Abstract
Purpose:
Transforming myofibroblasts into adipocyte-like cells may be a viable option for treating Dupuytren disease. Human Dupuytren myofibroblasts (DMFs) and adipose-derived stem cells (ASCs) co-cultured in the presence of platelet-rich plasma (PRP) reprogrammed into lipid-laden cells. This treatment also reduced fibrosis markers in vivo We aimed to determine if this treatment transformed DMFs into adipocyte-like cells in vivo and to characterize contributing PRP factors to this transformation.
Methods:
DMFs and normal human dermal fibroblasts (NHDFs) were transplanted into the forepaws of rats (Rowett Nude [rnu/rnu]). Two months later, the paws were treated with saline, ASCs+PRP, or Clostridium histolyticum (clinical comparison) once a week for three treatments. The paw tissue was harvested one week after each treatment and subjected to Masson trichrome staining, collagen I and III, α-smooth muscle actin (SMA), and perilipin detection by immunohistochemistry. DMFs were co-cultured with ASCs and PRP or IGF-I or IGF-I depleted PRP. In addition, the IGF-I receptor was inhibited. Oil Red O or BODIPY detected lipid-laden cells.
Results:
Rodent paws implanted with DMFs showed enhanced α-SMA expression, imbalanced collagen III:I ratio, and reduced adipocytes compared to NHDFs. After treatment with ASCs+PRP, DMF paws demonstrated reduced α-SMA, a balanced collagen III:I ratio, and a replenishment of adipocytes. DMFs treated with ASCs+IGF-I transformed into adipocyte-like cells in vitro, which was validated by IGF-I-depletion and IGF-I receptor inhibition.
Conclusion:
ASCs+PRP reduces fibrosis markers and induces adipocyte renewal in vivo. As a PRP component, IGF-I works with ASCs to transform DMFs into adipocyte-like cells in vitro.
Clinical Relevance:
Identifying an active factor in PRP that synergizes with ASCs to transform DMFs into adipocyte-like cells contributes to finding a novel therapeutic for Dupuytren disease. Such a treatment may allow for less extensive surgical intervention coupled with therapeutic injection to reduce the recurrence of Dupuytren disease.
Keywords: adipose-derived stem cells, Dupuytren disease, fibrosis, lipodystrophy, myofibroblasts
Introduction
Dupuytren disease is a common fibrotic condition of the palm and digits. It may lead to flexion deformities of the digits and impaired hand function. Myofibroblasts are the primary cells responsible for the development of the disease 1. Surgical treatments encompassing a variety of approaches, percutaneous release, and collagenase clostridium histolyticum (CCH) are all standard treatments for Dupuytren disease. However, although these modalities have a reasonable safety profile, the tendency for recurrence with all modalities is high and usually occurs within two years of the treatment 2. In Dupuytren disease, there is an increase in total collagen with an increased ratio of type III to type I collagen in the diseased tissue 3. CCH, as a non-surgical treatment, decreases collagen by enzymatically breaking it down. It also decreases α-smooth muscle actin (α-SMA) levels in myofibroblasts in vitro, but its effect is temporary 4.
Anti-fibrotic treatments typically aim to target excess ECM deposition 5,6. However, fibrosis is also associated with the loss or reduction of lipid-laden cells, a process known as lipodystrophy. Loss of these cells critically affects tissue function and integrity, especially in the skin 7. Understanding the mechanisms and implications of fibrotic lipodystrophy is crucial when considering therapeutic strategies in fibrosis 8.
Adipose-derived stem cells (ASCs), as a source of mesenchymal stem cells, are postulated to be a means to decrease fibrosis 9. Adipose tissue is an excellent source for obtaining many ASCs (tens of thousands per ml of lipoaspirate) using liposuction, a minimally invasive procedure 10. In addition, the therapeutic potential of ASCs for anti-fibrosis is enhanced when the ASCs are cultured with platelet-rich plasma (PRP), a concentration of platelets from whole blood containing numerous anti-fibrotic and pro-adipogenic growth factors 11,12. We previously demonstrated that PRP and ASCs provide factors that induce myofibroblasts derived from Dupuytren tissue to transform into adipocyte-like lipid-laden cells in vitro. In addition, this treatment reduced fibrosis markers in a rodent model. However, adipocyte-like cells were not detected in vivo under the treatment regime in the previous study 13.
Thus, in the current study, we first aimed to develop a treatment regime to determine whether the adipocyte-like cells generated in vitro could also be generated in vivo. The notion of lipogenic and myogenic fibroblastic phenotype switching as a mode of marking fibrosis or scarring progression and resolution has been demonstrated previously 14,15. However, from a translational perspective, our method of inducing this transformation using PRP has limitations, mainly related to its composition 16. Thus, as a second aim in this study, we examined components of PRP, as determined via an extensive literature search, to determine if any of these factors resulted in the production of lipid-filled adipocyte-like cells in the presence of the ASC secretome as a replacement for PRP. We selected insulin-like growth factor I (IGF-I) as a potential candidate and then demonstrated lipogenic effects similar to PRP.
MATERIALS AND METHODS
Fibroblast Isolation and Culture
The palmar fascia from discarded surgical tissue collected at the University of California Irvine Medical Center (UCIMC) in the Department of Plastic Surgery (Institutional Review Board (IRB):2019-5468) were used to isolate Dupuytren myofibroblasts (DMFs). The normal human dermal fibroblasts (NHDFs) were purchased from ATCC (Gaithersburg, Maryland). The Dupuytren tissue was trimmed to remove excess fat tissue, soaked in dispase (5U/mL) (Stem Cell Technologies, Seattle, WA), washed in enzyme-free cell dissociation buffer (Millipore, Temecula, CA), and then in HEPES buffered saline (Sigma-Aldrich, St. Louis, MO). The tissue was digested using type II collagenase (Gibco, Waltham, MA) and halted with 10% fetal bovine serum (FBS) (Gibco). The digested tissue was filtered through a cell strainer (Fisher, Waltham, MA), centrifuged, and resuspended in Minimum Essential Media (MEM) (Gibco) containing 10% FBS and 1% penicillin and streptomycin (P/S) (Corning, Glendale, AZ). The cells were grown to confluence and split using trypsin (Gibco). The cells used for the experiments were from passages 4-8.
Characterization of Myofibroblasts
After the myofibroblasts were isolated, the expression of α-SMA was assessed. NHDFs were plated, left untreated, or treated without TGF-β1 treatment (15ng/ml for 48 hr) to induce myofibroblast differentiation, and DMFs were plated simultaneously without treatment. Immunofluorescence was performed by fixing the cells and using a primary antibody to detect α-SMA (Cell Signaling Technologies, Danvers, MA) and fluorescently conjugated secondary antibody (Alexa Fluor 488, Invitrogen, Waltham, MA). The nuclei were counter-stained with DAPI (Millipore). The fluorescence was quantified using ImageJ. The percent of α-SMA positive cells were NHDF 7.6% (±2.1), NHDF+ TGF-β1 29.8% (±4.4), and DMF 43.3% (±9.9), verifying the isolation of myofibroblasts from the Dupuytren disease tissue.
ASC Isolation and Culture
The lipoaspirate was obtained from healthy individuals undergoing elective liposuction at the Department of Plastic Surgery, University of California, Irvine (UCI). The tissue was washed in phosphate-buffered saline (PBS) and digested with type I collagenase I (Stem Cell Technologies). Dulbecco’s Modified Eagle Medium (DMEM) (Corning) supplemented with 10% FBS and 1% P/S was used to stop the digestion. The solution was filtered through a cell strainer (Fisher), centrifuged, and the cell pellet was resuspended in DMEM with 10% FBS and 1% P/S. The ASCs were characterized as described previously 17. The cells were used at passages 4-8.
Dupuytren Animal Model
The isolated DMFs and NHDFs were implanted (8x106 cells/paw) into the left and right forepaws of 27 male nude rats (Rowett Nude (rnu/rnu athymic), respectively, to establish a Dupuytren disease animal model 18 under Institutional Animal Care and Use Committee (IACUC) approval. The rats were randomly allocated to three groups 2 months after implanting the cells. Based on our previous study, a power analysis revealed that we needed two rats to achieve at least 95% power to test our hypothesis using a two-sided paired t-test at 2.5% significance level. Bonferroni correction was applied to account for multiple group comparisons. We selected three animals per group to account for any issues with the experiment. The forepaws were treated with 100 μl of saline or a mix of PRP (1%) + ASCs (1x106 cells/paw in 100 μl), or CCH (0.58mg/ml in 100 μl) (to compare to the clinical treatment) (n=9/group). Our previous in vitro data revealed that lipid-filled cells were only induced when ASCs and PRP were combined13. Therefore, we did not assess these components alone in the rats. The forepaw tissue was harvested after one week for nine rats (n=3/group). The 18 remaining rats received the treatments described above. After another week, the forepaw tissue was harvested from 9 rats (n=3/group), and the nine remaining rats received the last treatments. Finally, one week later, the forepaw tissue from the final nine rats was harvested. The harvested tissues were fixed in formalin and paraffin-embedded following standard protocols. After sectioning, Masson’s trichrome staining (Polysciences; Warrington, PA) was performed to assess collagen (Figure S1).
Immunohistochemistry (IHC) was done to evaluate α-SMA, collagen I, collagen III, and perilipin expressions. Briefly, the sections were blocked in hydrogen peroxide after deparaffinization and rehydration. Antigen retrieval was done using a trypsin enzymatic kit (Abcam, Burlingame, CA). A Mouse and Rabbit Specific HRP/DAB (ABC) Detection IHC kit (Abcam) was used, and the primary antibodies were α-SMA (1:100; Cell Signaling, Danvers, MA), Collagen I alpha 1 (1:100; Novus Biologicals, Centennial, CO), Collagen III alpha 1 (1:100; Novus Biologicals), and perilipin (1:100; Cell Signaling). The slides were mounted and imaged at 4X. Five fields of view per slide were used to quantify the protein expression (ImageJ (NIH)). IHC images were deconvoluted to obtain the DAB-positive area. The mean gray value was used to determine the signal intensity of each image. Intensity units were then converted to Optical Density (OD) utilizing the following formula: OD = log(maximum intensity/mean intensity), where maximum intensity = 255. Adipocyte count and area were calculated using the Adipocyte Tools plugin. Adipocytes were segmented using the percentile thresholding method with a minimum adipocyte size of 100 and a maximum size of 200,000.
Cell Culture and Treatments
DMFs were seeded (0.5x105 cells/well) into 24-well plates. The cells were treated at about 80% confluency (~1.9x105 cells/well). Untreated DMFs served as a negative control. DMFs co-cultured with ASCs (transwell culture system; ASCs were added at 1x105 cells/insert) and treated with PRP (1%) served as a positive control, as determined previously 13. The PRP was acquired from healthy donors collected at UCI in the Department of Dermatology using an Eclipse PRP (3.2 platelet multiplier and leukocyte poor) (Eclipse, The Colony, TX). The PRP (10 donors) was pooled to account for individual variability. For the experimental treatments, DMFs were treated with recombinant human IGF-I (Irvine Scientific, Santa Ana, CA;10ng/ml or 100ng/ml) alone or with the ASCs co-culture. To inhibit IGF, NVP-AEW541 (MedChemExpress, Monmouth Junction, NJ), an inhibitor of the insulin-like growth factor 1 receptor (IGF-1R), or BMS-536924 (MedChemExpress), a competitive and selective IGF-1R inhibitor, was used at 10μM to treat the DMFs. The cells were assessed at four weeks using Oil Red-O (MilliporeSigma, Temecula, CA) or Bodipy (ThermoFisher, Waltham, MA) to reveal lipid droplets. The cells were imaged and analyzed using ImageJ (Figure S1).
IGF Depletion from PRP and Validation
Pooled PRP was rotated with an IGF-I antibody (2.5μg; Peprotech; Cranbury, NJ) or an isotype control antibody overnight at 4°C. Then, protein A/G beads (SellekChem, Houston, TX) were added to the PRP, which was rotated for 20 min at 4°C. The beads were collected using a magnet, and the depleted PRP was collected. An ELISA (RayBioTech, Norcross, GA) was conducted following the manufacturer’s instructions to verify the depletion of IGF from the PRP.
Statistical Analysis
All the quantitative data are expressed as the mean ± SEM. To assess differences between groups, an unpaired, two-tailed t-test was performed. A value of p < 0.05 was considered statistically significant.
Results
Validation of the Dupuytren Rodent Model
Paws implanted with DMFs demonstrated more collagen accumulation than those implanted with NHDFs, as shown by the Masson’s Trichrome staining (Figure 1A). The DMF paws had significantly (p < 0.05) more collagen I (Figures 1A and B) and higher collagen III expression (not significant (NS); p > 0.05) (Figures 1A and C) than those implanted with NHDFs. Furthermore, the type III to I collagen ratio was higher in the DMF paws (Figure 1D). Finally, the expression of α-SMA within the tissue (not associated with the blood vessels) was significantly greater in the DMF paws (p > 0.05) (Figures 1A and E). These increased disease markers in the paws implanted with DMFs compared to those with NHDFs validated the original data and our previous findings 13,18.
Figure 1.
Establishing a Model of Dupuytren Disease In Vivo
(A). NHDFs and DMFs were implanted into the paws of rnu/rnu rats. After about two months, the paw tissue was harvested. Masson’s trichrome and IHC staining was conducted to detect Collagen I and III and α-SMA (n =3/group). Representative images are shown and labeled. Scale bar = 100um. IHC staining was quantified using ImageJ for Collagen I (B) and Collagen III (C). (D) The ratio of the type III/I Collagen was determined. (E) IHC staining was quantified using ImageJ for α-SMA. The data are presented as the mean±standard deviation. * p <0.05 (AU; arbitrary units)
Fibrotic Marker Reduction with ASCs+PRP Treatment
Masson’s trichrome staining was similar across all time points for the DMF paws (a representative image is shown in Figure 2A). After the first two treatments with CCH or ASCs+PRP, there was a reduction in overall collagen. However, after the third treatment with ASCs+PRP, the reduction was greater than the CCH-treated paws (Figure 2A).
Figure 2.
ASCs+PRP Treatment Reduces Fibrosis Markers In Vivo
DMFs were implanted into the paws of rnu/rnu rats. After about two months, the paw tissue was treated with saline, ASC+PRP, or CCH once a week for three weeks. The paw tissue was harvested one week after each treatment. (A). Masson’s trichrome was conducted. A represented image of saline-treated DMF-implanted paws served as a comparison. Scale bar = 100um. IHC was performed to detect Collagen I (B) and III (C). (D) The ratio of Collagen III/I is presented. (E) IHC was performed to detect α-SMA. ImageJ was used to quantify the staining (n =3/group). The data are presented as the mean±standard deviation. *p<0.05; **p<0.01 (AU; arbitrary units)
CCH and ASCs+PRP reduced collagen I expression after the first treatment. However, after the second treatment, collagen I expression increased with CCH but declined with ASCs+PRP compared to the control (NS; p >0.5) (Figure 2B). After one treatment, CCH and ASCs+PRP reduced collagen III expression. The difference between the reduced levels of the treatment groups was significant (p < 0.01); the ASCs+PRP treatment demonstrated a more substantial reduction. For the second treatment, the decrease was significant only for the ASCs+PRP-treated paws. After the third treatment, collagen III expression was unchanged with CCH and lower with ASCs+PRP (p >0.05) (Figure 2C). After the three treatments, the collagen III/I ratio was 1.4 for the DMF group, 1.9 for the CCH treatment, and 1.0 for the ASCs+PRP treatment (Figure 2D). Thus, the overall decrease in collagen was likely due to the reduction of collagen III.
The expression of α-SMA was greatest in the paws implanted with DMF at the 2-treatment timepoint, likely reflecting the highest activity of the myofibroblasts. CCH and ASCs+PRP significantly inhibited α-SMA expression (45% (p < 0.05) and 64% (p < 0.01), respectively) (Figure 2E). However, the inhibition was greater for ASCs+PRP. Thus, treating rodent paws with ASCs+PRP effectively reduced collagen and α-SMA.
Emergence of Adipocytes with ASCs+PRP Treatment
The adipocyte counts for the NHDF paws were significantly greater (p < 0.01) than the DMFs at the 2-week treatment time. CCH treatment did not change the adipocyte counts in the DMF paws. In contrast, ASCs+PRP treatment led to a significant increase (p < 0.01) in adipocyte counts, reaching a number similar to the NHDF paws. After 1 and 3 treatments, the adipocyte count was greatest for the ASCs+PRP group (Figure 3).
Figure 3.
ASCs+PRP Treatment Induces Adipocyte Reemergence In Vivo
DMFs were implanted into the paws of rnu/rnu rats. After about two months, the paw tissue was treated with saline, ASC+PRP, or CCH once a week for three weeks. The paw tissue was harvested one week after each treatment. IHC was performed to detect perilipin. Perilipin served as a marker of adipocytes. Adipocytes were counted using ImageJ and the Adipocyte Tools plugin. The data are presented as the mean±standard deviation. *p<0.05; **p<0.01
IGF-I Replaces PRP Transforming DMFs into Lipid-Filled Cells in the Presence of ASCs
IGF-I (10ng/ml) demonstrated lipogenic effects similar to PRP (ASCs+PRP:87% lipid-laden cells and ASCs+IGF-I 83% lipid-laden cells) (Figure 4). IGF-I was then used at increasing concentrations to optimize the transformation, and 100ng/ml was selected (Figure S2).
Figure 4.
DMFs treated with ASCs+IGF-I Transform into Lipid-Filled Cells
DMFs were left untreated, co-cultured with ASCs and PRP (1%) as a positive control, treated with IGF-I (100ng/ml), or treated with ASCs+IGF-I (100ng/ml). The cells were fixed, stained with Oil Red O, and imaged at 10X. Representative images are shown. The scale bar represents 50 μm.
Next, IGF-I was depleted from the pooled PRP, and the depletion was confirmed by ELISA (p < 0.01) (Figure S3). When DMFs were co-cultured with IGF-I-depleted PRP and ASCs, lipid-laden cells were reduced by 64%. In contrast, the cells treated with ASCs+PRP and ASCs+IGF-I showed significant induction of lipid-filled cells (p < 0.05 and p < 0.01, respectively) (Figure 5A and 5B).
Figure 5.
DMFs treated IGF-Depleted PRP and ASCs Show Reduced Lipid-Filled Cells
DMFs were left untreated, co-cultured with ASCs and PRP (1%) as a positive control, treated with ASCs+IGF-I (100ng/ml), or treated with ASCs and IGF-I-depleted PRP (1%). (A) The cells were subjected to BODIPY staining and imaged at 10X. Representative images are shown. The scale bar represents 50 μm. (B). The staining was quantified using ImageJ. The data are presented as the mean±standard deviation. *p<0.05; **p<0.01
Finally, treating DMFs with IGF-I receptor inhibitor (NVP-AEW541) blocked the induction of lipid-filled cells when DMFs were treated with ASCs+PRP and ASCs+IGF-I. ASCs+PRP showed 84% lipid-laden cells and IGF-I receptor inhibition reduced it to 13%. ASCs+IGF-I showed 80% lipid-laden cells and IGF-I receptor inhibition lowered it to 11% (Figure 6). Similar results were demonstrated with BMS-536924, another IGF-I receptor inhibitor (data not shown).
Figure 6.
IGF-I Receptor Inhibition Prevents DMF Transformation into Lipid-Filled Cells
DMFs were left untreated, co-cultured with ASCs and PRP (1%), treated with ASCs+IGF-I (100ng/ml), or treated with an IGF-I receptor inhibition with the ASCs+PRP or ASCs+IGF-I. The cells were fixed, stained with Oil Red O, and imaged at 10X. Representative images are shown. The scale bar represents 50 μm.
Discussion
Previously, we showed that ASCs+PRP had potential as a treatment for Dupuytren disease in vivo, as demonstrated by a rebalancing of collagen I and III in rodent paws implanted with DMFs. Furthermore, we showed that ASCs+PRP transformed DMFs into lipid-filed adipocyte-like cells 13. Here, we utilized the ASCs+PRP treatment again in a rodent model with altered treatment schedules and incubation times. The rodent paws implanted with DMF and treated with ASCs+PRP demonstrated a rebalance in collagen I and III, reduction in α-SMA, and induction of adipocytes. Emerging evidence indicates that lipid-filled cells secrete fibroprotective factors in tissues, and their loss during fibrotic transformation contributes to tissue dysfunction 19,20. The replenishment of fat cells within these paws after ASCs+PRP occurred after all treatment time points but was significant after two treatments. CCH treatment did not induce adipocytes, which is expected as CCH targets collagen 4. These findings suggest that an injection of ASCs+PRP might reduce the fibrotic condition and replenish lipid-filled cells, decreasing the fibrotic condition.
From a translational perspective, PRP has limitations 16. The PRP utilized in these studies was pooled from 10 donors. In the clinic, this would likely be an autologous treatment. Thus, it was essential to determine which factor in PRP contributed to adipocyte-like cell conversion in the presence of ASCs. We identified that IGF-I functioned like PRP to generate adipocyte-like cells from DMFs co-cultured with ASCs. IGF-I is a viable candidate because it induces adipogenesis in vivo by recruiting endogenous cells 21. Finding a factor from PRP that replaces its function in vitro is a means to alleviate the translational limitations of PRP utilization and offers a target for future mechanistic studies.
Depleting IGF-I from PRP revealed a reduction in the formation of lipid-filled cells when DMFs were co-cultured with ASCs. This slight production of lipid-laden cells by the depleted PRP suggests that another PRP component may also contribute to this process or that the small amount of IGF-I that remained contributed to the process. Two IGF-I receptor inhibitors reduced the ability of IGF-I and ASCs to transform DMFs into lipid-laden cells to a greater degree than IGF-I-depleted PRP. These findings suggest that the DMFs utilize IGF-I receptor-mediated signaling to generate lipid-laden cells. Interestingly, IGF-I alone is not capable of doing this alone. Co-culturing with ASCs was needed. Thus, synergy occurs with IGF-I and an ASC-secreted factor that induces this process. IGF-synergistic action is common. IGF-I synergizes with EGF to promote human esophageal epithelial cells and glucocorticoid to induce myogenesis 22,23.
A proteomics study revealed increased expression of the IGF-I receptor in Dupuytren tissue compared to non-diseased tissue 24. While IGF-I has not been demonstrated to have an anti-fibrotic function, we hypothesize that the ASC-secreted factor synergizing with IGF-I creates an anti-fibrotic response. Assessing the ASC secretome to identify this factor(s) is a logical next step.
We aim to create a cell-free therapy as we develop this Dupuytren disease treatment strategy 25. The ASC secretome is a set of complex soluble factors, including cytokines, chemokines, growth factors, and vesicles. These ASC-secreted factors work in paracrine and have been demonstrated to target lung, liver, and cardiac fibrosis and systemic sclerosis 26–29. Thus, identifying which factor from the ASC secretome works with IGF-I to transform DMFs into adipocyte-like cells would yield a cell-free therapy. Furthermore, this strategy could serve as a means to reduce the need for moderate or radical fasciectomy and limit the surgery to a percutaneous contracture release and biologic injection aimed at reducing fibrosis and replenishing subdermal fat loss and reducing recurrence.
This study has limitations. First, the animal model does not perfectly replicate Dupuytren disease in humans. However, DMFs, as the principal effector cells, were implanted into a physiologically relevant orthotopic anatomical location in rats and monitored over time. Thus, we observed the transition into adipocytes and scar tissue evolution. Using human cells in this model created an incubator to reasonably emulate the human cellular transformation sequence. Second, we must fully characterize the lipid-laden cells generated in vitro. These cells were termed adipocyte-like because they do not possess the morphological features of mature adipocytes. The nature of these cellular transitions during fibrosis and healing is still not fully understood 8.
Dermal fat has largely been overlooked in fibrosis research. It is crucial to healthy skin, and its lipid depletion in a fibrotic context is likely to have a profound impact on skin health. The current study offers mechanisms for discovering new therapeutic targets for treating skin fibrosis, using Dupuytren disease as a model. However, research directed at fibrotic fat loss in the skin may also have implications for other organs that exhibit fibrotic lipodystrophy, such as the lungs and liver 14,30.
Supplementary Material
Figure S3 demonstrates the ELISA results confirming the depletion of IGF-I from PRP.
Figure S1 demonstrated the study design for the 2 phases of the project.
Figure S2 demonstrates optimizing the IGF-I dose used to treat DMFs.
Acknowledgments:
This study was supported by a grant from the American Foundation for Surgery of the Hand (2022 Andrew J. Weiland Basic Science Grant).
References
- 1.Verjee LS, Verhoekx JS, Chan JK, et al. Unraveling the signaling pathways promoting fibrosis in Dupuytren’s disease reveals TNF as a therapeutic target. Proc Natl Acad Sci U S A. 2013;110(10):E928–937. Published online 2013/February/23. doi: 10.1073/pnas.1301100110 [DOI] [PMC free article] [PubMed] [Google Scholar]
- 2.Nayar SK, Pfisterer D, Ingari JV. Collagenase Clostridium Histolyticum Injection for Dupuytren Contracture: 2-Year Follow-up. Clin Orthop Surg. 2019;11(3):332–336. Published online 2019/September/03. doi: 10.4055/cios.2019.11.3.332 [DOI] [PMC free article] [PubMed] [Google Scholar]
- 3.Brickley-Parsons D, Glimcher MJ, Smith RJ, Albin R, Adams JP. Biochemical changes in the collagen of the palmar fascia in patients with Dupuytren’s disease. J Bone Joint Surg Am. 1981;63(5):787–797. Published online 1981/June/01. [PubMed] [Google Scholar]
- 4.Syed F, Thomas AN, Singh S, Kolluru V, Emeigh Hart SG, Bayat A. In vitro study of novel collagenase (XIAFLEX(R)) on Dupuytren’s disease fibroblasts displays unique drug related properties. PLoS One. 2012;7(2):e31430. Published online 2012/March/03. doi: 10.1371/journal.pone.0031430 [DOI] [PMC free article] [PubMed] [Google Scholar] [Retracted]
- 5.Li Y, Zhang W, Gao J, et al. Adipose tissue-derived stem cells suppress hypertrophic scar fibrosis via the p38/MAPK signaling pathway. Stem Cell Res Ther. 2016;7(1):102. Published online 2016/August/04. doi: 10.1186/s13287-016-0356-6 [DOI] [PMC free article] [PubMed] [Google Scholar] [Retracted]
- 6.Spiekman M, Przybyt E, Plantinga JA, Gibbs S, van der Lei B, Harmsen MC. Adipose tissue-derived stromal cells inhibit TGF-beta1-induced differentiation of human dermal fibroblasts and keloid scar-derived fibroblasts in a paracrine fashion. Plast Reconstr Surg. 2014;134(4):699–712. Published online 2014/10/31. doi: 10.1097/PRS.0000000000000504 [DOI] [PubMed] [Google Scholar]
- 7.Shook BA, Wasko RR, Mano O, et al. Dermal Adipocyte Lipolysis and Myofibroblast Conversion Are Required for Efficient Skin Repair. Cell Stem Cell. 2020;26(6):880–895 e886. Published online 2020/April/18. doi: 10.1016/j.stem.2020.03.013 [DOI] [PMC free article] [PubMed] [Google Scholar]
- 8.Jussila A, Zhang B, Kirti S, Atit R. Tissue fibrosis associated depletion of lipid-filled cells. Exp Dermatol. 2024;33(3):e15054. Published online 2024/March/23. doi: 10.1111/exd.15054 [DOI] [PMC free article] [PubMed] [Google Scholar]
- 9.Vanderstichele S, Vranckx JJ. Anti-fibrotic effect of adipose-derived stem cells on fibrotic scars. World J Stem Cells. 2022;14(2):200–213. Published online 2022/April/19. doi: 10.4252/wjsc.v14.i2.200 [DOI] [PMC free article] [PubMed] [Google Scholar]
- 10.Fraser JK, Wulur I, Alfonso Z, Hedrick MH. Fat tissue: an underappreciated source of stem cells for biotechnology. Trends Biotechnol. 2006;24(4):150–154. Published online 2006/February/21. doi: 10.1016/j.tibtech.2006.01.010 [DOI] [PubMed] [Google Scholar]
- 11.Kalen A, Wahlstrom O, Linder CH, Magnusson P. The content of bone morphogenetic proteins in platelets varies greatly between different platelet donors. Biochem Biophys Res Commun. 2008;375(2):261–264. Published online 2008/August/19. doi: 10.1016/j.bbrc.2008.08.014 [DOI] [PubMed] [Google Scholar]
- 12.McLaughlin M, Gagnet P, Cunningham E, et al. Allogeneic Platelet Releasate Preparations Derived via a Novel Rapid Thrombin Activation Process Promote Rapid Growth and Increased BMP-2 and BMP-4 Expression in Human Adipose-Derived Stem Cells. Stem Cells Int. 2016;2016:7183734. Published online 2016/January/30. doi: 10.1155/2016/7183734 [DOI] [PMC free article] [PubMed] [Google Scholar]
- 13.Ziegler ME, Staben A, Lem M, et al. Targeting Myofibroblasts as a Treatment Modality for Dupuytren Disease. J Hand Surg Am. 2023;48(9):914–922. Published online 2023/July/23. doi: 10.1016/j.jhsa.2023.06.007 [DOI] [PubMed] [Google Scholar]
- 14.El Agha E, Moiseenko A, Kheirollahi V, et al. Two-Way Conversion between Lipogenic and Myogenic Fibroblastic Phenotypes Marks the Progression and Resolution of Lung Fibrosis (vol 20, pg 261, 2017). Cell Stem Cell. 2017;20(4):571–571. doi: 10.1016/j.stem.2017.03.011 [DOI] [PubMed] [Google Scholar]
- 15.Plikus MV, Guerrero-Juarez CF, Ito M, et al. Regeneration of fat cells from myofibroblasts during wound healing. Science. 2017;355(6326):748–752. Published online 2017/January/07. doi: 10.1126/science.aai8792 [DOI] [PMC free article] [PubMed] [Google Scholar]
- 16.Nazaroff J, Oyadomari S, Brown N, Wang D. Reporting in clinical studies on platelet-rich plasma therapy among all medical specialties: A systematic review of Level I and II studies. PLoS One. 2021;16(4):e0250007. Published online 2021/April/24. doi: 10.1371/journal.pone.0250007 [DOI] [PMC free article] [PubMed] [Google Scholar]
- 17.Banyard DA, Salibian AA, Widgerow AD, Evans GR. Implications for human adipose-derived stem cells in plastic surgery. J Cell Mol Med. 2015;19(1):21–30. Published online 2014/November/27. doi: 10.1111/jcmm.12425 [DOI] [PMC free article] [PubMed] [Google Scholar]
- 18.Satish L, Palmer B, Liu F, et al. Developing an animal model of Dupuytren’s disease by orthotopic transplantation of human fibroblasts into athymic rat. BMC Musculoskelet Disord. 2015;16:138. Published online 2015/June/08. doi: 10.1186/s12891-015-0597-z [DOI] [PMC free article] [PubMed] [Google Scholar]
- 19.Fang F, Liu L, Yang Y, et al. The adipokine adiponectin has potent anti-fibrotic effects mediated via adenosine monophosphate-activated protein kinase: novel target for fibrosis therapy. Arthritis Research & Therapy. 2012;14(5)doi:ARTN R229 10.1186/ar4070 [DOI] [PMC free article] [PubMed] [Google Scholar]
- 20.Jenke A, Schur R, Röger C, et al. Adiponectin attenuates profibrotic extracellular matrix remodeling following cardiac injury by up-regulating matrix metalloproteinase 9 expression in mice. Physiol Rep. 2017;5(24)doi:ARTN e13523 10.14814/phy2.13523 [DOI] [PMC free article] [PubMed] [Google Scholar]
- 21.Hu L, Yang G, Hagg D, et al. IGF1 Promotes Adipogenesis by a Lineage Bias of Endogenous Adipose Stem/Progenitor Cells. Stem Cells. 2015;33(8):2483–2495. Published online 2015/May/27. doi: 10.1002/stem.2052 [DOI] [PMC free article] [PubMed] [Google Scholar]
- 22.Pansters NA, Langen RC, Wouters EF, Schols AM. Synergistic stimulation of myogenesis by glucocorticoid and IGF-I signaling. J Appl Physiol. 2013;114(9):1329–1339. doi: 10.1152/japplphysiol.00503.2012 [DOI] [PubMed] [Google Scholar]
- 23.Qureshi FG, Tchorzewski MT, Duncan MD, Harmon JW. EGF and IGF-I synergistically stimulate proliferation of human esophageal epithelial cells. J Surg Res. 1997;69(2):354–358. doi:DOI 10.1006/jsre.1997.5080 [DOI] [PubMed] [Google Scholar]
- 24.Pavelic SK, Sedic M, Hock K, et al. An integrated proteomics approach for studying the molecular pathogenesis of Dupuytren’s disease. J Pathol. 2009;217(4):524–533. doi: 10.1002/path.2483 [DOI] [PubMed] [Google Scholar]
- 25.Hu C, Zhao L, Zhang L, Bao Q, Li L. Mesenchymal stem cell-based cell-free strategies: safe and effective treatments for liver injury. Stem Cell Res Ther. 2020;11(1):377. Published online 2020/September/05. doi: 10.1186/s13287-020-01895-1 [DOI] [PMC free article] [PubMed] [Google Scholar]
- 26.Driscoll J, Patel T. The mesenchymal stem cell secretome as an acellular regenerative therapy for liver disease. J Gastroenterol. 2019;54(9):763–773. Published online 2019/July/05. doi: 10.1007/s00535-019-01599-1 [DOI] [PMC free article] [PubMed] [Google Scholar]
- 27.Li X, Ma T, Sun J, et al. Harnessing the secretome of adipose-derived stem cells in the treatment of ischemic heart diseases. Stem Cell Res Ther. 2019;10(1):196. Published online 2019/June/30. doi: 10.1186/s13287-019-1289-7 [DOI] [PMC free article] [PubMed] [Google Scholar]
- 28.Vasse GF, Van Os L, De Jager M, et al. Adipose Stromal Cell-Secretome Counteracts Profibrotic Signals From IPF Lung Matrices. Front Pharmacol. 2021;12:669037. Published online 2021/August/17. doi: 10.3389/fphar.2021.669037 [DOI] [PMC free article] [PubMed] [Google Scholar]
- 29.Velier M, Simoncini S, Abellan M, et al. Adipose-Derived Stem Cells from Systemic Sclerosis Patients Maintain Pro-Angiogenic and Antifibrotic Paracrine Effects In Vitro. J Clin Med. 2019;8(11)Published online 2019/November/20. doi: 10.3390/jcm8111979 [DOI] [PMC free article] [PubMed] [Google Scholar]
- 30.Hernandez-Gea V, Friedman SL. Pathogenesis of liver fibrosis. Annu Rev Pathol. 2011;6:425–456. Published online 2010/November/16. doi: 10.1146/annurev-pathol-011110-130246 [DOI] [PubMed] [Google Scholar]
Associated Data
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Supplementary Materials
Figure S3 demonstrates the ELISA results confirming the depletion of IGF-I from PRP.
Figure S1 demonstrated the study design for the 2 phases of the project.
Figure S2 demonstrates optimizing the IGF-I dose used to treat DMFs.