Tendinopathy accounts for approximately 30% of the total number of diagnosed injuries and is more common in sports that involve repetitive loading of a particular tendon or tendon group. Tendon injuries can be acute or chronic and can be influenced by a variety of factors, including increasing age, gender, type of exercise and physical activity, occupation, and certain comorbidities, including metabolic or cardiovascular disease, alone or in combination. Achilles tendinopathy (AT) is clinically defined as an overload injury to the Achilles tendon caused by continuous, prolonged, and intense functional demands, which can affect the distal insertion or the midportion of the tendon. AT is distinguished by pain and swelling in and around the tendon, reducing both quality of life and physical activity. AT management is one of the most critical challenges in orthopedic settings. Currently available treatments for AT include prolonged monitoring, physiotherapy, injectable medications, shockwave therapy, orthosis, oral medication, and surgery.[5,6] Most patients receive multiple treatments over time, consequently increasing healthcare consumption. The majority of such treatments are largely palliative and pose the risk of serious adverse effects and poor long-term benefits. Regenerative medicine is a new modality that is gaining popularity in AT treatment. Mesenchymal stem cells (MSCs) are multipotent adult stem cells that have a wide range of applications in tissue repair and regeneration. Adipose-derived stem cells have been shown to promote wound healing and scar repair.[9–12] MSCs have the ability to improve tendon healing and repair. However, the sourcing of MSCs along with the associated donor and ethical issues are major challenges; therefore, it is essential to seek novel and safe sources of stem cells for use in regenerative medicine.
Human adult stem cells are capable of improving hair follicle development and promote hair growth.[14,15] Recently, hair follicle-derived MSCs have attracted the attention of the research community. Hair follicles are easy to access and contain stem cells of various developmental origins, including MSCs. Human hair follicle-derived mesenchymal stem cells (hHF-MSCs) were identified by Liu et al as cells that express the MSC immunophenotype and exhibit multilineage differentiation potential. In single-cell culture, hHF-MSCs demonstrate the ability to differentiate into myogenic, melanocytic, and neuronal cell lineages, as well as adipocyte, chondrocyte, and osteocyte lineages. Implanted hHF-MSCs enhanced the repair of severe peripheral nerves and the spinal cord. Yari et al reported that hHF-MSC transplantation affected wound closure, epithelial thickness, collagen formation, and SDF-1α and CXCR4 expression in wound healing. These results revealed the repair and regeneration potential of hHF-MSCs. However, the role of hHF-MSCs in AT was still unclear. The present study aimed to investigate the temporal effects of hHF-MSCs on AT repair and regeneration in rabbits.
This study was approved by the Ethic Committee of Zhejiang Provincial People's Hospital and the Institutional Animal Care and Use Committee approved all the experiments designed in this study (HB2004022). Written informed consent was obtained from all participants.
Isolation and culture of hHF-MSCs
Human scalp skin was obtained under sterile conditions via skin biopsy from a donor, and hHF-MSCs were isolated using a previously described protocol. First, skin tissues were rinsed with phosphate buffered saline (PBS) containing 1% penicillin/streptomycin; then, the underlying adipose tissues were removed, the tissue was dissected into 24 mm segments and digested using 1 mg/mL collagenase type I at 37°C. Then, after 4 h, the epidermis was peeled away from the dermal layer, and single-hair follicles were released from the dermis. After filtration through a 40-mm cell strainer and thorough washing with PBS, the isolated hHF-MSCs were seeded into a 100-mm cell culture dish and cultured in Dulbecco's modified eagle medium (DMEM), supplemented with 10% fetal bovine serum (FBS) and 10 ng/mL basic fibroblast growth factor (bFGF) in an incubator with 5% CO2 at 37°C. Once cells reached approximately 90% confluence, subculturing was performed using 0.1% trypsin (containing 0.02 mol/L ethylenediamine tetraacetate [EDTA]).
Identification and multidifferentiation assessment
Cells were washed and incubated with the appropriate dilution of control/specific antibodies. Following 45 min of incubation, the cells were washed for 30 min and analyzed through flow cytometry (Beckman™, Brea, CA, USA) following a second washing step.
Osteogenic differentiation assays
hHF-MSCs were incubated in an osteogenic medium for 21 days, and dexamethasone (1 × 10−7 mol/L), ascorbic acid (50 mg/L), and β-glycerolphosphate (10 mol/L) were added to the medium. The osteogenic medium was removed and cells were rinsed with PBS, followed by staining for 2 min by alizarin red. After washing with a differentiation solution for 15 s, the cells were rinsed with PBS and observed under the microscope.
Adipogenic differentiation assays
hHF-MSCs were incubated in an adipogenic medium, which was supplemented with dexamethasone (0.25 μmol/L), isobutylmethylxanthine (0.5 mmol/L), indomethacin (100 μmol/L), and insulin (10 mg/L) for 21 days. Cells were rinsed twice with PBS, fixed with 4% formaldehyde for 30 min, and stained with Oil Red O.
Chondrogenic differentiation assays
Cells were cultured in chondrogenic medium (low-glucose DMEM) at 37°C and 5% CO2 for 21 days. The cells were then fixed with formaldehyde for 1 h after removing from the medium and stained with toluidine blue for 3 h.
Animals and experimental design
Yuhang Kelian Rabbit Co., Ltd. (Hangzhou, China) supplied healthy male New Zealand rabbits. All animals were divided into four groups: control, tendinopathy, hHF-MSCs treatment, and positive cell treatment (umbilical cord MSCs [UCMSC]). Apart from the control group, all groups were injected with 2400 U/2 mL collagenase I administered at 1 cm above the Achilles tendon insertion. To confirm the induction of AT, rabbit gait was observed followed by gross anatomy of the Achilles tendon. Hematoxylin–eosin (HE) staining to determine the number and distribution of prokaryotic cells and arrangement of collagen fibers was performed for pathological observation and the content of hydroxyproline was additionally detected. For tendinopathy treatment, purified hHF-MSCs were injected in and around the Achilles tendon. The entire treatment lasted for 48 days, and the total number of cells collected was 5 × 106 per collection, with three administrations in total at 72-h intervals. The fourth treatment was performed 10 days before sacrifice. The Achilles tendon was anatomically and pathologically observed, and the general health of all animals was noted. Each animal's Achilles tendon was biomechanically assessed to determine its maximum load.
For HE staining, paraffin-embedded tissues were de-waxed three times using xylene, followed by a fourth dewaxing step of 15 min and gradient alcohol dehydration (100%, 95%, and 80% ethanol). The tissues were stained with hematoxylin for 5 min and then with eosin for 3 to 5 min at room temperature. After dehydration using gradient alcohol and xylene transparent application, tissues were sealed with neutral gum.
For immunohistochemical (IHC) staining, paraffin-embedded tissues were dewaxed using xylene, followed by gradient alcohol dehydration. Tissues were incubated with 3% H2O2 at 37°C for 15 min to block and inactivate endogenous peroxidase. Citric acid buffer (0.01 mol/L, pH 6.0) was applied to repair tissue antigens for 10 min. Tissues were incubated with primary antibody at 4°C overnight and then transferred to room temperature for 30 min to reach equilibrium before being incubated with secondary antibodies at 37°C for 30 min. Tissues were stained with 3,3′-diaminobenzidine after rinsing with PBS, followed by water flushing and sealing with neutral gum after counterstaining with hematoxylin.
A tensile testing machine (Instron 5900®; Instron Corporation™, MA, USA) was used to measure biomechanical properties. To prevent drying, samples (n = 10 per group) were immersed in PBS. With three technical replicate readings per group, the rupture point of the Achilles tendon was monitored and recorded as the maximum tensile force.
Quantitative real-time reverse-transcription polymerase chain reaction (qPCR)
Tenascin-C (TNC) and matrix metalloproteinase (MMP)-9 transcriptomic expression from tissue samples were detected by qPCR, accordingly. Briefly, total RNA was isolated from tendon tissue samples using total RNA Rapid Extraction Kit® (Generay™, Shanghai, China) and measured through a Nanodrop® platform. cDNA synthesis was performed using HiScript II Q RT SuperMix® for qPCR (Vazyme™, Nanjing, China). The cDNA was then subjected to reverse-transcription polymerase chain reaction using ChamQ® SYBR Color qPCR Master Mix kit (Vazyme™). The forward and reverse primers for TNC were 5′-AATTCTGACCACCCCCAGGA-3′ and 5′-ACTGTGGTTCTGGCTCTGTG-3′. The forward and reverse primers for MMP-9 were 5′-GCCCCAGCGAAAGACTCTAC-3′ and 5′-TTGTCCTTGTCGTAGCTGGC-3′. The forward and reverse primers for glyceraldehyde 3-phosphate dehydrogenase (GAPDH) were 5′-TGCCGCCTGGAGAAAGC-3′ and 5′-CGACCTGGTCCTCGGTGTAG-3′. Quantitative PCR was conducted using a real-time PCR system. The relative transcriptomic expression value of all investigated genes was normalized against GAPDH and calculated using 2−ΔΔCt method.
Enzyme-linked immunosorbent assay
Following tissue homogenization, total proteomic content for all samples was extracted. TNC/MMP-9 transcriptomic levels within tendon tissues were detected using Rabbit TNC ELISA Kit® (Jianglaibio™, Shanghai, China) and Rabbit MMP-9 ELISA Kit (Jianglaibio™) as per the manufacturer's instructions. This measurement was conducted using a microplate reader (SpectraMax Plus 384®, Molecular Devices™, California, USA).
All data were analyzed using SPSS 13.0 (SPSS; Chicago, IL, USA), and the quantitative values were reported as mean ± standard deviation (SD) and analyzed using the t test, one-way analysis of variance, or one-way multivariate analysis of variance with repeated measurements as appropriate. Differences with a P value of <0.05 were considered statistically significant.
Characterization of hHF-MSCs
Flow cytometry revealed that hHF-MSCs were positive for the mesenchymal stem-cell markers CD49, CD105, CD90, and CD73 and negative for HLA-DR, CD34, CD14, CD45, and CD117 [Figure 1].
Lipid droplets stained with oil red O were red in color under the microscope and the lipid droplets could be observed in hHF-MSCs after the induction of adipogenic differentiation [Figure 2]. Osteogenic differentiation revealed that calcium nodules were stained dark red. Chondrogenic differentiation results revealed that acid mucopolysaccharide represents cartilage differentiation that was stained with toluidine blue. Differentiation analysis revealed that cells can differentiate into multiple types, implying that our current hHF-MSCs originated from MSCs.
Successful establishment of the AT animal model
After AT induction, the gait of the rabbit changed significantly. The Achilles tendon of the control group was glossy, healthy, white in color, whereas that in the AT group was red, swollen, and dull [Figure 3A and 3B].
The number and distribution of prokaryotic cells and the arrangement of collagen fibers were observed by HE staining. Compared with the control group, collagen fibers in the AT group showed a loose and disordered arrangement and uneven distribution [Figure 3C]. Furthermore, the nucleus of prokaryotic cells was blue or purplish blue and the cytoplasm was pink in color. Quantification of prokaryotic cells in Achilles tendon showed fewer prokaryotic cells in the AT group than in the control (cross section, t = 4.670, P = 0.01; longitudinal section, t = 4.578, P = 0.01, Figure 3D).
hHF-MSCs promotes repair of AT
There was little difference between the groups in terms of body weight changes (F = 0.540, P = 0.820, Figure 4A). Gait changes were observed in AT group, and anatomical observation revealed that Achilles tendon was red, swollen and dull, but in hHF-MSCs group, Achilles tendon was healthy, white and shiny, which was better than positive cell group (UCMSC group) [Figure 4B]. The hydroxyproline content in the AT group decreased significantly compared with the control group, indicating that the AT animal model was successfully established. Hydroxyproline levels were higher in the hHF-MSCs and UCMSC groups than in the AT group, and the difference was greater in the hHF-MSCs treatment group (F = 549.263, P < 0.001; AT vs. control, P < 0.001, hHF-MSCs or UCMSC vs. AT, P < 0.001; Figure 4C). Biomechanical analysis revealed that the maximum load of the Achilles tendon in AT group was significantly lower than that of the control (P < 0.05). The maximum load of the hHF-MSCs and UCMSC groups were upregulated compared with that of the AT group (F = 6.11, P = 0.018; UCMSC vs. AT, P = 0.047, hHF-MSCs vs. AT, P = 0.003, Figure 4D, P < 0.05).
Effect of hHF-MSCs on AT by pathological analysis in a rabbit model
The arrangement of collagen fibers in the AT group was found to be disordered on pathological examination, with a large area of inflammatory cells infiltration, which destroyed the structure of collagen fibers. The number of prokaryotic cells decreased significantly, and inflammatory cell aggregation around blood vessels was obvious, but prokaryotic cells increased in the hHF-MSCs group (cross section, F = 168.46, P < 0.001; longitudinal section, F = 121.93, P < 0.001; Figures 5 and 6). Compared with AT group, the collagen fibers in hHF-MSCs group were arranged in order with lesser inflammatory cells infiltration, and the number of prokaryotic cells was normal. Additionally, there was no obvious fiber necrosis and aggregation of inflammatory cells around the blood vessels. This therapeutic effect was better than that in the UCMSC group.
Collagen I and III were stained brown or yellowish brown by IHC staining [Figure 7]. Collagen I and III expression was lower in the AT group than in the control group and higher in the hHF-MSCs group than in the AT group. Furthermore, the average optical density (AOD) of IHC staining was also measured using image-Pro Plus, and the results were expressed as mean ± SD. The AOD also revealed that hHF-MSCs could promote the expression of collagen I (F = 17.97, P = 0.001, hHF-MSCs vs. AT, P = 0.001) and collagen III (F = 7.44, P = 0.01, hHF-MSCs vs. AT, P = 0.007, Figure 8).
Molecular mechanism underlying hHF-MSCs action on tendinopathy
To investigate the possible molecular mechanism of the therapeutic effect of hHF-MSCs on tendinopathy, we examined TNC and MMP-9 mRNA and protein levels in serum and tissue [Figure 9]. TNC mRNA and protein levels were found to be significantly upregulated in the hHF-MSCs group compared with the AT group (for mRNA level, F = 844.97, P < 0.001, hHF-MSCs vs. AT, P < 0.001; for protein level, F = 49.03, P < 0.001, hHF-MSCs vs. AT, P < 0.001, Figure 9), whereas MMP-9 mRNA and protein levels were significantly downregulated in the hHF-MSCs group compared with tendinopathy group (for mRNA level, F = 1174.69, P < 0.001, hHF-MSCs vs. AT, P < 0.001; for protein level, F = 262.87, P < 0.001; hHF-MSCs vs. AT, P < 0.001, Figure 9).
The management of tendinopathy remains a major challenge. Consequently, no specific treatment can be recommended in clinical implications. Platelet-rich plasma and adipose-derived stromal vascular fraction cell-based therapy have been reported to be used in regenerative medicine.[22–24] Their application, however, was limited because of the inconvenient sample collection. Recent research has revealed that the use of stem cells is an intriguing strategy for tendon repair and regeneration, with promising results in terms of effects and safety. Currently, two types of MSCs are commonly used: bone marrow-derived cells and adipose tissue-derived cells. Because of their strong anti-inflammatory and immune-modulatory properties, exosome secreted by adipose-derived MSCs can be used to treat neurodegenerative conditions as well as Corona Virus Disease 2019 (COVID-19).[28,29] These results suggested that the MSC can represent an effective, autologous, and safe therapy for regeneration. However, the issues associated with sourcing it and other ethical issues of MSCs limits its clinical application. Recently, hHF-MSCs have drawn the attention of researchers because of their several advantages, including easy accessibility; easy culture without MHC class I expression, thus reducing the chance of transplant rejection; high proliferative capacity; multipotential properties; and the possibility of autologous use without side effects. In our study, we extracted stem cells from hair follicles and characterization using a trilineage-induced differentiation assessment showed that our present hHF-MSCs were originated from MSCs.
The ability of the hHF-MSCs to promote AT repair in vivo was then tested using a rabbit AT model. Anatomical examination revealed that the Achilles tendon in the hHF-MSCs group was healthy, white, and glossy and overall better than that in the AT group. The hydroxyproline levels increased after hHF-MSCs treatment, implying that hHF-MSCs can reverse tendinopathy-induced hydroxyproline downregulation. Previous research has shown that bone marrow- and adipose tissue-derived stem cells can help with tendon healing.[30,31] This is consistent with our findings that stem cells from hair follicles are more promising for AT treatment. From a biomechanical point of view, a healed tendon is not as efficient as an uninjured one. Our present study also indicated that hHF-MSC treatment can increase the maximum load of the Achilles tendon. Although this load did not reach normal levels, it has greatly improved compared with that of the AT group. This indicated that hHF-MSCs can promote repair of AT.
Tendinopathy was also reported to show a disordered arrangement of collagen fibers. In patients with classical symptoms of AT, changes in collagen fibers structure were evident, with a loss of the normal parallel bundles. Tendon repair after injury mainly involves five stages: inflammation, cell proliferation, cell migration, remodeling, and spatial organization of type I collagen. Pathological analysis in our study revealed the collagen fibers in the HFSC group were arranged in order with few inflammatory cells infiltration, and the number of prokaryotic cells was normal. Furthermore, there was no evidence of fiber necrosis or inflammatory cell aggregation around blood vessels. However, compared with the tendinopathy group, the expression of collagen I and III was increased in the hHF-MSCs group. The findings show that hHF-MSCs are easier to differentiate in tenogenic cells and express more tenogenic genes and collagen I and III, thereby making hHF-MSCs more promising for tendon healing than any other MSCs. Previous studies reported that the expression of genes encoding tendon-associated molecules, including TNC, was upregulated during tendon repair,[36,37] suggesting that TNC plays an important role in tendon repair. hHF-MSCs treatment in our study also significantly induced TNC upregulation, suggesting that tendinopathy treatment by hHF-MSCs promotes the regeneration of collagen fiber, which may be due to the upregulation of tenogenesis genes, such as TNC. Gelatinases, including MMP-9, are a member of the MMP superfamily with collagenolytic activity. A study on rat flexor tendon healing found that MMP-9 was involved in collagen I degradation during the early stages of healing. Our study determined that tendinopathy model induction could increase MMP-9 levels in the Achilles tendon. Moreover, hHF-MSCs can promote AT repair by upregulating TNC and downregulating MMP-9.
In conclusion, we isolated and cultivated hHF-MSCs and verified them through trilineage-induced differentiation assessment. Treatment with hHF-MSCs promoted the repair of AT by upregulation of collagen I and III. Further analysis revealed that hHF-MSCs treatment in tendinopathy promoted the regeneration of collagen fiber that may be because of the upregulation of TNC and downregulation of MMP-9; thus indicating that hHF-MSCs are more promising for AT.
We thank Liwen Bianji (Edanz) (www.liwenbianji.cn) for editing the English text of a draft of this manuscript.
This work was supported by Medical Science and Technology Project of Zhejiang Province (No. 2022RC102), Shanghai “Rising Stars of Medical Talents” Youth Development Program, Youth High-level Talent Special Support Plan of Zhejiang Province, and Innovation High-level Talent Special Support Plan from Health Commission of Zhejiang Province.
Conflicts of interest
1. Macedo CSG, Tadiello FF, Medeiros LT, Antonelo MC, Alves MAF, Mendonca LD. Physical therapy service delivered in the polyclinic during the Rio 2016 paralympic games. Phys Ther Sport
2019; 36:6267. doi: 10.1016/j.ptsp.2019.01.003.
2. Ilaltdinov AW, Gong Y, Leong DJ, Gruson KI, Zheng D, Fung DT, et al. Advances in the development of gene therapy, noncoding RNA, and exosome-based treatments for tendinopathy
. Ann N Y Acad Sci
2021; 1490:312. doi: 10.1111/nyas.14382.
3. Cook JL, Purdam CR. Is tendon pathology a continuum? A pathology model to explain the clinical presentation of load-induced tendinopathy
. Br J Sports Med
2009; 43:409416. doi: 10.1136/bjsm.2008.051193.
4. Maffulli N, Khan KM, Puddu G. Overuse tendon conditions: time to change a confusing terminology. Arthroscopy
1998; 14:840843. doi: 10.1016/s0749-8063(98)70021-0.
5. Challoumas D, Kirwan PD, Borysov D, Clifford C, McLean M, Millar NL. Topical glyceryl trinitrate for the treatment of tendinopathies: a systematic review. Br J Sports Med
2019; 53:251262. doi: 10.1136/bjsports-2018-099552.
6. Murphy MC, Travers MJ, Chivers P, Debenham JR, Docking SI, Rio EK, et al. Efficacy of heavy eccentric calf training for treating mid-portion Achilles tendinopathy
: a systematic review and meta-analysis. Br J Sports Med
2019; 53:10701077. doi: 10.1136/bjsports-2018-099934.
7. Lu H, Yang H, Shen H, Ye G, Lin XJ. The clinical effect of tendon repair for tendon spontaneous rupture after corticosteroid injection in hands: a retrospective observational study. Medicine
2016; 95:e5145:1–4 doi: 10.1097/MD.0000000000005145.
8. Pittenger MF, Mackay AM, Beck SC, Jaiswal RK, Douglas R, Mosca JD, et al. Multilineage potential of adult human mesenchymal stem cells. Science
1999; 284:143147. doi: 10.1126/science.284.5411.143.
9. Gentile P. New strategies in plastic surgery: autologous adipose-derived mesenchymal stem cells contained in fat grafting improves symptomatic scars. Front Biosci
2021; 26:255257. doi: 10.52586/4940.
10. Gentile P, Garcovich S. Concise review: adipose-derived stem cells (ASCs) and adipocyte-secreted exosomal microRNA (A-SE-miR) modulate cancer growth and promote wound repair. J Clin Med
2019; 8:855:1–13 doi: 10.3390/jcm8060855.
11. Gentile P, Garcovich S. Systematic review: adipose-derived mesenchymal stem cells, platelet-rich plasma and biomaterials as new regenerative strategies in chronic skin wounds and soft tissue defects. Int J Mol Sci
2021; 22:1538:1–14 doi: 10.3390/ijms22041538.
12. Gentile P, Sterodimas A, Calabrese C, Garcovich S. Systematic review: advances of fat tissue engineering as bioactive scaffold, bioactive material, and source for adipose-derived mesenchymal stem cells in wound and scar treatment. Stem Cell Res Ther
2021; 12:318:1–16 doi: 10.1186/s13287-021-02397-4.
13. Bianco ST, Moser HL, Galatz LM, Huang AH. Biologics and stem cell-based therapies for rotator cuff repair. Ann N Y Acad Sci
2019; 1442:3547. doi: 10.1111/nyas.13918.
14. Gentile P, Garcovich S. Advances in regenerative stem cell therapy in androgenic alopecia and hair loss: Wnt pathway, growth-factor, and mesenchymal stem cell signaling impact analysis on cell growth and hair follicle development. Cells
2019; 8:466:1–21 doi: 10.3390/cells8050466.
15. Gentile P, Scioli MG, Bielli A, Orlandi A, Cervelli V. Stem cells from human hair follicles: First mechanical isolation for immediate autologous clinical use in androgenetic alopecia and hair loss. Stem Cell Investig
2017; 4:58:1–10 doi: 10.21037/sci.2017.06.04.
16. Kiani MT, Higgins CA, Almquist BD. The hair follicle: an underutilized source of cells and materials for regenerative medicine. ACS Biomater Sci Eng
2018; 4:11931207. doi: 10.1021/acsbiomaterials.7b00072.
17. Liu JY, Peng HF, Gopinath S, Tian J, Andreadis ST. Derivation of functional smooth muscle cells from multipotent human hair follicle mesenchymal stem cells. Tissue Eng Part A
2010; 16:25532564. doi: 10.1089/ten.TEA.2009.0833.
18. Yu H, Kumar SM, Kossenkov AV, Showe L, Xu X. Stem cells with neural crest characteristics derived from the bulge region of cultured human hair follicles. J Invest Dermatol
2010; 130:12271236. doi: 10.1038/jid.2009.322.
19. Zeisel A, Hochgerner H, Lonnerberg P, Johnsson A, Memic F, van der Zwan J, et al. Molecular architecture of the mouse nervous system. Cell
2018; 174:9991014. e22. doi: 10.1016/j.cell.2018.06.021.
20. Yari A, Heidari F, Veijouye SJ, Nobakht M. Hair follicle stem cells promote cutaneous wound healing through the SDF-1alpha/CXCR4 axis: an animal model. J Wound Care
2020; 29:526536. doi: 10.12968/jowc.2020.29.9.526.
21. Wang B, Liu XM, Liu ZN, Wang Y, Han X, Lian AB, et al. Human hair follicle-derived mesenchymal stem cells
: Isolation, expansion, and differentiation. World J Stem Cells
2020; 12:462470. doi: 10.4252/wjsc.v12.i6.462.
22. Gentile P, Alves R, Cole JP, Andjelkov K, Van Helmelryck T, Fernandez J, et al. AIRMESS - Academy of International Regenerative Medicine & Surgery Societies: recommendations in the use of platelet-rich plasma (PRP), autologous stem cell-based therapy (ASC-BT) in androgenetic alopecia and wound healing. Expert Opin Biol Ther
2021; 21:14431449. doi: 10.1080/14712598.2021.1908995.
23. Gentile P, Calabrese C, De Angelis B, Dionisi L, Pizzicannella J, Kothari A, et al. Impact of the different preparation methods to obtain autologous non-activated platelet-rich plasma (A-PRP) and activated platelet-rich plasma (AA-PRP) in plastic surgery: wound healing and hair regrowth evaluation. Int J Mol Sci
2020; 21:431:1–9 doi: 10.3390/ijms21020431.
24. Gentile P, Casella D, Palma E, Calabrese C. Engineered fat graft enhanced with adipose-derived stromal vascular fraction cells for regenerative medicine: clinical, histological and instrumental evaluation in breast reconstruction. J Clin Med
2019; 8:504:1–21 doi: 10.3390/jcm8040504.
25. Behfar M, Sarrafzadeh-Rezaei F, Hobbenaghi R, Delirezh N, Dalir-Naghadeh B. Enhanced mechanical properties of rabbit flexor tendons in response to intratendinous injection of adipose derived stromal vascular fraction. Curr Stem Cell Res Ther
2012; 7:173178. doi: 10.2174/157488812799859874.
26. Costa-Almeida R, Calejo I, Reis RL, Gomes ME. Crosstalk between adipose stem cells and tendon cells reveals a temporal regulation of tenogenesis by matrix deposition and remodeling. J Cell Physiol
2018; 233:53835395. doi: 10.1002/jcp.26363.
27. Gentile P, Piccinno MS, Calabrese C. Characteristics and potentiality of human adipose-derived stem cells (hASCs) obtained from enzymatic digestion of fat graft. Cells
2019; 8:282:1–20 doi: 10.3390/cells8030282.
28. Gentile P. SARS-CoV-2: the “uncensored” truth about its origin and adipose-derived mesenchymal stem cells as new potential immune-modulatory weapon. Aging Dis
2021; 12:330344. doi: 10.14336/AD.2021.0121.
29. Gentile P, Sterodimas A, Pizzicannella J, Calabrese C, Garcovich S. Research progress on mesenchymal stem cells (MSCs), adipose-derived mesenchymal stem cells (AD-MSCs), drugs, and vaccines in inhibiting COVID-19 disease. Aging Dis
2020; 11:11911201. doi: 10.14336/AD.2020.0711.
30. Goncalves AI, Gershovich PM, Rodrigues MT, Reis RL, Gomes ME. Human adipose tissue-derived tenomodulin positive subpopulation of stem cells: a promising source of tendon progenitor cells. J Tissue Eng Regen Med
2018; 12:762774. doi: 10.1002/term.2495.
31. Tan SL, Ahmad RE, Ahmad TS, Merican AM, Abbas AA, Ng WM, et al. Effect of growth differentiation factor 5 on the proliferation and tenogenic differentiation potential of human mesenchymal stem cells in vitro. Cells Tissues Organs
2012; 196:325338. doi: 10.1159/000335693.
32. Guevara-Alvarez A, Schmitt A, Russell RP, Imhoff AB, Buchmann S. Growth factor delivery vehicles for tendon injuries: mesenchymal stem cells and platelet rich plasma. Muscles Ligaments Tendons J
2014; 4:378385. doi: 10.32098/mltj.03.2014.18.
33. Benazzo F, Stennardo G, Mosconi M, Zanon G, Maffulli N. Muscle transplant in the rabbit's Achilles tendon. Med Sci Sports Exerc
2001; 33:696701. doi: 10.1097/00005768-200105000-00003.
34. Astrom M, Gentz CF, Nilsson P, Rausing A, Sjoberg S, Westlin N. Imaging in chronic Achilles tendinopathy
: a comparison of ultrasonography, magnetic resonance imaging and surgical findings in 27 histologically verified cases. Skeletal Radiol
1996; 25:615620. doi: 10.1007/s002560050146.
35. Voleti PB, Buckley MR, Soslowsky LJ. Tendon healing: repair and regeneration. Annu Rev Biomed Eng
2012; 14:4771. doi: 10.1146/annurev-bioeng-071811-150122.
36. Guerquin MJ, Charvet B, Nourissat G, Havis E, Ronsin O, Bonnin MA, et al. Transcription factor EGR1 directs tendon differentiation and promotes tendon repair. J Clin Invest
2013; 123:35643576. doi: 10.1172/JCI67521.
37. Scott A, Sampaio A, Abraham T, Duronio C, Underhill TM. Scleraxis expression is coordinately regulated in a murine model of patellar tendon injury. J Orthop Res
2011; 29:289296. doi: 10.1002/jor.21220.
38. Vargova V, Pytliak M, Mechirova V. Matrix metalloproteinases. Exp Suppl
2012; 103:133. doi: 10.1007/978-3-0348-0364-9_1.
39. Oshiro W, Lou J, Xing X, Tu Y, Manske PR. Flexor tendon healing in the rat: a histologic and gene expression study. J Hand Surg Am
2003; 28:814823. doi: 10.1016/s0363-5023(03)00366-6.