune cells, especially macrophages, transition from pro- to anti-inflammatory states, contributing to resolution and repair 35 , 36 . The interaction between these cell types is crucial in determining whether healing leads to regeneration or chronic degeneration 32 . MSC-derived exosomes are gaining attention as a promising cell-free therapeutic approach for tendon healing, particularly through enhancing intercellular communication and modulating key signaling pathways involved in inflammation and regeneration 37 and their ability to influence macrophage polarization 38 , 39 . Several experimental model studies have demonstrated that MSC-derived exosomes control inflammation and enhance tendon repair 37 , 40 – 42 . These exosomes activate main signaling pathways including PI3K/AKT and MAPK/ERK1/2, reducing apoptosis 42 , regulating the balance between ECM synthesis and degradation, and contributing to a more regenerative microenvironment 43 , 44 . Additionally, they modulate the immune response by downregulation of pro-inflammatory cytokines and upregulation of anti-inflammatory markers, ultimately reducing scar formation and improving biomechanical properties 45 . Additionally, MSC-derived exosomes contribute to tendon healing through the shift of macrophages toward a phenotype that supports tissue regeneration 39 . Furthermore, combining MSC-derived exosomes with fibrin scaffolds has demonstrated superior tendon healing outcomes by enhancing histological scores, upregulating tenogenic markers, and improving biomechanical properties, highlighting the synergistic potential of this approach in tendon regeneration 44 . Since tendonitis encompasses both inflammatory dysregulation and tissue degeneration, an intralesional therapeutic strategy that combines immunomodulatory and regenerative modalities could result in improved outcomes. This study investigates the effectiveness of PRF, either individually or combined with MSC-derived exosomes, in a donkey model of SDF tendonitis. It is hypothesized that MSC-derived exosomes may attenuate excessive inflammation while augmenting the regenerative capacity of PRF by facilitating more effective intercellular communication and signaling. Materials and methods Animals, ethics endorsement, and consent for participation The Medical Research Ethics Committee of the Faculty of Veterinary Medicine, Mansoura University, approved this prospective, blinded, placebo-controlled experimental study (Code No: Ph.D./101). All procedures involving animals were carried out following institutional guidelines and national regulations for the care and use of laboratory animals. The study design and reporting adhered to the ARRIVE guidelines. Thirty-three clinically healthy male donkeys (36–48 ± 6.51 months old; 170 ± 11.68 kg) were obtained from the Faculty of Veterinary Medicine farm, Mansoura University, and housed in isolation stables at the Mansoura Veterinary Teaching Hospital, Egypt. Before enrollment, all donkeys underwent clinical and ultrasonographic evaluations to confirm soundness. Clinical assessment included general and orthopedic physical examinations, with palpation of the tendons and evaluation of gait at a walk and trot in straight lines and circles on a hard surface. Ultrasonographic examination of the SDFTs was performed bilaterally to assess echogenicity and fiber alignment, ensuring the absence of pre-existing tendon pathology. Donkeys exhibiting lameness, swelling, heat, pain on palpation, or ultrasonographic abnormalities were excluded from the study. Animals were kept under constant conditions and received a balanced diet of chopped wheat straw, bran, and whole corn. One week before the experiment, donkeys were dewormed with doramectin (1 ml/50 kg BW, IM; Dectomax, Zoetis, USA). Study design SDF tendonitis was induced in both forelimbs of 30 donkeys at the mid-metacarpal region. After seven days, 27 donkeys were randomly assigned to one of three treatment groups ( n = 9 each) using a computer-generated randomization sequence: the Placebo group, treated with saline; the PRF group, treated with I-PRF; and the PRF/Exosome group, treated with I-PRF and MSCs-derived exosomes. The remaining three donkeys (6 limbs) were retained as untreated tendonitis models (positive controls), while additional three healthy animals (6 limbs) were maintained as negative controls. Tendon healing was assessed through clinical, ultrasonographic, histopathological, immunohistochemical (IHC), and biomechanical evaluations at specific time points, as detailed later, with six forelimbs ( n = 6) evaluated per group at each time point (Fig. 1 ). Fig. 1. Open in a new tab The time diagram illustrates the sequential phases of the experimental design. SDF tendonitis induction The SDF tendonitis induction was accomplished under light general anesthesia using a modified previously described technique 46 . Donkeys received intravenous sedation with acepromazine (0.05 mg/kg, Castran, 15 mg/mL) followed 20 min later by xylazine HCl (1.1 mg/kg, Xyla-Ject, 20 mg/mL). Subsequently, butorphanol (0.05 mg/kg, Torbugesic 10 mg/mL) was administered. General anesthesia was induced using IV injection of thiopental (Anapental 500 mg, Corden Pharma, Egypt) at a dose rate of 2 mg/kg BW. On a lateral recumbency, the metacarpal (MC) regions of both limbs were aseptically prepared and scraped using povidone iodine. Under ultrasonographic guidance, an insulin needle was inserted laterally into the central SDFT at the mid-metacarpal region (midpoint between the distal end of the accessory carpal bone and the highest of the proximal sesamoid bones). A core lesion was then created via a sole injection of 2.5 mg collagenase (125 CDU/mg; Sigma, St. Louis, MO, USA) diluted in 0.3 mL saline. A pressure bandage was applied for 24 h post-injection. A three-day course of intramuscular procaine penicillin G (20,000 IU/kg SID) was administered, beginning one day before tendonitis induction and continuing for two days thereafter. Additionally, all animals administered systemic Flunixin meglumine (Flamicure 5%, Pharma Swede, Egypt) at a dose of 1.1 mg/kg once daily for three consecutive days starting immediately after collagenase injection, to control post-induction pain. The donkeys were stall rested for 24 h post-induction, then subjected to daily 5–15 min walking sessions for 7 days to simulate the mechanical load and stress associated with naturally occurring injuries. Autologous injectable PRF (I-PRF) preparation Briefly, 4 ml of venous blood was drawn into a sterile plain plastic tube and immediately subjected to centrifugation at 700 rpm for three minutes (22 g RCF) using a rotor set at a 45° angle with a 40 mm radius. This process resulted in the separation of an upper I-PRF fraction from the underlying red blood cell layer, and the obtained I-PRF was administered within 5–10 min to prevent clotting 47 . Mesenchymal stem cell-derived exosome (MSC-derived Exosome) preparation Exosomes were prepared immediately before treatment by reconstituting a 3 gm allogenic lyophilized powder (ExSommé, Bioluga ® Canada) in 2 ml of the supplied excipient, followed by the addition of 1 ml of sterile distilled water. The mixture was gently mixed until a clear and homogeneous injectable solution was obtained. According to the manufacturer, ExSommé ® contains lyophilized exosome concentrates derived from 6 × 10⁶ bone marrow-derived MSCs, with a protein concentration of 500 µg/ml (± 10%) as determined by BCA protein assay. Manufacturer-provided characterization indicated a mean particle size of approximately 130 nm measured by dynamic light scattering, typical cup-shaped vesicle morphology observed by transmission electron microscopy, and high expression (> 95%) of canonical exosomal markers CD63, CD81, and CD9. Treatment, follow-up, and controlled exercise On the 7th day post-collagenase injection (Ti7 = T0), local injection of the treatment inoculum was accomplished on a lateral recumbency, after sedation and induction of thiopental anesthesia, and under complete aseptic preparation of the metacarpal regions of both forelimbs. Based on group allocation, the treatment inoculum of 3 mL volume contained either 3 mL of normal saline solution (placebo group), 1.5 mL of I-PRF + 1.5 mL saline (PRF group), or 1.5 mL I-PRF + 1.5 mL exosomes (PRF/Exosome group) and was administered using an 18-gauge needle under ultrasonographic guidance. The injection was distributed across three sites: intralesional [at the maximum injured zone (MIZ)], 0.5 cm above, and 0.5 cm below it. Following treatment, a pressure bandage was subsequently applied to the metacarpal region for one day. All animals were subjected to a progressively increasing exercise program, as previously described 48 . After a 3-day rest period following treatment, the exercise program commenced on day 4 with a 5-minute walking session per day. Walking time was extended at a rate of five minutes each week until reaching 30 min per day by week 7. Starting from week 8, the program incorporated trotting, beginning with 2 min of trotting per day and gradually increasing by 2-minute increments every two weeks. Evaluations of tendon healing Clinical evaluation Animals were carefully examined for degree of lameness, pain to pressure, hotness upon palpation, and presence of discomfort daily through the induction days (Ti1-Ti7) ( Tables 1 and 2 ) . Following treatment, these evaluations were continued daily for the initial week, and subsequently at 2, 4, 8, 14, and 20 weeks post-treatment. Lameness scores were assessed in accordance with the modified scores by Riex et al. 49 . Pain and hotness were semi-quantitatively scored from 0 to 3, where 0 represented normal findings, 1 indicated mild symptoms, 2 denoted moderate manifestations, and 3 signified severe signs 48 . The donkeys were examined for the presence of discomfort using the modified donkey chronic pain scale 50 . Total discomfort scores ranged from 0 for comfort, 1–3 for mild discomfort, 4–6 for moderate discomfort, and 7–9 for severe discomfort. Table 1. Clinical assessment scores for tendon healing 48 , 49 . Score Description Heat 0 Normal findings 1 Mild increase 2 Moderate increase 3 Severe increase Pain 0 Normal findings 1 Mild reaction 2 Moderate reaction 3 Severe reaction Modified clinical score assessment for lameness 0 Normal gait 1 Mild, intermittent lameness or difficult to observe, regardless of surface 2 Mild lameness, intermittent at a walk but consistently present under certain conditions (e.g., hard surface, weight-bearing) 3 Mildly abnormal gait and/or stiff walk 4 Reluctance to move when motivated/severe lameness at a walk 5 No movement or lying down; limb barely touches the ground (minimal weight-bearing or resting limb in flexion posture) Tendon shape upon palpation 0 Smooth, uniform normal thickening 1 Minimal irregularity or slight thickening 2 Notable irregularity 3 severe irregularity with a soft consistency 4 Hard nodular area Intensifying weight bearing (static examination) 0 Even weight distribution 1 Slight reduction in weight-bearing, slight preference for the opposite limb 2 Notable reduction in weight-bearing, notable preference for the opposite limb 3 Notable offloading of the limb Open in a new tab Table 2. Discomfort score according to a modified donkey pain scale 50 . General appearance 0 Alert and/or is interacting with a mate/group. 1 Mildly depressed and/or decreased interaction with mate/group. 2 Moderately depressed and/or aggressive, or no reaction with mate/group. 3 Severely depressed. Standing Posture score 0 Quietly standing and/or one hind leg resting. 1 Slightly tucked up abdomen and/or mild weight shifting. 2 Extremely tucked up abdomen and/or hunched back and/or stretching limbs/body and/or mild muscle tremors. 3 Sits on hind quarters and/or extreme muscle tremors. Resting posture (lying down -Excluding Auto Grooming) 0 Does not lie down or rest lying down. 1 Attempts to lie down or is lying down < 50% of the time. 2 Lying down > 50% of the time. 3 Lies down in an abnormal position: on its side with stretched limbs or on its back, and/or is repeatedly rolling. 0–9 Total discomfort score Open in a new tab Additionally, tendon shape and weight-bearing ability were examined at T0 and T20. To assess static load-bearing capacity, a controlled load equal to 30% of the animal’s body weight was applied. All evaluations were independently and blindly performed by two clinicians to ensure objective assessment. Ultrasonographic evaluation All tendons were evaluated using B-mode ultrasonography before collagenase injection (N), on day 7 post-collagenase injection, which also marked the start of treatment (Ti7 = T0), then at 1 week post-treatment (T1), at 2 week post-treatment (T2), and subsequently at two weeks intervals up to 20 weeks (T2–T20).This was carried out with a 7.5–10 MHz linear scanner in both longitudinal and cross-sectional planes. Fiber echogenicity score (FES) was classified into four grades: grade 0 indicated normal echogenicity, grade 1 represented hypoechogenicity, grade 2 corresponded to heterogeneous or mixed echogenicity, and grade 3 denoted anechoic areas. Fiber alignment score (FAS) was determined based on the assessed proportion of parallel fibers as score 0 when more than 75% of fibers were parallel, score 1 when 50–74% were parallel, score 2 when 25–49% were parallel, and score 3 when less than 25% demonstrated parallel alignment (Table S1 ) 48 . At the level of the MIZ, both the T-CSA and the L-CSA were quantified (mm²) using transverse sonograms. The proportion of lesion (Lesion %) was subsequently calculated by dividing the lesion cross-sectional area (L-CSA) by the total tendon cross-sectional area (T-CSA) and multiplying the result by 100 51 . Because significant intergroup variations were detected in baseline (T0) values of Lesion% and T-CSA, relative changes were assessed longitudinally within each group. For this purpose, baseline values were normalized to 1 (T0/T0), and measurements recorded at subsequent time points were normalized by dividing them by their respective baseline (T0) values 52 . US evaluations were performed by M.N., whereas image analyses were carried out by an independent radiologist blinded to treatment allocation. Each variable was measured in triplicate, and the mean of these values was then calculated and used for the subsequent statistical analysis. Tissue harvest At the scheduled time points of T8, T14, and T20 weeks post-treatment, donkeys were sedated via IV injection of xylazine HCl at a dose of 2 mg/kg. Euthanasia was then achieved by rapid IV injection of a pentobarbital overdose 51 . At T8 & T14, the SDFTs were cautiously separated from the surrounding tissues. For histological and IHC evaluations, 2 cm tendon samples were harvested, including the central lesion site as well as adjacent proximal and distal healthy tissue. At T20, 7 cm-long SDFT specimens were harvested and longitudinally sectioned into two halves. One-half was submitted for biomechanical testing. In parallel, a 2 cm segment was excised from the second half and processed for histological and IHC analysis, following the same protocol as in earlier time points. Additionally, donkeys in the control positive group were euthanized on day 7 following collagenase injection to represent the baseline lesion state (T0) for histopathological evaluation. Meanwhile, control negative animals, which did not receive any tendon injury or treatment, served as the source of normal tendon specimens. Histopathological evaluation In weeks 8, 14, and 20, collected SDFT samples were processed following the routine histopathological preparation protocol. For histological assessment, three randomly selected sections from each sample were evaluated in a blinded manner by a pathologist. A modified semi-quantitative multiparametric scoring system was employed, as conducted in Table S2 53 . This scoring system involved the quantitative evaluation of six parameters, including: fiber structure, fiber arrangement, rounding of the nuclei, inflammatory cell infiltration, increased vascularity, and cell density. Each parameter was graded on a scale of 0 to 3, where 0 indicated normal arrangement, 1 represented mild abnormalities, 2 moderate tendon abnormalities, and 3severe tendon abnormalities. Immunohistochemistry (IHC) IHC was conducted to assess the relative proportions of collagen types I & III in tendon Sect. (5 μm thick) fixed on adhesion slides (Thermo Scientific, Fischer Scientific, UK) and processed following standard deparaffinization and rehydration protocols 48 . For collagen type I staining, the sections underwent enzymatic antigen retrieval using Pronase and Hyaluronidase to unmask collagen epitopes embedded in dense matrix regions. Non-specific sites were blocked with goat serum, and the samples were then incubated with a primary rabbit polyclonal antibody against type I collagen. Detection was achieved with an HRP-conjugated secondary antibody and DAB chromogen, producing a brown signal at positive sites. Nuclei were counterstained with hematoxylin before dehydration and mounting. Collagen type III staining employed a heat-induced antigen retrieval method using EDTA buffer, followed by suppression of endogenous peroxidase activity. After blocking with goat serum, sections were exposed to a primary rabbit polyclonal anti-collagen type III antibody. Signal amplification was accomplished using a biotinylated secondary antibody in conjunction with an ABC-HRP complex, and visualization was performed with DAB as described for type I. All rinsing steps were carried out with phosphate-buffered saline, ensuring removal of unbound reagents between incubations. This approach enabled clear differentiation and quantification of the two collagen types within the tendon matrix. For quantitative analysis, high-resolution digital images of selected regions of interest (ROIs) were extracted at magnifications up to ×40 and processed using ImageJ software (FIJI, National Institutes of Health, USA). Color deconvolution was applied to isolate the brown DAB chromogen signal from the hematoxylin counterstain, resulting in a DAB-specific monochrome image. A threshold was applied to highlight positively stained areas. The image was then binarized, and watershed segmentation was used to separate overlapping elements. Within each ROI, the DAB-positive area was quantified using frequency analysis, and the area fraction (percentage of positive staining relative to the selected area) was calculated. All measurements were performed in at least three representative ROIs per section, and the results were expressed as mean ± SEM 54 . Biomechanical examinations Immediately after collection, specimens were preserved in phosphate-buffered saline at − 20 °C, taking care to avoid repeated freeze-thaw cycles. Prior to testing, the samples were gradually brought to room temperature to allow complete thawing. The 7 cm-long SDFT specimens were loaded into a tensile testing device (Model LRX, Lloyd Instruments Ltd, UK), with a jaw separation of 4.2 cm. Sandpaper was placed between the sample ends and the clamps to minimize slippage during testing. After stabilization, the midsection’s width and depth were recorded in the transverse plane with a digital caliper to determine the cross-sectional area. A small preload of 5 N was applied for five cycles at 1 Hz to remove specimen laxity. Each specimen was then subjected to uniaxial tension until failure using a 200 kgf load cell at a steady crosshead speed of 10 mm/min. The peak force recorded before rupture was taken as the failure load (N), while failure stress (MPa) was calculated by dividing this load by the sample’s cross-sectional area. The strain (%) was determined by expressing the elongation at the point of failure relative to the specimen’s initial length 55 . Statistical analysis Statistical evaluation was performed in SPSS v28 (IBM Corp., Armonk, NY). Data distribution was evaluated with the Shapiro–Wilk test. Normally distributed variables are summarized as mean (SD), whereas ordinal data are reported as median (min–max). Parametric variables were evaluated with two-way ANOVA followed by Tukey adjustment. To further control for Type I errors across groups and time points, post-hoc analyses were adjusted using the Bonferroni method. Paired t-tests compared T-CSA at T20 with pre-induction (Normal) and baseline (T0). Groups were compared for change from baseline (T0) for lesion percentage and T-CSA. Nonparametric data were analyzed using Kruskal–Wallis (between-group), Friedman (within-group over time), and Wilcoxon signed-rank tests (T0 vs. T20). Statistical significance was defined as P ≤ 0.05, and all graphs and visual representations were generated using GraphPad Prism version 8.4.3. Results Clinical findings During the tendonitis induction period, pain and heat scores peaked within the first two days post-collagenase injection, with median scores of 3 (3–3), displaying severe pain reactions upon palpation and a marked increase in local temperature. By the 3rd day, a significant reduction was observed compared to day 1 ( P = 0.027), with scores declining to 2 (1–2), where animals exhibited moderate pain and heat responses. By the 4th day, a further significant decrease was noted compared to day 3 ( P = 0.016), with the score reaching 1 (0–1), representing mild heat and pain responses. By the 7th day, pain and heat responses had significantly diminished to 0 (0–1) compared to day 4 ( P = 0.006, r = 0.65, 95% CI 0.30–0.85), reflecting normal findings in most animals, and only a few animals showed a mild response. Similarly, all animals exhibited a severe lameness during the first two days following collagenase injection, with a median score of [S = 5 (5–5)]. Where animals were either in a recumbent position or maintained their limbs in a flexed posture while standing, lightly touching the ground with minimal weight-bearing. By the 4th day, a significant reduction in the lameness score was observed compared to day 1 ( P = 0.000), decreasing to 2 (2–3), where animals exhibited either a mild abnormal gait (S = 3) or a consistently difficult observable lameness (S = 2). Further significant improvement was noted by the 6th day ( P = 0.039), with the lameness score decreasing to 1 (1–2), representing intermittently difficult-to-detect lameness. By the 7th day, the lameness score had significantly reduced to 0 (0–1), representing a return to normal gait (S = 0) in most animals, with only a few exhibiting intermittent difficult-to-detect lameness (S = 1), with significant differences compared to day 6 ( P = 0.034) and non-significance difference compared to the normal value ( P = 0.576, r = 0.19, 95% CI − 0.15–0.50). Concerning discomfort assessments, the general appearance score revealed a mild to moderate decrease in alertness and group interaction 1 (1–2) during the first two days following collagenase injection. By the 4th day, a significant improvement was observed compared to day 1 ( P = 0.006), with the score reducing to 1 (0–1), where most animals exhibited a mild decrease in alertness and interaction with their group (S = 1), while others had returned to normal activity (S = 0). By the 6th day, the general appearance score had significantly returned to 0 [S = 0 (0–0)], representing full activity, with all animals being alert and interacting normally, with no significant difference compared to day 5 ( P = 0.276). A mild alteration in the standing posture score was observed during the first two days post-collagenase injection [S = 1 (1–1)], where animals were observed in mild weight shifting. By the 4th day, a significant improvement was noted compared to day 1 ( P = 0.000), as all animals returned to a normal standing posture [S = 0 (0–0]. A mild alteration in resting posture was observed during the first three days post-collagenase injection, as animals either attempted recumbency or limited time in a lying position (S = 1). By the 5th day, a significant improvement was noted compared to day 1 ( P = 0.000), with the score returning to the normal value [S = 0 (0–0)]. Finally, the overall discomfort score exhibited a mild increase in the first three days post collagenase injection [S = 3 (3–5)]. By the 5th day, a significant reduction was observed compared to day 1 ( P = 0.000), with scores decreasing to 0 (0–1), where only mild signs of discomfort persisted in a few animals ( n = 13/30). By the 6th day, the total discomfort score had further decreased to score 0 (0–0), indicating a complete return to normal behavior, with a significant difference compared to day 1 ( P = 0.000, r = 0.85, 95% CI 0.70–0.92) and a non-significant difference compared to the normal value ( P = 1.000, r = 0.0, 95% CI − 0.05–0.05) (Table S3, Fig. 2 ). Fig. 2. Open in a new tab Showing the clinical evaluation parameters during the tendonitis induction period (Ti1–Ti7) in comparison to the normal baseline value (N). Regardless of the biomaterials used for treatment, there was no significant difference in the clinical observation parameters between all groups ( P > 0.05) at any time point after treatment, except for intensifying weight-bearing and tendon shape scores at T20. Additionally, Time had no significant effect on these parameters in any group compared to T0 ( P > 0.05), except for intensifying weight-bearing and tendon shape scores. Regarding the pain and heat scores, the placebo and PRF/Exosome groups returned to normal scores by day 3, while the PRF group by day 4 post-treatment. In terms of lameness scores, donkeys in the PRF/Exosome group became sound by day 3 post-treatment, those in the PRF group by day 4, and those in the placebo group by day 7. At T0, all groups exhibited an irregular tendon shape with soft consistency [S = 3(3–3)], with no significant differences between them ( P = 1.000). By T20, the placebo group expressed hard nodular area in the tendon [S = 4(4–4)], while the PRF group showed minimal tendon thickening (S = 1 (0–1)), with a non-significant difference from normal values ( P = 0.149). Additionally, the PRF/Exosome group demonstrated near-complete recovery at T20, with most donkeys ( n = 7/9) achieving normal tendon shape and consistency [S = 0 (0–1)] with non-significant difference compared to normal ( P = 0.773) (Table S4, Fig. 3 A). Fig. 3. Open in a new tab Showing the clinical evaluation of palpable shape of the tendons ( A ), and intensifying weight carrying score ( B ) in both placebo and treated groups at T0 & T20. Within-group comparisons between the two time points are indicated by colored horizontal lines: black lines for the Placebo group, blue lines for the PRF group, and green lines for the PRF/Exosome group. Between-group comparisons at time point T20 are indicated by red horizontal lines. Statistical significance is denoted as follows:* p < 0.05, ** p < 0.01, *** p < 0.001, & ns = not significant. Regarding intensifying weight-bearing, all groups displayed notable offloading of the limb [S = 3 (3–3)] at T0, with no significant differences between them ( P = 1.000). At T20, the placebo group showed only a slight reduction in weight-bearing ability [S = 1 (1–1)], with no significant difference from normal values ( P = 0.083). Conversely, all PRF and PRF/Exosome groups achieved balanced weight distribution (S = 0), showing a significant improvement compared with baseline ( P < 0.05). The PRF/Exosome group fully normalized weight-bearing ability ( P = 1.000), whereas the PRF group exhibited a non-significant decrease relative to normal values ( P = 0.386) (Table S4, Fig. 3 B). Ultrasonographic findings The normal tendon cross-sectional areas (T-CSA), lesion cross-sectional areas (L-CSA), and the corresponding lesion percentages and the corresponding lesion sizes for each treatment group are summarized in Table S5. During weeks 4 and 6 (T4 & T6), the lesion % in the placebo and PRF groups continued to rise significantly ( P ≤ 0.05), reaching the highest level during this period, with no significant difference observed between the two groups ( P > 0.05). In contrast, the PRF/Exosome group exhibited only a slight, non-significant increase in lesion % at T4 compared to T0 ( P = 0.9952), followed by a gradual and steady decline by T6. From week 8 onward, all groups exhibited a gradual decline in lesion %, though at varying rates. In the placebo group, the first drop below T0 was observed at T14, although this reduction was not statistically significant ( P = 0.2065). A significant reduction in lesion % in the placebo group was first observed at T18 ( P < 0.0001). In comparison, the PRF group showed its first significant reduction below T0 as early as T10, with a marked statistically significant difference compared to the placebo group ( P < 0.0001). The PRF/Exosome group continued a steady decline after T6, with the first significant reduction below T0 occurring at T10 ( P = 0.0493) and a highly significant decrease by T12 ( P < 0.0001). The disappearance of lesions occurred at varying times across the treatment groups. In the placebo group, the anechoic core lesion had completely resolved by T18 and was replaced by hyperechoic dots. In the PRF group, complete lesion resolution was noted at T20. The PRF/Exosome group achieved complete lesion resolution by T18 (Table S6, Fig. 4 A, B). Fig. 4. Open in a new tab Showing the Lesion % ( A ), proportional change in lesion % over time relative to T0 ( B ), FES ( C ), and FAS ( D ) in both control and treated groups. Times with different small letters are significant in a group at P > 0.05. * There is a significant difference compared to the placebo group at the same time, at p < 0.05. # There is a significant difference compared to the PRF group at the same time, at p < 0.05. Concerning the tendon cross sectional area (T-CSA) during the first two weeks following treatment, all groups exhibited a significant increase ( P < 0.05) except the PRF/Exosome group which showed only slight a non-significant increase ( P = 0.981) compared to T0, with this increase showed a significantly lower value compared to the placebo ( P = 0.0001) and PRF ( P = 0.001) groups. After this initial phase, T-CSA values across all groups exhibited fluctuating patterns, alternating between increases and decreases. At T20, the placebo group showed a significant decrease in T-CSA (35.38 mm² ± 0.63) compared to T0 ( P = 0.001). The PRF and PRF/Exosome groups showed consistently higher T-CSA values than T0 throughout the study, with a slight, non-significant decrease observed at T20 (Table S7). Compared to the normal tendon size, the T-CSA at T20 remained higher in all groups to varying degrees. The placebo group showed a slight increase of 13% ( P < 0.0001). The PRF/Exosome group exhibited a mild increase of 25%, while the PRF group showed a marked increase of 34% ( P < 0.0001) relative to the normal T-CSA. Compared to T0, the FES showed a significant decrease for the first time at T10 across all groups, with varying scores: [S = 2 (2–2) in both the placebo ( P = 0.031, r = 0.70, 95% CI 0.40–0.90) and PRF groups ( P = 0.028, r = 0.73, 95% CI 0.42–0.91); and 2 (1–2) in the PRF/Exosome group ( P = 0.037, r = 0.70, 95% CI 0.40–0.90). Nearly normal echogenicity was observed at T18 in the PRF/Exosome group [S = 0.5 (0–1)], and at T20 in the PRF group [S = 0 (0–1)]. In the placebo group, the lesion was significantly replaced by hyperechoic tissue at both T18 and T20 ( P = 0.000). There was no significant difference in FES between the placebo group and both PRF and PRF/Exosome groups at most time points ( P > 0.05) except at T18 and T20 ( P < 0.05). The PRF/Exosome group showed a significant difference compared to the PRF group just at T8, 12, and T14 ( P < 0.05), where it showed a more rapid improvement in FES at those time points (Table S8, Fig. 4 C). Regarding FAS, there was no significant difference between the placebo and PRF groups at all time points ( P > 0.05). The PRF/Exosome group exhibited a significant improvement in FAS compared to the PRF and Placebo groups at T18 ( P = 0.042, r = 0.47, 95% CI 0.05–0.70). Compared to T0, the FAS in the Placebo group showed a significant decrease for the first time at T16 ( P = 0.004), representing fibers with 25–49% of the normal alignment pattern [S = 2(2–2)], and remained unchanged thereafter. This parameter decreased significantly in the PRF and PRF/Exosome groups earlier at T12 ( P < 0.05). At T20, a non-significant decrease was observed in the PRF group [S = 1.5 (1–2)] compared to T12, and the PRF/Exosome group exhibited a significant decrease [S = 1(0–1)] compared to T12, representing fibers with 50–74% of the normal alignment pattern (Table S9, Fig. 4 D). Histopathological findings Tendons from the control positive group (at day 7 post-collagenase injection; Ti7) displayed marked histological alterations, where the normal tendon structure was replaced by a dense inflammatory infiltrate composed predominantly of neutrophils, lymphocytes, and macrophages. Moreover, pronounced neovascularization was observed, accompanied by multifocal hemorrhages. The administration of regenerative biologics to the SDFT induced notable histopathological improvements. Statistical analysis revealed significant differences among treatment groups in all histopathological assessment scores at T20. Moreover, time exerted a positive effect on improving these scores across the PRF/Exosome group. However, no significant temporal changes were observed in fiber arrangement, fiber structure, and cell density within the placebo group, nor in fiber arrangement, fiber structure, and nuclear morphology within the PRF group, where values remained statistically nonsignificant compared to T0 ( P > 0.05) (Table 3 ; Fig. 5 ). Table 3. Shows results of histopathological assessment scores between the different groups. Group Evaluation times Friedman’s test P value T0 (Control positive group) T8 T14 T20 Angiogenesis Placebo 3(3–3) a 3(3–3) a 1.5(1–2) bc 1(1–1) c 17.471 0.001 PRF 3(3–3) a 3(3–3) a 3(2–3) a* 2(1–2) b* 16.286 0.001 PRF/Exosome 3(3–3) a 3(2–3) a 1(1–2) b# 1(1–2) b 16.043 0.001 Kruskal-wallis 0.000 15.046 13.065 11.764 P value 1.000 0.002 0.004 0.008 Inflammatory cell infiltration Placebo 3(3–3) a 3(3–3) a 1.5(1–2) b 1(1–1) b 17.471 0.001 PRF 3(3–3) a 3(3–3) a 1.5(1–2) b 2(1–2) b* 17.694 0.001 PRF/Exosome 3(3–3) a 3(2–3) a 1(1–2) b 1(1–2) b 16.043 0.001 Kruskal-wallis 0.000 10.767 1.874 11.764 P value 1.000 0.013 0.599 0.008 Fiber structure Placebo 3(3–3) 3(3–3) 3(3–3) 3(3–3) 0.000 1.000 PRF 3(3–3) a 3(3–3) a 3(2–3) a 2.5(2–3) a 7.364 0.061 PRF/Exosome 3(3–3) a 3(3–3) a 2(2–3) ab* 1(1–2) b*# 16.680 0.001 Kruskal-wallis 0.000 0.000 16.560 19.943 P value 1.000 1.000 0.001 0.000 Fiber arrangement Placebo 3(3–3) 3(3–3) 3(3–3) 3(3–3) 0.000 1.000 PRF 3(3–3) a 3(3–3) a 2.5(2–3) a 2(2–3) a 8.053 0.045 PRF/Exosome 3(3–3) a 3(3–3) a 1.5(1–2) b*# 1(0–1) b*# 17.471 0.001 Kruskal-wallis 0.000 0.000 16.962 20.125 P value 1.000 1.000 0.001 0.000 Rounding of nuclei Placebo 3(3–3) a 3(3–3) a 3(2–3) a 1(0–1) b 16.286 0.001 PRF 3(3–3) a 3(3–3) a 3(2–3) a 2(1–3) a* 7.333 0.062 PRF/Exosome 3(3–3) a 3(3–3) a 2(1–2) b*# 1(0–2) b 17.583 0.001 Kruskal-wallis 0.000 0.000 12.230 9.922 P value 1.000 1.000 0.007 0.019 Cell density Placebo 3(3–3) a 3(3–3) a 3(3–3) a 3(2–3) a 6.000 0.112 PRF 3(3–3) a 3(3–3) a 2.5(2–3) ab 2(2–3) b 12.000 0.007 PRF/Exosome 3(3–3) a 3(3–3) a 1.5(1–2) b*# 1(1–2) b*# 16.846 0.001 Kruskal-wallis 0.000 0.000 16.962 17.084 P value 1.000 1.000 0.001 0.001 Total histological score Placebo 3.0 ± 0.0 3.00 ± 0.00 2.5 ± 0.77 b 2.0 ± 1.09 - - PRF 3.0 ± 0.0 a 3.00 ± 0.00 a 2.5 ± 0.58 ab 2.0 ± 0.20 b - - PRF/Exosome 3.0 ± 0.0 a 3.00 ± 0.00 a 1.5 ± 0.45 b* 1.0 ± 0.00 b# - - Open in a new tab Times with different superscript letters are significantly different at p < 0.05, df = 3 *significant difference compared to the placebo group in the same time at p < 0.05, df = 3 #significant difference compared to the PRF group in the same time at p < 0.05, df = 3 Fig. 5. Open in a new tab Representative photomicrograph of tendon from different treatment groups. AT T8 ( A - D ) Placebo group showing diffuse severe replacement of tendon tissue by many blood vessels surrounded by abundant inflammatory cells composed of many neutrophils, lymphocytes, and macrophages. ( E - H ) PRF group showing focal to coalescing inflamed fibrovascular tissue with newly formed blood vessels surrounded by many lymphocytes, macrophages, and immature round fibroblasts. ( I - L ) PRF/Exosome group showing disorganized tendon with severe replacement by many blood vessels surrounded by abundant lymphocytes, plasma cells, and rounded fibroblasts. AT T14 ( Q - T ) Placebo group showing highly wavy tendon with a focal few blood vessels surrounded by low numbers of inflammatory cells as lymphocytes, plasma cells, and mild numbers of spindle fibroblasts. ( U - X ) PRF group showing less organized wavy tendon fibers with multifocal angiogenesis, few extravasated RBCs with many rounded fibroblasts, and mild numbers of inflammatory cells as lymphocytes. A1:D1) PRF/Exosome group showing little wavy arranged tendon fibers with few focal blood vessels surrounded by mild to moderate numbers of extravasated RBCs, admixed with rounded fibroblasts and a few macrophages, lymphocytes. AT T20 I1:L1 Placebo group showing disorganized tendon fibers with few angiogenesis, scattered extravasated RBCS, haphazard rounded to spindle fibroblasts. M1:P1) PRF group showing mild focal to coalescing blood vessels surrounded by moderate numbers of spindle fibroblasts. Q1:T1) PRF/Exosome group showing little wavy bundles and a few regular arrangements of tendon fibers with a few focal aggregations of inflammatory cells and spindle fibroblasts. Thin arrow = neovascularization, thick arrow = inflammation with fibroblast proliferation (either rounded = red arrowhead or spindle = black arrowhead), w = wavy tendon arrangement, hw = highly wavy disorganized collagen, lw = little wavy. Image magnification=100x, 400x. By T20, all groups showed a statistically significant reduction in inflammatory cell infiltration and angiogenesis compared to T0, although the extent of improvement varied (Placebo: P = 0.003, r = 0.97, 95% CI 0.60–0.99; PRF: P = 0.01, r = 0.83, 95% CI 0.50–0.96; PRF/Exosome: P = 0.007, r = 0.90, 95% CI 0.55–0.98). The placebo group recorded the lowest scores (S = 1 [1–1]), corresponding to minimal inflammatory cell infiltration and angiogenesis. The PRF/Exosome group also improved [S = 1 (1–2)] at T20, yet retained higher scores than the placebo group. In contrast, the PRF group showed the least improvement [S = 2 (1–2)], representing persistent moderate inflammation. Concerning the fiber structure of the newly formed tissue at the AOI, the PRF/Exosome group exhibited mildly to moderately fragmented fibers (S = 1 [1–2]), with a significant improvement from T0 ( P = 0.002, r = 1.03, 95% CI 0.65–1.0). In contrast, the PRF group displayed moderate to severe fragmentation (S = 2.5[2–3]), with a non-significant reduction compared to T0 ( P = 0.061). The placebo group consistently showed severely fragmented fibers throughout the study period. Intergroup comparison at the end of the study revealed that the PRF/Exosome exhibited significantly better outcomes than the PRF group ( P = 0.034, r = 0.50, 95% CI 0.05–0.75) and the placebo group ( P = 0.003, r = 0.68, 95% CI 0.40–0.90). Similarly, the PRF/Exosome group exhibited notable improvement in fiber arrangement at T20 compared to T0 ( P = 0.003, r = 0.97, 95% CI 0.60–0.99), with scores reflecting slightly loose, wavy fibers (S = 1 [1–1]). The PRF group retained higher scores at T20, representing moderately loose to completely disorganized fiber architecture [S = 2 (2–3)]. The placebo group consistently showed severely disrupted fiber arrangement throughout the study period [S = 3 (3–3)], with no identifiable fiber pattern. At T20, the PRF/Exosome group demonstrated significantly improved scores compared to the PRF group ( P = 0.021, r = 0.54, 95% CI 0.10–0.75) and the placebo group ( P = 0.001, r = 0.73, 95% CI 0.40–0.90). At T20, the PRF/Exosome group exhibited significantly improved nuclear morphology compared to T0 ( P = 0.001, r = 1.03, 95% CI 0.65–1.0), with scores ranging from spindle-shaped to moderately rounded nuclei (S = 1 [0–2]), but did not differ significantly from the PRF group ( P = 0.071). In contrast, the PRF group maintained higher scores (S = 2 [1–3]), representing the persistence of slightly to severely rounded nuclei, and differed significantly from the placebo group ( P = 0.004, r = 0.68, 95% CI 0.40–0.90). The placebo group at T20 showed significant improvement from T0 ( P = 0.004, r = 0.97, 95% CI 0.60–0.99), with lower scores (S = 1 [0–1]) representing predominantly spindle-shaped or slightly rounded nuclei, and no significant differences were observed when compared to the PRF/Exosome groups ( P = 0.277). Compared to the pattern of cell density observed at T0, a significant improvement was detected at T20 in the PRF/Exosome groups ( P = 0.004, r = 0.97, 95% CI 0.60–0.99), with scores reflecting a slight to moderate increase in cellularity (S = 1 [1–2]). In contrast, both the PRF and placebo groups showed higher scores, demonstrating a moderate to severe increase in cell density (S = 2 [2–3] and 3 [2–3], respectively). Intergroup comparisons at T20 revealed significantly improved scores in the PRF/Exosome groups relative to both the PRF group ( P = 0.017, r = 0.57, 95% CI 0.10–0.75) and the placebo group ( P = 0.001, r = 0.73, 95% CI 0.40–0.90). Across the study timeline, all groups showed a gradual improvement in total histological scores from T8 to T20. The PRF/Exosome group showed the most notable improvement (1.0 ± 0.00) at T20, significantly differing from the PRF group ( P = 0.027, r = 0.52, 95% CI 0.05–0.75). In contrast, the PRF and placebo groups retained higher scores at T20 (2.0 ± 0.20 and 2.0 ± 1.09), with no significant difference between them ( P = 0.603). Immunohistochemical findings Time exerted a consistent and significant effect in reducing collagen type III area percentage across all groups ( P < 0.05). At T20, the PRF/Exosome group exhibited the lowest collagen III content (0.39 ± 0.16%), which was significantly lower than both the PRF group (1.65 ± 0.12%, P < 0.0001) and the placebo group (10.89 ± 0.45%, P < 0.0001). Nevertheless, all treated tendons still showed significantly higher collagen III levels at T20 compared to the normal values (Table S10, Fig. 6 A). Fig. 6. Open in a new tab Immunohistochemical semi-quantitative analysis of collagen type III ( A ) and collagen type I ( B ) in SDFT sections at different time points across treatment groups. Time had a significant effect in increasing the collagen type I area percentage across all treatment groups ( P < 0.05). At T8, the placebo group showed minimal collagen type I content (0.022 ± 0.005%), with no significant change at T14 (0.020 ± 0.014%, P = 0.985). In contrast, the PRF (0.92 ± 0.11%, P = 0.0056) and PRF/Exosome (0.13 ± 0.05%, P = 0.0002) groups demonstrated significantly higher levels compared to the placebo group. By T20, collagen type I markedly increased in all biomaterials treated groups, with significantly higher values recorded in the PRF/Exosome (21.93 ± 0.79%, P < 0.0001) group, followed by the PRF group (10.17 ± 0.64%, P = 0.0024), in comparison to the placebo group (0.494 ± 0.122%) (Table S10, Fig. 6 B). All groups exhibited significantly lower collagen type I values compared to the normal tendon baseline ( P < 0.05). Biomechanical finding All treated groups exhibited significantly lower failure stress values compared to the normal tendon tissue ( P < 0.05) at T20. Among the treated groups, PRF/Exosome showed the highest failure stress values (60.24 ± 3.5 MPa), with a significant difference compared to the placebo group (44.7 ± 3.5 MPa, P < 0.0001) and a non-significant difference compared to the PRF group (55.3 ± 5.7 MPa, P = 0.098). At T20, the mean (± SD) strain % was highest in the PRF/Exosome group (42.9 ± 5.5%), significantly greater than the PRF (23.6 ± 6.1%) and placebo (15.32 ± 5.3%) groups ( P < 0.05). When compared to the normal tendon (59.5 ± 7.9%), strain values remained significantly lower in all groups ( P < 0.05) (Table S11, Fig. 7 ). Fig. 7. Open in a new tab Representing the mean ± SD of failure load (N), failure stress (MPa), and strain (%) of SDFT specimens at T20 across experimental groups. Groups with different superscript letters differ significantly ( P < 0.05). Discussion This study assessed the regenerative potential of I-PRF, alone or combined with MSCs-derived exosomes, in a collagenase‑induced SDF tendonitis donkey model. Donkey SDFT lesions resemble the horse condition in clinical presentation, lesion morphology, and prolonged healing 56 . Although species‑ and use‑related differences exist, most notably in tendon dimensions, biomechanical properties, and pain‑related behaviour 57 , 58 , the fundamental anatomical architecture and load‑sharing function of the horses and donkeys’ SDFT remain highly conserved 59 . These similarities allow careful extrapolation of the current outcomes to horses 60 . Moreover, as athletic tendinopathies in humans share similar pathogenic mechanisms, this large-animal model also provides relevant translational insight for sports medicine applications 61 . This prospective, blinded, placebo-controlled experimental study demonstrated the superior therapeutic efficacy of intralesional I-PRF combined with MSC-derived exosomes (PRF/Exosome group) in promoting healing of collagenase-induced SDFT injuries in donkeys. Compared to the placebo group, this combination therapy significantly reduced collagen type III expression, enhanced collagen type I deposition, improved fiber echogenicity and alignment, and enhanced biomechanical performance. Moreover, the PRF/Exosome group outperformed I-PRF alone in improving the collagen I/III ratio and total histological score. I-PRF alone also conferred marked benefits compared to the placebo group, including enhancement of fiber echogenicity, a more favorable collagen I/III balance, and increased failure stress. In contrast, placebo-treated tendons exhibited disrupted architecture, hyperechoic nodularity, disorganized fibrotic matrix, and diminished mechanical strength, suggesting an inferior fibrotic healing. In this study, clinical signs were assessed spending semi-quantitative clinical grading systems, although practical, may not provide complete accuracy. Incorporating advanced techniques, such as computerized gait analysis or thermography 62 , 63 , could have offered a more objective and detailed evaluation of both inflammation severity and tendon healing. The daily clinical assessments conducted during the tendonitis induction period offer valued visions into the development of inflammation. The observed peak in pain, heat, lameness, and discomfort scores within the first few days aligns with the expected acute inflammatory response triggered by collagenase-induced tissue disruption, which leads to the rapid onset of clinical signs. This pattern is consistent with other findings, which observed similar acute inflammatory responses post-induction 64 . The significant improvement in these parameters, reaching normal or near-normal points by the end of the first week, suggests that the acute phase had largely subsided before treatment initiation. Consequently, following treatment, no significant differences in these parameters were observed among groups, as these parameters primarily reflect the transient acute inflammatory phase, which had already resolved across all groups before intervention 10 . These results are consistent with previous research on both naturally occurring and experimentally induced SDF tendonitis in donkeys and horses 60 , 65 . Nevertheless, some studies have reported differing outcomes, observing sustained mild elevations in these clinical parameters following treatment 66 – 68 . Ultrasound remains a dependable device for noticing and spotting tendon injuries by assessing lesion size, location, and extent 69 . However, its accuracy is operator-dependent and influenced by probe pressure and measurement consistency 70 . Without strict standardization, lesion dimensions may be underestimated. Advanced imaging modalities such as MRI and CT offer greater objectivity in evaluating tendon healing 71 . In ultrasonographic assessment of tendon healing, relying on actual lesion size has major limitations in both longitudinal follow-up within the same group and intergroup comparisons. Tendon dimensions change dynamically during inflammation and remodeling 72 , and apparent lesion enlargement may reflect overall tendon swelling rather than true pathological progression. Additionally, natural variability in tendon size across individuals 69 can affect the clinical interpretation of lesion size; what appears mild in a large tendon may be severe in a smaller one 73 . To minimize these limitations, lesion percentage was used as a standardized parameter, allowing adjustment for individual tendon size and enabling more reliable intra-group comparisons 67 . Because baseline lesion percentages differed among groups, intergroup comparisons were based on relative changes rather than absolute values 52 . Baseline values (T0) were normalized within each group, and subsequent time points were expressed as ratios relative to T0. However, despite normalization, baseline variability in lesion percentage and T-CSA may still influence the magnitude of the observed healing response, as tendons with greater initial injury severity may exhibit different relative recovery patterns than those with milder lesions, potentially limiting direct intergroup comparisons. This should be considered when interpreting the relative treatment effects. A persistent increase in lesion percentage and T-CSA during the repair phase indicates continued tissue damage caused by ongoing excessive inflammation. While the initial inflammatory response facilitates tissue clearance and healing by releasing growth factors and cytokines 29 , prolonged excessive inflammation disrupts collagen synthesis and shifts its production toward the weaker type III collagen instead of the stronger type I, leading to misaligned scar tissue 74 . Notably, the pattern of these increases varied significantly between groups, underscoring the differential efficacy of the treatments in regulating inflammation and ECM turnover to promote effective and controlled tendon repair. The placebo group exhibited a significant increase in Lesion % and T-CSA up to week 8, reflecting the natural inflammatory reaction to tendon injury. In contrast, the PRF group demonstrated a slight reduction in lesion % during the first two weeks, indicating early stabilization. This effect may be related to mechanisms reported in previous studies, including PRF’s potential role in promoting macrophage polarization from the pro-inflammatory M1 to the reparative M2 phenotype and downregulation of IL-1β and IL-6 75 , 76 . However, due to PRF’s limited duration of cytokine release (up to 14 days) 18 , lesion % increased again at weeks 4 to 8, approaching levels similar to the placebo group. The PRF/Exosome group showed a slight, non-significant increase in lesion % up to week 4, followed by a gradual, non-significant decrease at weeks 6 and 8; this trend may reflect a more regulated healing response than both the placebo and PRF groups, as suggested by previous studies reporting potential immunomodulatory effects of MSC-derived exosomes, including modulation of macrophage polarization and downregulation of pro-inflammatory cytokines 42 , 77 . Importantly, these differences were mirrored by histological findings at week 8 post-treatment, where the PRF/Exosome group revealed a non-significant reduction in inflammatory markers compared to placebo and PRF groups, while both the latter retained the highest histological inflammation scores, reflecting persistent and more pronounced inflammatory activity. The timing of lesion resolution different markedly within groups, further underlining the difference effectiveness of the treatments. In the placebo group, the delayed anechoic lesion resolution observed at week 18 indicates a prolonged inflammatory and proliferative phase typical of untreated tendon injuries. Similar findings were reported by other studies 78 , 79 , who noted a persistent lesion expansion up to 16 weeks in untreated tendons. The PRF/Exosome group achieved lesion resolution slightly by T18, yet demonstrated a steady decline in lesion percentage throughout the study, with a significant improvement over the PRF and placebo groups. This pattern suggests a more regulated healing response than the placebo and PRF groups 80 , 81 . Similarly, the PRF group exhibited lesion resolution by the end of the study, likely due to its limited ability to sufficiently regulate early inflammation compared to the combination therapy 82 . The combined use of ultrasonographic, histological, and immunohistochemical assessments provided an integrated evaluation of tendon repair quality. In the placebo group, initial anechoic defects progressed to hyperechoic, structurally disorganized patterns consistent with fibrotic scar formation, a finding supported by palpable nodular consistency, histological evidence of collagen misalignment and a predominance of type III collagen. In contrast, the PRF/Exosome group demonstrated progressive structural improvement with lesion resolution by T18 and a favorable collagen I/III profile at T20, indicating more advanced matrix maturation despite mild residual enlargement. The PRF group exhibited intermediate findings, with persistent remodeling features and lower collagen type I expression compared to the combination therapy, suggesting incomplete structural restoration. The changeable T-CSA pattern detected across groups reflects the dynamic nature of tendon healing, encompassing inflammation, cellular proliferation, and matrix remodeling 83 . Early increases corresponded to inflammatory edema and cellular infiltration, whereas later variations likely reflected ongoing extracellular matrix turnover and tissue reorganization. Notably, T-CSA remained above normal levels at the study endpoint in all groups, consistent with previous reports 51 , 52 , 65 . Although the placebo group approached near-normal dimensions, the PRF and PRF/Exosome groups exhibited greater residual enlargement (34% and 25%, respectively), with the PRF group demonstrating significantly higher T-CSA at T20. This persistent enlargement suggests ongoing remodeling rather than complete structural normalization. The absence of significant differences in lameness scores among treated groups should not be interpreted as equivalent functional recovery. Routine clinical assessment during walk and trot may underestimate biomechanical deficits that become evident under higher mechanical loading, such as athletic activity 84 . Moreover, lameness alone is an insensitive indicator of tendon repair quality, as chronic tendon pain is often minimal in equine species 85 . Although the recurrence rate of tendon injury has been suggested as a more reliable parameter for assessing long-term functionality 86 , the improved intensifying weight-bearing symmetry observed in combination-treated groups at the end of this study suggests enhanced mechanical integrity. However, more sensitive assessment tools, particularly direct tendon tensile strength measurements, remain essential for a definitive assessment of tendon repair quality 46 , 87 . Accordingly, both combination therapies exhibited significantly higher biomechanical performance compared to the PRF and placebo groups, with no statistical difference between them. These findings are consistent with previous studies, where MSCs-based therapies significantly improved tendon biomechanics compared to controls, although biomechanical values remained below those of native tendon tissue 51 , 85 . In this study, IHC evaluation was not restricted to visual inspection; instead, quantitative assessment was performed using digital morphometric analysis with image-processing software to calculate the area percentage of positively stained regions 88 . This semi-quantitative method provides a reliable estimation of collagen distribution within defined tissue zones, although it does not reflect absolute protein concentrations 89 . The present findings demonstrate a clear correspondence between the IHC profile and the biomechanical behavior of the repaired tendon tissue. Increased collagen type I expression in the combination-treated group at T20 paralleled its superior mechanical properties, reflecting more advanced matrix maturation and functional restoration 90 . These results are in line with previous reports showing that these therapies enhance collagen type I deposition and mechanical competence 51 , 85 , although variability in IHC outcomes has been described in some studies 67 . Collagenase-induced tendonitis remains one of the most widely accepted and biologically relevant models for equine tendon injury research 46 , 51 , 60 , 91 . Although it does not fully replicate the progressive microdamage observed in naturally occurring cases, it reliably reproduces key pathological features, including central core lesions, matrix degradation, and a pronounced inflammatory response 51 . A recognized limitation is variability in lesion size among animals. In contrast, surgically induced models offer greater uniformity but often lack the characteristic inflammatory and degradative responses of spontaneous tendonitis 48 , 92 . While the influence of bacterial collagenase on the efficacy of intralesional biomaterials remains unclear, several studies have reported successful application of biologics 7–12 days after induction 51 , 78 . We acknowledge that direct in-house characterization of the MSC-derived exosomes was not performed in this study. All animals received exosomes from the same production batch, ensuring batch-to-batch consistency. Biological quality and functional integrity were further supported by previously published studies using the same product 93 , 94 and by the observed effects in both the current pilot and experimental study. Bone marrow-derived stem cells were selected as the source of exosomes due to their suitability for tendon regeneration 95 . Their exosomes have been reported in previous studies to enhance tenogenic gene expression and potentially modulate inflammation by promoting M2 macrophages and upregulating IL-4 and IL-10 to limit scar formation 40 , 96 , as well as carrying miRNAs that support extracellular matrix remodeling, thereby contributing to improved tendon structure and function 44 . Effective doses of MSC-derived exosomes in preclinical tendon models have ranged from 100 to 200 µg per lesion in rodents to approximately 500 µg per lesion in larger animals such as dogs 37 , 93 , 94 . No standardized dose has been established for equine tendons, and in this study, the exosome dose was determined based on manufacturer instructions and previous studies using the same product 94 . Nonetheless, it would have been preferable to conduct a study to evaluate the response to different doses and determine the optimal therapeutic concentration. Nonetheless, these methodological constraints should be considered when interpreting the findings. While long-term follow-up studies are essential to evaluate re-injury rates and safe return to function, the current 20-week study was specifically designed to capture the key stages of tendon healing, including inflammation resolution, matrix remodeling, and structural maturation. This timeframe aligns with previous experimental studies and allows for a comprehensive assessment of tissue quality 51 , 97 . Conclusion In this collagenase-induced tendonitis model in donkeys, intralesional administration of I-PRF combined with MSC-derived exosomes was associated with greater structural and functional improvement compared to the saline-treated group, as reflected by improved collagen fiber alignment, increased collagen type I deposition, decreased collagen type III expression, and enhanced tensile strength. PRF/Exosome-treated tendons also showed reduced T-CSA and better total histological scores and collagen expression compared to I-PRF therapy alone. These findings suggest that the combined biologic approach may contribute to tendon matrix remodeling and functional repair, but the underlying mechanisms remain speculative. Further studies, particularly in naturally occurring cases, are warranted to confirm these effects and to evaluate long-term rehabilitation outcomes. Study limitation The absence of long-term follow-up assessing re-injury rates and functional recovery limits the direct translational relevance of these findings to clinical practice in equine or human athletes. Future studies on naturally occurring tendonitis with long-term follow-up may help determine the durability of these findings and their implications for athletic return. Supplementary Information Below is the link to the electronic supplementary material. Supplementary Material 1 (34.7KB, docx) Author contributions GK, EM, AR, AS, and MN played a role in the conception and design of the study. MN, AS, and AR conducted the experimental procedures. MN collected the data. Histological processing and interpretation were conducted by II. MN and AS drafted the manuscript. AS, AR, EM, and GK thoroughly reviewed the manuscript and approved the final version. Funding Open access funding provided by The Science, Technology & Innovation Funding Authority (STDF) in cooperation with The Egyptian Knowledge Bank (EKB). Data availability All data produced or analyzed in this study are provided within this published article and its supplementary materials, including detailed results presented in the attached supplementary tables. Declarations Competing interests The authors declare no competing interests. Ethics The Medical Research Ethics Committee at the Faculty of Veterinary Medicine, Mansoura University, granted approval for this study (Approval Code: Ph.D./101). All procedures were performed in accordance with the relevant institutional guidelines and national regulations. The study was conducted and reported in accordance with the ARRIVE guidelines. Consent for publication All authors have provided their consent for publication. Footnotes Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. References 1. Olivier, A., Nurton, J. & Guthrie, A. An epizoological study of wastage in thoroughbred racehorses in Gauteng, South Africa. J. S. Afr. Vet. Assoc. 68 (4), 125–129 (1997). [ DOI ] [ PubMed ] [ Google Scholar ] 2. Lam, K., Parkin, T., Riggs, C. & Morgan, K. Descriptive analysis of retirement of Thoroughbred racehorses due to tendon injuries at the Hong Kong Jockey Club (1992–2004). Equine Vet. J. 39 (2), 143–148 (2007). [ DOI ] [ PubMed ] [ Google Scholar ] 3. Dabbene, I. et al. Clinical findings and prognosis of interference injuries to the palmar aspect of the forelimbs in Standardbred racehorses: A study on 74 cases. Equine Vet. J. 50 (6), 759–765 (2018). [ DOI ] [ PubMed ] [ Google Scholar ] 4. Bertuglia, A., Bullone, M., Rossotto, F. & Gasparini, M. Epidemiology of musculoskeletal injuries in a population of harness Standardbred racehorses in training. BMC Vet. Res. 10 (1), 1–10 (2014). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 5. Alzola, R., Easter, C., Riggs, C., Gardner, D. & Freeman, S. Ultrasonographic-based predictive factors influencing successful return to racing after superficial digital flexor tendon injuries in flat racehorses: a retrospective cohort study in 469 Thoroughbred racehorses in Hong Kong. Equine Vet. J. 50 (5), 602–608 (2018). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 6. Citro, V. et al. Tendon tissue engineering: An overview of biologics to promote tendon healing and repair. J. tissue Eng. 14 (1), 1–24 (2023). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 7. Dyson, S. Medical management of superficial digital flexor tendonitis: A comparative study in 219 horses (1992-2000). Equine Vet. J. 36 (5), 415–419 (2004). [ DOI ] [ PubMed ] [ Google Scholar ] 8. O’meara, B., Bladon, B., Parkin, T., Fraser, B. & Lischer, C. An investigation of the relationship between race performance and superficial digital flexor tendonitis in the Thoroughbred racehorse. Equine Vet. J. 42 (4), 322–326 (2010). [ DOI ] [ PubMed ] [ Google Scholar ] 9. Tamura, N. et al. A retrospective cohort study investigating risk factors for the failure of Thoroughbred racehorses to return to racing after superficial digital flexor tendon injury. Vet. J. 235 , 42–46 (2018). [ DOI ] [ PubMed ] [ Google Scholar ] 10. Smith, R. K. Treatment of tendinopathies. Equine Veterinary Educ. 36 (12), 659–672 (2024). [ Google Scholar ] 11. Kovac, M. et al. Gene therapy using plasmid DNA encoding VEGF164 and FGF2 genes: a novel treatment of naturally occurring tendinitis and desmitis in horses. Front. Pharmacol. 9 , 978 (2018). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 12. Willow, R., Guzmán, K. E., Panek, C. L. & Colbath, A. C. Stem cells and platelet-rich plasma for the treatment of naturally occurring equine tendon and ligament injuries: A systematic review and meta-analysis. J. Am. Vet. Med. Assoc. 262 (S1), S50–S60 (2024). [ DOI ] [ PubMed ] [ Google Scholar ] 13. Roshini, S. T., Arunprasad, A., William, B. J., Jeyaraja, K. & Priyadharshini, K. Autologous platelet rich plasma for regeneration of tendon injuries in horses. Indian J. Anim. Res. 53 (10), 1354–1360 (2019). [ Google Scholar ] 14. Chalidis, B., Givissis, P., Papadopoulos, P. & Pitsilos, C. Molecular and biologic effects of platelet-rich plasma (PRP) in ligament and tendon healing and regeneration: A systematic review. Int. J. Mol. Sci. 24 (3), 2744 (2023). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 15. Bonilla-Gutiérrez, A. F., López, C. & Carmona, J. U. Regenerative therapies for the treatment of tenodesmic injuries in horses. J. Equine Veterinary Sci. 73 , 139–147 (2019). [ Google Scholar ] 16. Carmona, J. U. & López, C. Efficacy of Platelet-Rich Plasma in the Treatment of Equine Tendon and Ligament Injuries: A Systematic Review of Clinical and Experimental Studies. Veterinary Sci. 12 (4), 382 (2025). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 17. Montano, C. et al. The use of platelet-rich plasma for treatment of tenodesmic lesions in horses: A systematic review and meta-analysis of clinical and experimental data. Animals 11 (3), 793 (2021). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 18. Zwittnig, K. et al. Growth factor release within liquid and solid PRF. J. Clin. Med. 11 (17), 5070 (2022). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 19. Zuleika, P., Saleh, I., Murti, K., Liberty, I. A. & Surono, A. Platelet-rich fibrin: A systematic review of its action. J. Nat. Sci. Med. 7 (4), 242–252 (2024). [ Google Scholar ] 20. Farmani, A. R. et al. Application of platelet rich fibrin in tissue engineering: focus on bone regeneration. Platelets 32 (2), 183–188 (2021). [ DOI ] [ PubMed ] [ Google Scholar ] 21. Wong, C-C. et al. Cytokine and growth factor delivery from implanted platelet-rich fibrin enhances rabbit Achilles tendon healing. Int. J. Mol. Sci. 21 (9), 3221 (2020). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 22. Yousefi, A. R., Kazemi, D. & Rezaei, M. Effect of Platelet-Rich Fibrin on Biomechanical and Biochemical Aspects Of Tendon Healing: A Preliminary Animal Study. J. Musculoskelet. Res. 25 (03), 2250016 (2022). [ Google Scholar ] 23. Abd Al-Hussein, S. S., Al-Dirawi, A. A. & Al-Khalifah, R. M. Histopathological Study of the Effect of Xenogeneic Platelet-Rich Fibrin on Achilles Tendon Healing in Rabbit. Revista Bionatura . 8 (2), 63 (2023). [ Google Scholar ] 24. Abdelhakiem, M. A., Hussein, A., Seleim, S. M., Abdelbaset, A. E. & Abd-Elkareem, M. Silver nanoparticles and platelet-rich fibrin accelerate tendon healing in donkey. Sci. Rep. 13 (1), 3421 (2023). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 25. Dakin, S. A review of the healing processes in equine superficial digital flexor tendinopathy. Equine Veterinary Educ. 29 (9), 516–520 (2017). [ Google Scholar ] 26. Mosca, M. J. et al. Trends in the theory that inflammation plays a causal role in tendinopathy: A systematic review and quantitative analysis of published reviews. BMJ Open. Sport. Exerc. Med. 4 (1), e000332 (2018). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 27. Najafi, Z., Rahmanian-Devin, P., Baradaran Rahimi, V., Nokhodchi, A. & Askari, V. R. Challenges and opportunities of medicines for treating tendon inflammation and fibrosis: A comprehensive and mechanistic review. Fundam. Clin. Pharmacol. 38 (5), 802–841 (2024). [ DOI ] [ PubMed ] [ Google Scholar ] 28. Chisari, E., Rehak, L., Khan, W. S. & Maffulli, N. Tendon healing in presence of chronic low-level inflammation: a systematic review. Br. Med. Bull. 132 (1), 97–116 (2019). [ DOI ] [ PubMed ] [ Google Scholar ] 29. Arvind, V. & Huang, A. H. Reparative and maladaptive inflammation in tendon healing. Front. Bioeng. Biotechnol. 9 (1), 719047 (2021). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 30. Fang, Y. et al. Collagen denaturation in post-run Achilles tendons and Achilles tendinopathy: In vivo mechanophysiology and magnetic resonance imaging. Sci. Adv. 10 (40), eado2015 (2024). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 31. Ackerman, J. E., Best, K. T., Muscat, S. N. & Loiselle, A. E. Metabolic regulation of tendon inflammation and healing following injury. Curr. Rheumatol. Rep. 23 (3), 1–9 (2021). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 32. Russo, V. et al. Tendon immune regeneration: Insights on the synergetic role of stem and immune cells during tendon regeneration. Cells 11 (3), 434 (2022). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 33. Chisari, E., Rehak, L., Khan, W. S. & Maffulli, N. The role of the immune system in tendon healing: a systematic review. Br. Med. Bull. 133 (1), 49–64 (2020). [ DOI ] [ PubMed ] [ Google Scholar ] 34. Tarafder, S. et al. Tendon stem/progenitor cells regulate inflammation in tendon healing via JNK and STAT3 signaling. FASEB J. 31 (9), 3991 (2017). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 35. Vinhas, A., Almeida, A. F., Gonçalves, A. I., Rodrigues, M. T. & Gomes, M. E. Magnetic stimulation drives macrophage polarization in cell to–cell communication with Il-1β primed tendon cells. Int. J. Mol. Sci. 21 (15), 5441 (2020). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 36. Wang, Y., Lu, X., Lu, J., Hernigou, P. & Jin, F. The role of macrophage polarization in tendon healing and therapeutic strategies: Insights from animal models. Front. Bioeng. Biotechnol. 12 , 1366398 (2024). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 37. Lu, V., Tennyson, M., Zhang, J. & Khan, W. Mesenchymal stem cell-derived extracellular vesicles in tendon and ligament repair—A systematic review of in vivo studies. Cells 10 (10), 2553 (2021). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 38. Chamberlain, C. S. et al. Extracellular vesicle-educated macrophages promote early Achilles tendon healing. Stem Cells 37 (5), 652–662 (2019). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 39. Shi, Y. et al. Exosomes derived from bone marrow stromal cells (BMSCs) enhance tendon-bone healing by regulating macrophage polarization. Med. Sci. monitor: Int. Med. J. experimental Clin. Res. 26 , e923328–e923321 (2020). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 40. Shi, Z., Wang, Q. & Jiang, D. Extracellular vesicles from bone marrow-derived multipotent mesenchymal stromal cells regulate inflammation and enhance tendon healing. J. translational Med. 17 (1), 211 (2019). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 41. Shen, H., Yoneda, S., Abu-Amer, Y., Guilak, F. & Gelberman, R. H. Stem cell‐derived extracellular vesicles attenuate the early inflammatory response after tendon injury and repair. J. Orthop. Res. 38 (1), 117–127 (2020). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 42. Zhang, M. et al. Tendon stem cell-derived exosomes regulate inflammation and promote the high-quality healing of injured tendon. Stem Cell Res. Ther. 11 (1), 1–15 (2020). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 43. Wang, Y. et al. Exosomes from tendon stem cells promote injury tendon healing through balancing synthesis and degradation of the tendon extracellular matrix. J. Cell. Mol. Med. 23 (8), 5475–5485 (2019). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 44. Yu, H. et al. Bone marrow mesenchymal stem cell-derived exosomes promote tendon regeneration by facilitating the proliferation and migration of endogenous tendon stem/progenitor cells. Acta Biomater. 106 , 328–341 (2020). [ DOI ] [ PubMed ] [ Google Scholar ] 45. Xu, T. et al. Comparative effects of exosomes and ectosomes isolated from adipose-derived mesenchymal stem cells on achilles tendinopathy in a rat model. Am. J. Sports Med. 50 (10), 2740–2752 (2022). [ DOI ] [ PubMed ] [ Google Scholar ] 46. Bernardi, N. S. et al. Applicability of ARFI elastography in detecting elasticity changes of the equine superficial digital flexor tendon with induced injury. Veterinary Radiol. 63 (6), 790–797 (2022). [ DOI ] [ PubMed ] [ Google Scholar ] 47. Miron, R. J. et al. Injectable platelet rich fibrin (i-PRF): opportunities in regenerative dentistry? Clin. Oral Invest. 21 , 2619–2627 (2017). [ DOI ] [ PubMed ] [ Google Scholar ] 48. Depuydt, E. et al. The evaluation of equine allogeneic tenogenic primed mesenchymal stem cells in a surgically induced superficial digital flexor tendon lesion model. Front. veterinary Sci. 8 , 641441 (2021). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 49. Reix, C., Burn, C., Pritchard, J., Barr, A. & Whay, H. The range and prevalence of clinical signs and conformation associated with lameness in working draught donkeys in P akistan. Equine Vet. J. 46 (6), 771–777 (2014). [ DOI ] [ PubMed ] [ Google Scholar ] 50. van Loon, J. P. et al. Objective assessment of chronic pain in donkeys using the donkey chronic pain scale (DCPS): A scale-construction study. Vet. J. 267 , 105580 (2021). [ DOI ] [ PubMed ] [ Google Scholar ] 51. Gaesser, A. M. et al. Evaluation of autologous protein solution injection for treatment of superficial digital flexor tendonitis in an equine model. Front. Veterinary Sci. 8 (1), 697551 (2021). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 52. Burk, J. et al. Treatment of naturally occurring tendon disease with allogeneic multipotent mesenchymal stromal cells: a randomized, controlled, triple-blinded pilot study in horses. Cells 12 (21), 2513 (2023). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 53. Movin, T., Gad, A., Reinholt, F. P. & Rolf, C. J. Tendon pathology in long-standing achillodynia: Biopsy findings in 40 patients. Acta Orthop. Scand. 68 (2), 170–175 (1997). [ DOI ] [ PubMed ] [ Google Scholar ] 54. Helps, S. C., Thornton, E., Kleinig, T. J., Manavis, J. & Vink, R. Automatic nonsubjective estimation of antigen content visualized by immunohistochemistry using color deconvolution. Appl. Immunohistochem. Mol. Morphology . 20 (1), 82–90 (2012). [ DOI ] [ PubMed ] [ Google Scholar ] 55. Ruan, D. et al. Early-stage primary anti-inflammatory therapy enhances the regenerative efficacy of platelet-rich plasma in a rabbit achilles tendinopathy model. Am. J. Sports Med. 49 (12), 3357–3371 (2021). [ DOI ] [ PubMed ] [ Google Scholar ] 56. O’Brien, C., Marr, N. & Thorpe, C. Microdamage in the equine superficial digital flexor tendon. Equine Vet. J. 53 (3), 417–430 (2021). [ DOI ] [ PubMed ] [ Google Scholar ] 57. Burden, F. & Thiemann, A. Donkeys are different. J. Equine Vet. Sci. 35 (5), 376–382 (2015). [ Google Scholar ] 58. Ribitsch, I. et al. Age-related changes of tendon fibril micro‐morphology and gene expression. J. Anat. 236 (4), 688–700 (2020). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 59. Hanot, P., Herrel, A., Guintard, C. & Cornette, R. Morphological integration in the appendicular skeleton of two domestic taxa: the horse and donkey. Proc. Royal Soc. B: Biol. Sci. 284 (1864), 20171241 (2017). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 60. Alaei, M. H. et al. Collagenase Induced Tendinitis in Forelimb Superficial Digital Flexor Tendon in Donkey (Equus asinus): Long Term Ultrasonographic and Biomechanical Assessments. Iran. J. Veterinary Surg. 16 (1), 47–55 (2021). [ Google Scholar ] 61. Shojaee, A. Equine tendon mechanical behaviour: Prospects for repair and regeneration applications. Vet. Med. Sci. 9 (5), 2053–2069 (2023). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 62. Hardeman, A. M. et al. Visual lameness assessment in comparison to quantitative gait analysis data in horses. Equine Vet. J. 54 (6), 1076–1085 (2022). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 63. Okur, S. et al. The effectiveness of thermography in determining localization of orthopedic diseases in horses. Van Veterinary J. 34 (1), 51–54 (2023). [ Google Scholar ] 64. Watts, A., Nixon, A., Yeager, A. & Mohammed, H. A collagenase gel/physical defect model for controlled induction of superficial digital flexor tendonitis. Equine Vet. J. 44 (5), 576–586 (2012). [ DOI ] [ PubMed ] [ Google Scholar ] 65. Geburek, F., Gaus, M., van Schie, H. T., Rohn, K. & Stadler, P. M. Effect of intralesional platelet-rich plasma (PRP) treatment on clinical and ultrasonographic parameters in equine naturally occurring superficial digital flexor tendinopathies–a randomized prospective controlled clinical trial. BMC Vet. Res. 12 , 1–16 (2016). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 66. Oloumi, M. M., Vosough, D., Derakhshanfar, A. & Nematollahi, M. H. The healing potential of Plantago lanceolata ointment on collagenase-induced tendinitis in burros (Equus asinus). J. Equine Veterinary Sci. 31 (8), 470–474 (2011). [ Google Scholar ] 67. Geburek, F., Lietzau, M., Beineke, A., Rohn, K. & Stadler, P. M. Therapy: Effect of a single injection of autologous conditioned serum (ACS) on tendon healing in equine naturally occurring tendinopathies. Stem cell. Res. 6 , 1–14 (2015). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 68. Shokry, A. A. M., M E-S, AA, A. T. & A T, M E-S, M. Autologous Bone Marrow-Derived Mesenchymal Stem Cells for Tendon Repair in Horses: Experimental Study. J. Biotechnol. Bioresearch . 3 (2), 1–11 (2021). [ Google Scholar ] 69. Ehrle, A., Lilge, S., Clegg, P. D. & Maddox, T. W. Equine flexor tendon imaging part 1: Recent developments in ultrasonography, with focus on the superficial digital flexor tendon. J. Veterinary J. 278 , 105764 (2021). [ DOI ] [ PubMed ] [ Google Scholar ] 70. Thoirs, K. A. & Childs, J. Are ultrasound measurements of Achilles tendon size reliable? A systematic review of rater reliability. Ultrasound Med. biology . 44 (12), 2476–2491 (2018). [ DOI ] [ PubMed ] [ Google Scholar ] 71. Johnson, S. A. et al. Longitudinal tendon healing assessed with multi‐modality advanced imaging and tissue analysis. Equine Vet. J. 54 (4), 766–781 (2022). [ DOI ] [ PubMed ] [ Google Scholar ] 72. Zellers, J. A., Cortes, D. H., Pohlig, R. T. & Silbernagel, K. G. Tendon morphology and mechanical properties assessed by ultrasound show change early in recovery and potential prognostic ability for 6-month outcomes. Knee Surg. Sports Traumatol. Arthrosc. 27 (9), 2831–2839 (2019). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 73. Mc Auliffe, S., Mc Creesh, K., Purtill, H. & O’Sullivan, K. A systematic review of the reliability of diagnostic ultrasound imaging in measuring tendon size: Is the error clinically acceptable? Phys. Ther. Sport . 26 , 52–63 (2017). [ DOI ] [ PubMed ] [ Google Scholar ] 74. Millar, N. L., Murrell, G. A. & McInnes, I. B. Inflammatory mechanisms in tendinopathy–towards translation. Nat. Rev. Rheumatol. 13 (2), 110–122 (2017). [ DOI ] [ PubMed ] [ Google Scholar ] 75. Nasirzade, J., Kargarpour, Z., Hasannia, S., Strauss, F. J. & Gruber, R. Platelet-rich fibrin elicits an anti‐inflammatory response in macrophages in vitro. J. Periodontol. 91 (2), 244–252 (2020). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 76. Zhang, J. et al. Anti-inflammation effects of injectable platelet-rich fibrin via macrophages and dendritic cells. J. Biomedical Mater. Res. Part. A . 108 (1), 61–68 (2020). [ DOI ] [ PubMed ] [ Google Scholar ] 77. Arabpour, M., Saghazadeh, A. & Rezaei, N. Anti-inflammatory and M2 macrophage polarization-promoting effect of mesenchymal stem cell-derived exosomes. Int. Immunopharmacol. 97 (1), 107823 (2021). [ DOI ] [ PubMed ] [ Google Scholar ] 78. Watts, A. E. et al. MicroRNA29a treatment improves early tendon injury. Mol. Ther. 25 (10), 2415–2426 (2017). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 79. Carlier, S. et al. Equine allogeneic tenogenic primed mesenchymal stem cells: a clinical field study in horses suffering from naturally occurring superficial digital flexor tendon and suspensory ligament injuries. Equine Vet. J. 56 (5), 924–935 (2024). [ DOI ] [ PubMed ] [ Google Scholar ] 80. Liu, H. et al. Adipose-derived mesenchymal stromal cell-derived exosomes promote tendon healing by activating both SMAD1/5/9 and SMAD2/3. Stem Cell. Res. 12 (1), 338 (2021). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 81. Trouski, F. F. & Parham, A. Exosomes Derived from Mesenchymal Stem Cells in the Treatment of Animal Tendon Injuries: A Review on Their Isolation and Application. Iran. J. Veterinary Med. 15 (3), 259–274 (2021). [ Google Scholar ] 82. Dakin, S. G. et al. Macrophage sub-populations and the lipoxin A4 receptor implicate active inflammation during equine tendon repair. PLoS One 7 (2), e32333 (2012). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 83. Leong, N. L. et al. Tendon and ligament healing and current approaches to tendon and ligament regeneration. J. Orthop. Research ® . 38 (1), 7–12 (2020). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 84. Verkade, M., Back, W. & Birch, H. Equine digital tendons show breed-specific differences in their mechanical properties that may relate to athletic ability and predisposition to injury. Equine Vet. J. 52 (2), 320–325 (2020). [ DOI ] [ PubMed ] [ Google Scholar ] 85. Geburek, F. et al. Effect of single intralesional treatment of surgically induced equine superficial digital flexor tendon core lesions with adipose-derived mesenchymal stromal cells: A controlled experimental trial. Stem Cell Res. Ther. 8 (1), 129 (2017). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 86. Beerts, C. et al. Tenogenically induced allogeneic peripheral blood mesenchymal stem cells in allogeneic platelet-rich plasma: 2-year follow-up after tendon or ligament treatment in horses. Front. Veterinary Sci. 4 (1), 158 (2017). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 87. Wagner, F. C., Reese, S., Gerlach, K., Böttcher, P. & Mülling, C. K. Cyclic tensile tests of Shetland pony superficial digital flexor tendons (SDFTs) with an optimized cryo-clamp combined with biplanar high-speed fluoroscopy. J. BMC veterinary Res. 17 (1), 223 (2021). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 88. De López, C. M. et al. Picrosirius red staining: revisiting its application to the qualitative and quantitative assessment of collagen type I and type III in tendon. J. Histochem. Cytochemistry . 69 (10), 633–643 (2021). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 89. Jacobson, K. R. et al. Comparative analysis of the extracellular matrix proteome across the myotendinous junction. J. Proteome Res. 19 (10), 3955–3967 (2020). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 90. Genç, E. et al. Investigation of the biomechanical and histopathological effects of autologous conditioned serum on healing of Achilles tendon. Acta Orthop. Traumatol. Turc. 52 (3), 226–231 (2018). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 91. Durgam, S. S., Stewart, A. A., Sivaguru, M., Wagoner Johnson, A. J. & Stewart, M. C. Tendon-derived progenitor cells improve healing of collagenase‐induced flexor tendinitis. J. Orthop. Research ® . 34 (12), 2162–2171 (2016). [ DOI ] [ PubMed ] [ Google Scholar ] 92. Schramme, M., Hunter, S., Campbell, N., Blikslager, A. & Smith, R. A surgical tendonitis model in horses: Technique, clinical, ultra-sonographic and histological characterisation. Vet. Comp. Orthop. Traumatol. 23 (04), 231–239 (2010). [ DOI ] [ PubMed ] [ Google Scholar ] 93. Elsherbiny, N. M. et al. MSCs–derived EVs protect against chemotherapy-induced ovarian toxicity: Role of PI3K/AKT/mTOR axis. J. Ovarian Res. 17 (1), 222 (2024). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 94. Rafat, A. et al. Potential regenerative effect of mesenchymal stem cells-derived microvesicles on healing of the ruptured achilles tendon in a dog model. J. Curr. Veterinary Res. 4 (2), 140–151 (2022). [ Google Scholar ] 95. Chen, J. et al. Regenerative properties of bone marrow mesenchymal stem cell derived exosomes in rotator cuff tears. J. Translational Med. 23 (1), 47 (2025). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 96. Huang, Y. et al. Bone marrow mesenchymal stem cell-derived exosomes promote rotator cuff tendon-bone healing by promoting angiogenesis and regulating M1 macrophages in rats. Stem Cell Res. Ther. 11 , 1–16 (2020). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 97. Carvalho, A. M. et al. Equine tendonitis therapy using mesenchymal stem cells and platelet concentrates: a randomized controlled trial. Stem Cell Res. Ther. 4 (4), 1–13 (2013). [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Associated Data This section collects any data citations, data availability statements, or supplementary materials included in this article. Supplementary Materials Supplementary Material 1 (34.7KB, docx) Data Availability Statement All data produced or analyzed in this study are provided within this published article and its supplementary materials, including detailed results presented in the attached supplementary tables. Articles from Scientific Reports are provided here courtesy of Nature Publishing Group ACTIONS View on publisher site PDF (2.9 MB) Cite Collections Permalink PERMALINK Copy RESOURCES Similar articles Cited by other articles Links to NCBI Databases Cite Copy Download .nbib .nbib Format: AMA APA MLA NLM Add to Collections Create a new collection Add to an existing collection Name your collection * Choose a collection Unable to load your collection due to an error Please try again Add Cancel Follow NCBI NCBI on X (formerly known as Twitter) NCBI on Facebook NCBI on LinkedIn NCBI on GitHub NCBI RSS feed Connect with NLM NLM on X (formerly known as Twitter) NLM on Facebook NLM on YouTube National Library of Medicine 8600 Rockville Pike Bethesda, MD 20894 Web Policies FOIA HHS Vulnerability Disclosure Help Accessibility Careers NLM NIH HHS USA.gov Back to Top