Taro Matsumoto M.D., Ph.D., Department of Medical Science, Division of Cell Regeneration and Transplantation, Nihon University School of Medicine, 30-1, Oyaguchi, Kami-cho, Itabashiku, Tokyo 173-8610, Japan. Email: firstname.lastname@example.org
Objectives: To examine the effects of mature adipocyte-derived dedifferentiated fat (DFAT) cell transplantation on urethral tissue regeneration and sphincter function.
Methods: Sixteen female Sprague–Dawley rats underwent vaginal distension (VD) for 3 h. Subsequently, green fluorescence protein (GFP)-labeled DFAT cells (1 × 106 in 20 µL saline, DFAT group, n = 8) or saline (20 µL, control group, n = 8) were injected into paraurethral connective tissue. Two weeks following VD, leak point pressure (LPP) was measured and an immunohistochemical analysis of the urethra was performed to evaluate urethral sphincter regeneration.
Results: The VD model was characterized by atrophy of the urethral sphincter and showed a decrease in LPP. DFAT cell transplantation resulted in a significant improvement of LPP (DFAT group: 37.3 ± 6.4 vs control group: 21.7 ± 5.7 mmHg, P < 0.01). Immunohistochemistry revealed that the striated muscle thickness and smooth muscle α−actin-positive area were significantly (P < 0.05) larger in the DFAT group than in the control group. DFAT cell transplantation enhanced macrophage accumulation followed by an increased number of cells in the proliferative state. Transplanted DFAT cells were observed in the damaged smooth muscle layer and showed positive staining for smooth muscle α−actin, suggesting conversion into the smooth muscle cell phenotype.
Conclusions: DFAT cell transplantation promotes sphincter muscle regeneration and improves LPP in the rat VD model.
Stress urinary incontinence (SUI) is one of the most prevalent forms of incontinence in women. Anatomical injury during childbirth is the most important risk factor for the development of SUI in later life.1 Rat models of childbirth made by vaginal distension (VD) demonstrate the development of SUI symptoms, as evidenced by lowered leak point pressure (LPP) on urodynamic testing.2
Adult stem cell therapy holds great potential as a new therapeutic strategy for the regeneration of damaged tissues, including the urinary tract. Recent reports show that an injection of muscle-derived stem cells (MDSC) and adipose-derived stem or stromal cells (ASC) facilitates the regeneration of sphincter muscles in a rat model with an impairment of urethral contraction.3 Since MDSC and ASC can be obtained easily in large quantities under local anesthesia, these cells exhibit potential advantages in cell therapy applications in patients with SUI. However, these cells are known to be highly heterogeneous, as they include fibroblasts, endothelial cells and other types of cells, because these cells are only a minor part of the cell population in stromal cell fractions extracted from muscle or adipose tissue.4,5 Therefore, other adult stem cell sources that can be easily isolated and expanded with high purity are still needed.
Mature adipocytes are the most abundant cell type in adipose tissue and are easily isolated without a painful procedure. It has been shown that mature adipocytes isolated from fat tissue can dedifferentiate into fibroblast-like cells with an in vitro dedifferentiation strategy, known as ceiling culture.6 Our group has established an adipogenic progenitor cell line derived from mature adipocytes, named dedifferentiated fat (DFAT) cells,7 which have shown the potential to differentiate into lineages of mesenchymal tissue similar to bone marrow mesenchymal stem cells.7,8 More recently, we reported that DFAT cells could differentiate into smooth muscle-like cells and contribute to bladder tissue regeneration.9 DFAT cells are easily isolated from a small amount (approximately 1 g) of subcutaneous adipose tissue and are readily expanded with high purity, regardless of the donor's age. These findings suggest that DFAT cells are applicable to many cell-based therapies for organ failure, including SUI.
In the present study we examined the effects of DFAT cell transplantation on LPP and urethra tissue regeneration in the VD model to explore the feasibility of DFAT cell-based therapy in patients with SUI.
The 8-week-old female Sprague–Dawley (SD) rats were purchased from CLEA Japan (Tokyo, Japan). All animal experiments were approved by the Animal Research and Care Committee at the Nihon University School of Medicine.
Cell isolation and culture
Green fluorescent protein (GFP)-labeled DFAT cells10 prepared from GFP transgenic rats (SD TgN [act-EGFP] OsbCZ-004) were used in the experiments. The preparation of DFAT cells from adipose tissue was performed as described previously.8 Briefly, approximately 1 g of isolated s.c. fat tissue was minced and digested using 0.1% collagenase solution (Collagenase type I; Koken, Tokyo, Japan) at 37°C for 1 h with gentle agitation. After filtration and centrifugation, the floating top layer containing unilocular adipocytes was collected. After being washed with phosphate-buffered saline (PBS), the cells (5 × 104) were placed in 25-cm2 culture flasks filled completely with Dulbecco's modified Eagle's medium (DMEM, Invitrogen, Carlsbad, CA, USA) supplemented with 20% fetal bovine serum (FBS, JRH Bioscience, Lenexa, KS, USA) and were incubated for 7 days. The adipocytes floated up and adhered to the top inner ceiling surface of the flask followed by their conversion to fibroblast-like DFAT cells. Subsequently, the medium was removed and the flasks were inverted so that the cells were on the bottom. The medium was changed every 3 days until the cells reached confluence. The isolation of rat dermal fibroblasts was performed as described previously.11
Differentiation assay in vitro
The differentiation of DFAT cells into the smooth muscle cell (SMC) lineage was performed as described previously.9 Briefly, cells were plated in 12-well dishes at a density of 5 × 104 cells, and incubated for one week in 5% FBS/DMEM containing 5 ng/mL transforming growth factor (TGF)-β1 (R&D Systems, Minneapolis, MN, USA). After the induction, cells were fixed, permeabilized, and incubated with mouse anti-smooth muscle α−actin (ASMA, DakoCytomation, Glostrup, Denmark) or anti-calponin-1 (Santa Cruz Biotechnology, Santa Cruz, CA, USA) antibodies, followed by anti-mouse IgG-Alexa-594 or anti-mouse IgG-Alexa-488 (Invitrogen) secondary antibodies. The nuclei were stained with Hoechst33342. Staining was visualized and photographed with a fluorescence microscope (Eclipse TE 2000-U; Nikon, Tokyo, Japan). Data were expressed as the percentage of ASMA-positive cells/total cells (nuclei) from 5 fields (100 × magnification) per well.
Cytokine antibody array
DFAT cells were plated in 60-mm dishes at a density of 5 × 104 cells, and incubated with 10% FBS/DMEM or 5% FBS/DMEM containing 5 ng/mL TGF-β1 for 1 week. The cells were then incubated with 0.5% FBS/DMEM for 3 days. Conditioned media were taken and analyzed for cytokines using an antibody array (RayBiotech, Norcross, GA, USA). The cytokines were measured according to the instruction manual. The signal was detected using a chemiluminescence imaging system (LAS-3000; Fujifilm, Tokyo, Japan) and quantified by Multi Gauge software (Fujifilm).
Rat VD model
A urethral sphincter injury by VD was made as described previously.2 Briefly, female SD rats were anesthetized with an isoflurane inhalation. A trimmed 10-Fr Foley catheter was then inserted into the vagina and the balloon was inflated with 3 mL of water for 3 h. A single 4-0 Vicryl stitch (Ethicon, Somerville, NJ, USA) was placed in the skin near the vagina to secure the catheter in place. After the balloon inflation was completed, the stitch and catheter were removed. At the indicated time period after VD, LPP testing was done with a modification of the method described previously.12 To avoid voiding reflex, we applied abdominal pressure gently and carefully during the testing. The LPP was tested at least 10 times on each rat and mean values were presented.
The urethra was harvested at the indicated time period, fixed, embedded in paraffin and sectioned (5 µm). The samples were incubated with rabbit anti-ASMA (Abcam, Cambridge, MA, USA) and mouse anti-sarcomeric actin (DakoCytomation) antibodies. In the other experiments the samples were incubated with rabbit anti-rat macrophage (Inter Cell Technologies, Jupiter, FL, USA) and mouse anti-topoisomerase IIα (DakoCytomation) antibodies. After three washes with PBS the samples were incubated with anti-mouse IgG-Alexa 488 and anti-rabbit IgG-Alexa 594 antibodies. The nuclei were stained with Hoechst33342. For the quantitative evaluation, tissue sections of the mid-urethra at the level of the striated muscle layer with maximum transverse diameter were selected in each sample. Following immunostaining, the diameter of the sarcomeric actin-positive layers and the area of the ASMA-positive regions were measured using a computerized digital morphometric analysis system (Adobe Photoshop CS3 extended, San Jose, CA, USA). The number of macrophage-positive cells, topoisomerase IIα−positive cells and apoptotic cells in muscle layers of the urethra were also counted from five randomly chosen fields (200 × magnification) in each sample. Apoptosis was determined by nuclear morphology in cells with condensed chromatin or fragmented nuclei after Hoechst33342 staining and it was confirmed by active caspase staining (Poly-Caspases Detection Kits; Immunochemistry Technologies, Bloomington, MN, USA).
DFAT cell transplantation
A total of 16 female SD rats underwent VD for 3 h. Subsequently, 1 × 106 DFAT cells in 20 µL saline (DFAT group, n = 8) or 20 µL saline alone (control group, n = 8) was administered by injection with a Hamilton microsyringe (Hamilton, Reno, NV, USA) into the paraurethral connective tissue at the mid-urethra, which was located at the level of the pubis symphysis. LPP testing was performed in both groups at 14 days after the administration. The urethras were then harvested, embedded in paraffin, and sectioned (5 µm). The samples were analyzed by immunohistochemistry as described in the immunohistochemistry section. To detect transplanted DFAT cells and their ability to undergo SMC differentiation, the sections were incubated with rabbit anti-GFP (Medical & Biological Laboratories, Nagoya, Japan) and mouse anti-ASMA antibodies, followed by anti-rabbit IgG-Alexa-488 and anti-mouse IgG-Alexa-594 secondary antibodies.
All LPP and histological data are presented as means ± SD. The Mann–Whitney U-test was used to compare parameters between the groups of rats, with P < 0.05 considered as statistically significant (Graphpad Prism ver 5.0) (Graphpad Software, La Jolla, CA, USA).
Properties of rat DFAT cells
We have previously reported that human DFAT cells efficiently (over 50%) differentiated into the SMC phenotype when the cells were incubated with 5% FBS/DMEM containing 5 ng/mL TGF-β1. Therefore, we first examined whether rat DFAT cells could differentiate into SMC lineages in response to the same culture conditions. The results showed that approximately 70% of DFAT cells exhibited ASMA-positive staining during the differentiation culture (Fig. 1). ASMA-positive cells were induced less efficiently (approximately 18%) when rat dermal fibroblasts were incubated with the differentiation media. The expression of calponin-1, a mature SMC marker, was also induced in DFAT cells during the differentiation culture (Fig. 1a).
We next investigated cytokine productions in supernatants of the rat DFAT cell culture by protein array analysis. As shown in Figure 2, DFAT cells predominantly secreted monocyte chemotactic protein-1, macrophage inflammatory protein (MIP)-3α, tissue inhibitor of metalloproteinase-1, lipopolysaccharide-induced C-X-C chemokine and vascular endothelial growth factor. Increased levels of these cytokines were detected in conditioned media subsequent to the SMC differentiation of DFAT cells. On the other hand, rat fibroblasts secreted smaller amounts of these cytokines (data not shown). These findings suggest that rat DFAT cells have an ability to differentiate into the SMC lineage with high efficiency and release soluble factors that stimulate lymphocyte migration, collagen production, and angiogenesis.
Characterization of the VD model
We next examined the functional and histological changes after VD in SD rats. After VD, LPP was significantly (P < 0.05) decreased by day 7, a level that was sustained until day 14, and subsequently recovered by day 28 (Fig. 3a). Immunohistochemistry revealed that VD resulted in the destruction and degenerative change in urethral sphincter at 3 days after VD. A marked atrophic change of smooth muscle and a thinning of striated muscle layers was observed on day 7 and lasted until day 14 (Fig. 3b–d). On day 28 the atrophy recovered and the thickness of the striated muscle layers had become comparable with non-injured muscle layers, although smooth muscle fibers often showed an irregular shape and varied size. The number of macrophages and apoptotic cells was increased in urethra tissue by day 3 after VD and reached a maximum at day 7, then gradually decreased (Fig. 3e,f). The number of topoisomerase IIα-positive cells, indicating the proliferative state of the cells, was predominantly increased by day 14 and then had rapidly decreased by day 28 (Fig. 3g). These findings indicate that VD causes a decrease in LPP with the atrophic change of striated muscle and smooth muscle layers but these changes were reversible by 28 days after VD.
DFAT cell implantation in VD injured urethra
We next examined the effects of DFAT cell transplantation on LPP and urethra tissue regeneration in the VD model. The results showed that LPP in the DFAT group (38.0 ± 5.6 mmHg) was significantly higher than that in the control group (20.5 ± 5.5 mmHg, P < 0.01) (Fig. 4a). Immunohistochemistry revealed that inner smooth muscle and external striated muscle layers in mid-urethras showed atrophy in the control group, whereas DFAT cell transplantation led an increase of smooth muscle and striated muscle mass with variable fiber orientation (Fig. 4b). Morphometric analysis revealed that striated muscle thickness and ASMA-positive areas were significantly (P < 0.05) larger in the DFAT group than in the control group (Fig. 4c,d). These findings indicated that DFAT cell transplantation promotes sphincter muscle regeneration and improves LPP in the VD model. The numbers of macrophages and topoisomerase IIα-positive cells in the mid-urethra tissue were significantly (P < 0.05) increased in the DFAT group in comparison with the control group (Fig. 5a,b). The number of apoptotic cells had decreased by approximately 50% in the DFAT group compared to the control group, although there were no statistically significant differences between the groups (Fig. 5c).
To identify the transplanted cells, immunohistochemistry of urethras from DFAT cell-transplanted rats was performed. The results showed that GFP-positive cells were efficiently distributed in injured smooth muscle layers (Fig. 6a), although the number of grafted cells was far fewer than in the observations of injected cells. The merged view showed that a large number of GFP-positive cells expressed ASMA (Fig. 6b). These data suggest that transplanted DFAT cells can convert to the SMC phenotype in vivo.
The VD model is widely used to study mechanisms of injury and tissue recovery of SUI, because the model can induce histological similarity to the injury that occurs naturally in childbirth and can be set up easily with reproducible results.13 In our pilot study, the model exhibited very similar levels of decreasing LPP and pathological changes in urethral tissue. Therefore we employed the VD model in the present study despite the less durable damage resulting from our technique. Our data as well as previous reports2,14 demonstrate that VD induces an extensive disruption of sphincter muscle layers including striated muscle and smooth muscle within a few days. Damaser et al.15 demonstrated that VD results in decreased blood flow and hypoxia of urogenital organs including the urethra, suggesting that hypoxic injury is a possible mechanism of injury leading to SUI. The increased level of hypoxia-inducible factor 1α expression in the urethra after VD also supports this hypothesis.16 VD leads to atrophy of the sphincter muscle and the increased number of apoptotic cells in the damaged urethra is probably due to urethral hypoxia. Our data showed that the number of macrophages was increased by day 3 after VD and reached a maximum at day 7, and subsequently the number of topoisomerase IIα-positive cells had increased by day 14. The atrophy of the urethral sphincter muscle subsequently recovered by day 28. These data suggest that VD induces reversible muscle damage with macrophage migration in the early phase, followed by an increase in the proliferative state of cells towards recovery of the urethral sphincter. These findings correspond to the time course of wound healing.17 It has been shown that infiltrated macrophages in the early phase play an important role in directing the wound healing process.18
In the VD model we showed that DFAT cell transplantation promoted the recovery of sphincter muscle tissue and improved LPP. Histologically, we found that a number of transplanted DFAT cells expressed ASMA, suggesting conversion to the SMC phenotype. Similar results were obtained in our previous DFAT cell transplantation study in a mouse bladder cryo-injury model.9 In this model, injected DFAT cells survived for at least 30 days and most expressed ASMA. These data, combined with the present data, suggest that transplanted DFAT cells can differentiate into SMC and contribute to improving LPP.
It has been reported that ASC can also differentiate into functional SMC with contracting capability.19 Interestingly, we could not find GFP/sarcomeric actin-double positive cells in any of the sections we examined, suggesting that DFAT cells cannot spontaneously differentiate into the skeletal muscle cell lineage. Our group showed that DNA demethylation using 5-azacytidine is required for the conversion of DFAT cells into the skeletal muscle cell phenotype in vitro.20 Certain pre-treatment of DFAT cells to induce genetic reprogramming may be needed to differentiate DFAT cells into striated muscle cells in vivo. We also found that DFAT cell transplantation promoted the intrinsic regeneration of sphincter muscle with macrophage accumulation. These findings suggest that DFAT cells also contribute to the regeneration of the sphincter muscle indirectly, promoting the wound healing process and angiogenesis and stimulating the differentiation of the remaining progenitor cells by paracrine mechanisms, as reported in other type of stem cells.21 Our protein array data (Fig. 2) showing that DFAT cells release a variety of soluble factors support this hypothesis.
DFAT cells have several properties that make them well suited for regenerative medicine in urological diseases. Compared to other stem cells such as ASC, we found that DFAT cells are quite homogeneous.8 This property of DFAT cells may lead to higher safety and efficacy for clinical cell therapies. In addition, DFAT cells can be obtained from donors regardless of age. In our study using human adipose tissue from a total of 18 donors ranging from 4 to 81 years of age, we successfully prepared DFAT cells from all the donors.8 The differentiation potential for smooth muscle-like cells was confirmed in all the donors we examined. These findings suggest that DFAT cells can be used for autologous transplantation in patients of various ages. Further studies are needed to evaluate the safety, functionality, and long-term viability of DFAT cell transplants using the model of durable urethral dysfunction.
DFAT cell transplantation promoted the regeneration of damaged urethral sphincters and improved LPP in the VD model rats. The transplanted DFAT cells were able to convert into the SMC phenotype and promoted macrophage accumulation followed by cell proliferation in the damaged urethral tissue. Because adipose tissue is abundant and easily accessible at most ages in humans, DFAT cell transplantation may be an attractive therapeutic strategy against SUI.
This work was supported by financial grants from the Ministry of Education, Science, Sports, and Culture of Japan (23390190, 80183648 and S0801033), by financial grants from the Japan Science and Technology Agency (08030216), and by a Nihon University Multidisciplinary Research Grant (10–027 and 11–017). We are grateful to Dr Noriyoshi Konuma, Dr Takayuki Masuko and Dr Zena Al-Bakri, for technical assistance with immunohistochemistry and Dr Chii Yamamoto with the cytokine array.