CTGF Induces Tenogenic Differentiation and Proliferation of Adipose-Derived Stromal Cells

Abstract

Intrasynovial tendons are paucicellular and hypovascular, resulting in a poor response to injury. Surgical repair of ruptured or lacerated tendons often lead to complications such as adhesions, repair site gapping, and repair site rupture. Adipose-derived stem cells (ASCs) have shown promise for enhancing tendon repair, as they have the capacity to differentiate into tendon fibroblasts and augment the healing response. Furthermore, connective tissue growth factor (CTGF) has been shown to promote tendon regeneration via the stimulation of endogenous tendon stem cells. Here, we evaluated the potential of CTGF to promote tenogenic differentiation of ASCs in vitro. Gene and protein expression, cell proliferation, and FAK and ERK1/2 signaling were assessed. CTGF increased tenogenic genes in mouse ASCs in a dose- and time-dependent manner. Western blot and immunostaining analyses demonstrated increases in tenogenic protein expression in CTGF-treated ASCs at all timepoints studied. CTGF increased ASC proliferation in a dose-dependent manner. CTGF induced phosphorylation of ERK1/2 within 5 min and FAK within 15 min; both signals persisted for 120 min. Blocking FAK and ERK1/2 pathways by selective inhibitors SCH772984 and PF573228, respectively, attenuated the CTGF-induced tenogenic differentiation and proliferation of ASCs. These results suggest that CTGF induces tenogenic differentiation of ASCs via the FAK and ERK1/2 pathway. Statement of clinical significance: Although prior research has led to advances in tendon operative techniques and rehabilitation methods, clinical outcomes after tendon repair remain variable, with high rates of repair site gapping or rupture.

Acute tendon injury and chronic tendinopathy affect a large portion of the aging population, leading to pain, disability, and large societal costs. For example, one-third of all acute injuries in workers are to the upper extremity, many of which are open wounds requiring extensive tendon surgery, including flexor tendon mid-substance repair.^1–3 These upper extremity injuries result in over 3 million restricted activity days and over 1.5 million days lost from work per year. Despite advances in operative techniques and rehabilitation methods, the outcomes after tendon repair are highly variable, resulting in a substantial clinical burden. Specifically, a high rate of gapping or repair site failure has been noted following the surgical repair of tendons of the hand, of the ankle, and of the rotator cuff.^4–6

Tendon healing after surgical repair progresses through phases of inflammation, proliferation, extracellular matrix (ECM) formation, and ECM remodeling.^7,8 The repair process is driven initially by high levels of inflammation and a propensity towards the formation of disorganized scar tissue. Due to the paucicellular and relatively avascular nature of tendon, the proliferative and remodeling phases of tendon healing are typically slow, resulting in only marginally successful recovery of tendon strength. Accordingly, therapeutic approaches have focused on promoting tendon regeneration through the delivery of mesenchymal stromal cells (MSCs), the expansion of endogenous tendon stem cells, and/or the delivery of tenogenic growth factors.^9,10 MSCs are readily available from autologous sources, including bone marrow and adipose tissue. Adipose-derived MSCs (ASCs) are particularly attractive because they are easier to obtain and more abundant than bone marrow-derived MSCs.¹¹ These cells have the capacity to differentiate into multiple lineages, including tendon fibroblasts, osteoblasts, chondrocytes, and adipocytes.¹² A number of strategies have been used to promote tenogenesis of MSCs, including growth factors and mechanical stimulation, but the molecular cues driving such process are unclear.^13–16

Previously, a number of growth factors have been shown to promote tenogenesis of cultured MSCs and endogeneous tendon stem cells, including connective tissue growth factor (CTGF), transforming growth factor beta (TGF-β), growth and differentiation factor 7 (GDF 7, a.k.a BMP-12), and GDF5.^17–20 In vitro and in vivo studies have recently demonstrated that CTGF, a member of the CCN family of proteins, is particularly attractive for tendon repair as it is capable of promoting tendon regeneration via the stimulation of endogenous tendon stem/progenitor cells, a process likely mediated by FAK/ERK1/2 signaling.^21–23

The goal of the current study was to evaluate the role of CTGF for tenogenic differentiation and proliferation of ASCs. To determine the tenogenic potential of CTGF, ASCs were isolated from mouse subcuticular fat tissue and cultured in vitro. Tenogenesis was evaluated using gene and protein expression and the mechanism of CTGF-induced tenogenesis was explored. Results showed that CTGF stimulated expression of tenogenic differentiation markers and proliferation in ASCs. Furthermore, FAK/ERK1/2 signaling was necessary for CTGF-induced tenogenesis of ASCs. These results suggested that CTGF plays a role in the tenogenic differentiation of ASCs, supporting the use of CTGF coupled with ASCs for improving tendon healing and providing a molecular and cellular basis for future tendon tissue engineering approaches.

MATERIALS AND METHODS

Cell Isolation and Culture

ASCs were isolated from subcutaneous fat of mice postmortem used for other studies approved by the Columbia University IACUC. The fat tissue (1 g per mouse) was minced into a fine slurry, digested with 0.5% Animal Free Collagenase/Dispase Blend II (MilliporeSigma,Germany) in dPBS at 37 °C for up to 1 h, and then centrifuged at 250 g for 5 min. The pellets were re-suspended in minimum essential medium alpha (alpha-MEM; Gibco, Life Technology, Carlsbad, CA) containing 10% heat inactivated fetal bovine serum (HI-FBS; Gibco, Life technology) and 1% antibiotic (1 x antibiotic-antimycotic, including 10 units/l penicillin G sodium, 10 mg/ml streptomycin sulfate) (Invitrogen, Carlsbad, CA). At 80%–90% confluence, cells were trypsinized, centrifuged, resuspended in growth medium as passage 1 (P1) cells, and incubated in 5% CO₂ at 37 °C, with fresh medium changes every 2–3 days. The plastic adherent ASCs were then maintained in alpha-MEM culture medium and passaged when they were 80–90% confluent. All of the cells were used for experiments at passage 2 unless otherwise specified. Prior studies demonstrated that this cell isolation approach produces mesenchymal stromal cells expressing CD29, CD44, and CD90 cell surface markers and that these cells have pluripotent (adipogenic, osteogenic, tenogenic, and chondrogenic) differentiation potential.^15,24,25

To examine the dose-response curve for the effect of CTGF on differentiation of ASCs, a total of 100,000 cells/well of ASCs were plated in 6 wells in differentiation medium (alpha-MEM supplemented with 2% FBS). On the following day, the cells were treated with 0, 1, 10, or 100 ng/ml of CTGF (BioVendor, Asheville, NC) in the same medium for up to 14 days. In cases where the ERK1/2 inhibitor SCH772984 (0–10,000 ng/ml) (Selleckchem, Houston, TX) or FAK inhibitor PF573228 (0–10,000 ng/ml) (Sigma, St. Louis, MO) was applied, the cells were pretreated with either of the drugs for 30 min prior to the addition of growth factors. Culture medium was changed every 2 to 3 days. Each experiment was repeated a minimum of three times.

Gene Expression: RNA Isolation and Real-Time RT-PCR

ASCs were isolated and grown in 6-well plates as previously described.¹⁵ At the indicated timepoints, cells were harvested for analysis of mRNA expression by real-time RT-PCR. RNA was extracted using an RNeasy Mini Spin Column (Qiagen Sciences, Hilden, Germany) according to the manufacturer’s instructions. Potential genomic DNA contamination was eliminated by treating RNA samples with DNase I (Qiagen, Hilden, Germany) during purification.

Real-time RT-PCR was performed in two steps. In the first step, total RNA (500 ng) was reversely transcribed into first-strand cDNA in a 20 μl reaction with random primers using the SuperScript VILO cDNA Synthesis Kit (Life Technologies, Carlsbad, CA) according to the manufacturer’s instructions. PCR of 1 μl of diluted cDNA from each sample was carried out in 20 μl reactions containing Platinum SYBR Green qPCR SuperMix-UDG and appropriate primers in the ABI StepOne Plus System from Applied Biosystems (Life technologies). The PCR thermal cycle was set as the following: 10 min at 95 °C for one cycle; 15 s at 95 °C and 40 s at 60 °C for 40 cycles. A melt curve analysis was performed at the end of each SYBR Green PCR. The efficiencies of all the primers were all over 90%. The relative quantity of target gene expression was analyzed using the comparative CT (2^−∆∆CT) method GAPDH was used as endogenous reference housekeeping gene. All RT-PCR primers used in this study were custom-designed and purchased from Integrated DNA Technologies (scleraxis [SCX] forward primer, 5′-tccgacgagaaaccctgc-3′; reverse primer, 5′-gcagcgtctcaatcttggag-3′; tenomodulin [TNMD] forward primer, 5′-tggtatcctggccttaactct-3′; reverse primer, 5′-ttctgttctggttatgggatcaa-3′; collagens type I(COL1A1) forward primer, 5′-caaagacgggagggcgag-3′; reverse primer, 5′-ctgtccagggatgccatctc-3′; collagens type III(COL3A1) forward primer, 5′-atgaggagccactagactgc-3′; reverse primer, 5′-ggtcaccatttctcccagga-3′; aggrecan [ACAN] forward primer, 5′-tccagaaccttcgctccaat-3′; reverse primer, 5′-gctgtgctcgatcaaagtcc-3′; runt-related transcription factor 2 [RUNX2] forward primer, 5′-agatgggactgtggttaccg-3′; reverse primer, 5′-tagctctgtggtaagtggcc-3′; GAPDH forward primer, 5′-atcaccatcttccaggagcg-3′).

Protein Expression

Western Blot Assay

To examine the expression of tenogenic markers, ASCs were cultured and treated with CTGF as described above. To study ERK1/2 and FAK phosphorylation, ASCs were cultured in 6-well plates at a density of 1 × 10⁵ cells/well and starved in growth medium overnight. On the following day, the cells were treated with 100 ng/ml CTGF for the indicated periods in duplicate. The dose of CTGF was chosen based on the strongest tenogenic dose from the gene expression study described above. In cases when the inhibitor SCH772984 (Selleckchem) or PF573228 (Sigma) was applied, the cells were pre-treated with either of the inhibitors for 30 min prior to the addition of CTGF.

Cellular protein was extracted in RIPA Lysis Buffer (Thermo Fisher Scientific, Waltham, MA) with Protease/Phosphatase Inhibitor Cocktail (Cell Signaling Technology, Danvers, MA). Proteins were separated by SDS–PAGE and transferred to nitrocellulose membranes (Bio-Rad, Hercules, CA) using a semi-dry transfer apparatus. Blots were blocked for 1 h in 5% non-fat dry milk dissolved in Tris-buffered saline containing 0.2% Tween-20 (TBST), then incubated in primary antibodies overnight at 4 ˚C. The following antibodies were used: Anti-Scx (1:500) and anti-Tnmd (1:50), purchased from Abcam (Cambridge, UK); antiphosphorylated FAK (sc-3283s), antiphosphorylated ERK1/2 (sc-292838), and anti-actin (sc-4970s), purchased from Cell Signaling Technology. Following primary antibody incubation, blots were washed three times in TBST and incubated in goat anti-rabbit IgG (H&L)-HRP conjugated secondary anitbody (1:2000) for 1 h at room temperature. Signals were visualized using an ECL chemiluminescent detection kit (Thermo Scientific, Waltham, MA).

Densitometric measurement of bands was performed on scanned immunoblot images using Image J software (Bethesda, MD). After subtracting the background, the mean grayscale value and integrated density were determined and used to calculate band volume. The freehand selection tool was used to identify the bands. Bands density was normalized to sample loading controls.

Immunofluorescence Staining

For tenogenic protein expression, cultured ASCs were fixed with 4% paraformaldehyde in PBS. After three washes with PBS, the cells were permeabilized with 0.5% Triton X-100 in PBS and blocked with 5% normal donkey serum (Jackson ImmunoResearch, West Grove, PA) in PBS containing 0.1% Tween-20 (PBST). Cells were then incubated with rabbit anti-Scx (1:100) or anti-Tnmd antibodies (1:200) (Santa Cruz Biotechnology, Santa Cruz, CA) at 4 °C overnight. After three washes with PBS, the cells were incubated with Cy3-conjugated Donkey AntiRabbit IgG (H þ L) (1:200) (Jackson ImmunoResearch) for 1 h at room temperature and then counterstained with DAPI (Sigma). All images were acquired using an inverted fluorescence microscope (Leica DMi8).

Cell Proliferation

The CCK-8 assay was used to measure cell viability in cell proliferation under starved conditions. The assay is based on the conversion of a water-soluble tetrazolium salt, 2-(2-methoxy-4-nitrophenyl)-3-(4-nitrophenyl)-5-(2,4-disulfophenyl)-2H-tetrazolium, monosodium salt (WST-8), to a water-soluble formazan dye upon reduction by dehydrogenases in the presence of an electron carrier. ASCs (2 × 10³ cells/ml) were grown in 96-well plates for up to 7 days and treated with CTGF over a range of concentrations (0, 10, 100 ng/ml). After 24 h, the ASCs were washed, and the extent of cell growth was assessed each day using a CCK-8 assay (Dojindo, Japan). CCK-8 solution (10 ml) was added to each well, followed by incubation for 2 h at 37 °C. Absorbance at 450 nm was determined using a multiplate reader (Biotek). For each concentration of CTGF, mean values of the mean absorbance levels from six wells were calculated.

Signaling Pathway Assessment

To explore potential non-canonical signaling processes involved in CTGF-induced tenogenesis, Cignal 45-Pathway Reporter Arrays were used to simultaneously assess 45 different signaling pathways by luciferase assay. Mouse ASCs (40,000 cells/well) were seeded into Cignal Finder 96-well plates (SABiosciences, Hilden, Germany) according to the manufacturer’s protocol. Reporter DNA constructs resident in each plate well were re-suspended with 50 ml Opti-MEM and then mixed with 50 μl diluted transfection reagent. ASCs were suspended in Opti-MEM supplemented with 5% FBS and 0.1 mM NEAA at a density of 6 × 10⁵ cells/ml. The 50 μl cell suspension was added into each plate well and mixed with DNA resident in the plate, and transfection complexes were re-suspended and mixed in wells. The cells were incubated at 5 % CO₂ and 37 °C up to 48 h. After 48 h, the cells were treated with DMSO or CTGF (100 ng/ml) for 6 h in fresh Opti-MEM. Cells were then lysed with 1 × PLB buffer (25 μl/well) by incubation on a shaking platform for 30 min at 25 °C in the dark. The lysates (20 μl) were transferred to wells of opaque 96-well plates for measurement of luciferase expression on a multiplate reader (Biotek).²⁶

Statistics

The effect of CTGF dose and time in culture were assessed using two-way ANOVAs with Fisher’s LSD post-hoc tests when appropriate (StatView 5.0, SAS Institute Inc.). p < 0.05 was considered statistically significant. The inhibitor experiments were assessed using one-way ANOVAs with post-hoc Tukey HSD tests, when appropriate. All data are shown as mean ± standard deviation.

RESULTS

CTGF Induced Tenogenic Differentiation of ASCs

ASCs were isolated from mouse subcutaneous fat and exposed to increasing concentrations of CTGF (1–100 ng/ml) for 3, 7, or 14 d. The effect of CTGF on ASC differentiation was investigated using gene and protein markers: tenogenic transcription factor scleraxis (SCX), tendon-specific marker tenomodulin (TNMD), tendon extracellular matrix collagens type I (COL1A1) and type III (COL3A1), cartilage extracellular matrix aggrecan (ACAN), and osteogenic transcription factor runt-related transcription factor 2 (RUNX2). CTGF dramatically increased expression of SCX and TNMD in a time and dose-dependent manner (SCX: Fig. 1A, p < 0.001 for dose and p = 0.0075 for time; TNMD: Fig. 1B, p = 0.0027 for dose and p < 0.001 for time). Expression of COL1A1 mRNA also increased in a time and dose-related fashion by up to fivefold (Fig. 1C; p < 0.001 for dose and time). COL3A1 levels in CTGF-treated ASCs were maximal at 14 days at the 10 ng/ml dose, but no changes were observed at other doses and timepoints (Fig. 1D). When examining the expression of cartilage matrix genes, CTGF dose-dependently increased ACAN expression (Fig. 1E; p < 0.01). When examining osteogenesis, CTGF time dependently suppressed RUNX2 expression in ASCs by up to 50% (Fig. 1F, p = 0.042). Based on the gene expression results, the most effective dose and treatment time for CTGF was 100 ng/ml for 14 days, respectively.

Figure 1.

(A–F) CTGF induced tenogenic (Scx, TNMD, and COL1A1) but not osteogenic (RUNX2) gene expression in mouse ASCs (mean fold change ± SD; *p < 0.05 for effect of dose; # p < 0.05 for effect of CTGF).

The tenogenic effect of CTGF demonstrated at the gene expression level was corroborated by protein-level assays. Specifically, protein expression was examined for tendon-specific SCX and TNMD using Western Blot analysis and immunocytochemistry at 3, 7, and 14 d for the 100 ng/ml CTGF dose. CTGF treatment induced dose-dependent increases in both SCX and TNMD by ASCs (Fig. 2).

Figure 2.

(A and B) CTGF induced SCX and TNMD protein expression in mouse ASCs, as determined by (A) Western blots and (B) immunocytochemistry (scale bar = 50 μm).

CTGF Induced Proliferation of ASCs

The CCK-8 assay was performed to determine the effect of CTGF on ASC proliferation. ASCs were treated with increasing CTGF concentrations (0, 10, and 100 ng/ml) for 1, 2, 3, 5, or 7 d. Proliferation was similar for the first 3 d of culture, regardless of CTGF treatment. However, CTGF increased ASC proliferation on days 5 and 7 in a dose-dependent manner (Fig. 3, p < 0.01 on both days 5 and 7).

Figure 3.

CTGF promoted proliferation in mouse ASCs (mean fold change ± SD; *p < 0.05 compared to 0 ng/ml CTGF).

CTGF-Induced Tenogenesis and Proliferation of ASCs Was Dependent on the ERK1/2 and FAK Pathway

To identify potential signaling pathways involved in CTGF-induced tenogenic dierentiation and proliferation of ASCs, a 45-pathway reporter array assay was used. Treatment of ASCs with 100 ng/ml of CTGF for 6 h induced upregulation and downregulation of a number of signaling pathways (Fig. 4A). The top five most responsive reporters were FOXO, ERSE, MTF1, TCF/LEF, and SRE. The SRE reporter represents binding related to the ERK1/2 and FAK pathway and is considered a canonical pathway related to cell dierentiation and proliferation and previously described as a critical pathway for CTGF-induced effects.^23,27

Figure 4.

(A) CTGF induced a number of signaling pathways in mouse ASCs, including FAK/ERK1/2 (indicated by an in increase in the marker SRE). (B) Western blot analysis revealed phosphorylation of ERK1/2 and FAK by CTGF in mouse ASCs.

To further explore the roles of ERK1/2 and FAK in CTGF-induced ASC differentiation and proliferation, Western blots were used to examine protein phosphorylation. CTGF induced robust phosphorylation of ERK1/2 within 5 min (Fig. 4B). A strong p-ERK1/2 signal persisted for 120 min following treatment. A p-FAK signal was detected in ASCs treated with CTGF for 15 min and persisted after 120 min. Selective inhibitors for ERK1/2 (SCH772984²⁸) or FAK (PF573228²⁹) were then applied to determine the necessity of this signaling pathway on CTGF-induced tenogenic differentiation of ASCs. The inhibitors were effective in blocking ERK1/2 and FAK phosphorylation (Fig. 5A and B). Inclusion of ERK1/2 inhibitor SCH772984 or FAK inhibitor PF573228 in cultures of CTGF-treated ASCs resulted in significant attenuation of tenogenesis-specific markers SCX, TNMD, and COL1A1 (Fig. 5C–E, p < 0.05 for all three genes). However, no change in COL3A1 gene expression was observed due to either inhibitor (data not shown). Furthermore, blocking FAK or ERK1/2 by selective inhibitor significantly reduced CTGF-induced proliferation of ASCs at days 5 and 7 (Fig. 5F, p < 0.05 at 5 d and 7 d for both ERK1/2 and FAK groups).

Figure 5.

Blocking ERK1/2 and FAK phosphorylation decreased the expression of tenocyte lineage markers in ASCs. (A and B) SCH772984 and PF562271 attenuated ERK1/2 and FAK phosphorylation, respectively, in a dose response manner. (C–E) CTGF enhanced expression of tenogenic markers SCX, TNMD, and COL1A1. Blocking ERK1/2 and FAK suppressed the tenogenic effect of CTGF. (F) Blocking ERK1/2 and FAK suppressed the proliferative effect of CTGF. (Mean fold change ± SD; *p < 0.05, #p < 0.05 compared to CTGF treatment and no inhibitor).

DISCUSSION

The current study explored the role of CTGF in driving tenogenesis and proliferation in mouse ASCs. Based on increased expression of the tendon markers SCX, TNMD, and COL1A1, CTGF induced tenogenic differentiation of ASCs in a dose- and time-dependent manner. The tenogenic effect of CTGF was modulated by FAK and ERK1/2 signaling, demonstrated by the signaling study using selective FAK and ERK1/2 inhibitors. These findings provide new insights into the cellular and molecular mechanisms of tenogenic differentiation and introduce a new basis for cell-based enhanced tendon repair.

In previous work, we examined the effect of the growth factors bFGF, PDGF-BB, and BMP12 on MSC differentiation and tendon healing. Early work focused on bFGF and PDGF-BB, which generally stimulated proliferation and matrix synthesis in fibroblastic cells. However, the growth factors tested neither promoted regeneration of tendon nor produced improvements in biomechanical properties.^17,18,30 CTGF has been shown to induce differentiation of bone marrow MSCs²¹ and tendon stem/progenitor cells (TSPCs)²³ into tendon fibroblasts both in vitro and in vivo. Consistent with the previous results, our findings showed that CTGF induced expression of SCX and TNMD at both protein and mRNA levels in ASCs in vitro.

In addition to the stimulation of tenogenic genes, we found that CTGF significantly increased ACAN expression in ASCs, an observation previously reported by Cordula et al.³¹ This effect may not necessarily result in cartilage formation, however, as proteoglycans are also essential components of tendon. Moreover, Lee et al.²¹ found COL2A1, a key marker for chondrogenesis, was suppressed during co-cultured with CTGF in bone marrow-derived MSCs. In contrast to teno- and chondro-genic marker gene expression, CTGF showed dose- and time-dependent suppression of RUNX2 in ASCs by up to 50%. These findings further support the notion that CTGF promotes tenogenic over osteogenic differentiation of ASCs. Although the current study did not test for adipogenesis in CTGF-treated ASCs, Cignal finder 45-pathway reporter array data showed that the Myc reporter (an early indicator of adipogenesis) was markedly down-regulated (Fig. 5A).^32,33 Notably, SCX and TNMD increased slightly with time in culture, even in the absence of CTGF. Therefore, under the conditions used in the current study, tendon differentiation was dominant over osteogenesis, chondrogenesis, and adipogenesis. In addition, Nanog, a classic pluripotency or stemness marker,³⁴ was also downregulated with CTGF, indicating loss of stemness and progression towards a fully differentiated cell phenotype.³⁵

The molecular mechanisms by which mesenchymal stem cells differentiate into tendon fibroblasts are poorly understood. Previous work demonstrated that BMP12 promotes tenogenesis in ASCs through Smad1/5/8, but not Smad2/3 and p38 signaling.¹⁵ In the current study, FAK/ERK1/2 was demonstrated to be necessary for CTGF-induced tenogenesis and proliferation of ASCs. Suppression of tenogenesis was achieved by blocking either FAK or ERK1/2 phosphorylation, suggesting that FAK and ERK1/2 have key roles in the process. Consistent with these results, a prior study demonstrated that siRNA knockdown of FAK and ERK1/2 attenuated tenogenesis and proliferation in a different study.²³ Furthermore, Liu et al.²² reported that CTGF-induced tenogenesis may involve direct physical interactions with BMP12 via the Smad1/5/8 pathway. In the current study, the Smad reporter in the 45-pathway reporter array was also upregulated, indicating that a BMP12-related pathway may be involved in CTGF-induced tenogenesis and proliferation. Similar to a recent report suggesting activation of AKT-mTOR signaling for tenogenesis of MSCs,³⁶ we noted upregulation of this pathway according to the signaling array (FOXO reporter). The interactions between these pathways for tenogenesis remain unclear, and the potential crosstalk between CTGF, TGF-β (BMP12), and mTOR signaling pathways is a limitation of the present study. Future studies will determine the necessity of these unexplored pathways for CTGF-induced tenogenesis.

CONCLUSIONS

In summary, this study demonstrated CTGF as a tenogenic growth factor, inducing ASCs towards a tendon fibroblast lineage. Mechanistic studies exploring the signaling pathway involved in tenogenesis also furthered our knowledge of how stem cells become tendon fibroblasts. As ASCs can be readily aspirated with a minimally invasive and clinically attractive procedures and differentiated into tendon fibroblasts, the study also presents an appealing and plentiful autologous cell source for tendon repair applications. Subsequent studies will explore the potential for this cell- and growth factor-based approach to improve tendon healing in flexor tendon and rotator cuff animal models.