Research Article | Open Access
Wnt-GSK3β/β-Catenin Regulates the Differentiation of Dental Pulp Stem Cells into Bladder Smooth Muscle Cells
Smooth muscle cell- (SMC-) based tissue engineering provides a promising therapeutic strategy for SMC-related disorders. It has been demonstrated that human dental pulp stem cells (DPSCs) possess the potential to differentiate into mature bladder SMCs by induction with condition medium (CM) from bladder SMC culture, in combination with the transforming growth factor-β1 (TGF-β1). However, the molecular mechanism of SMC differentiation from DPSCs has not been fully uncovered. The canonical Wnt signaling (also known as Wnt/β-catenin) pathway plays an essential role in stem cell fate decision. The aim of this study is to explore the regulation via GSK3β and associated downstream effectors for SMC differentiation from DPSCs. We characterized one of our DPSC clones with the best proliferation and differentiation abilities. This stem cell clone has shown the capacity to generate a smooth muscle layer-like phenotype after an extended differentiation duration using the SMC induction protocol we established before. We further found that Wnt-GSK3β/β-catenin signaling is involved in the process of SMC differentiation from DPSCs, as well as a serial of growth factors, including TGF-β1, basic fibroblast growth factor (bFGF), epidermal growth factor (EGF), hepatocyte growth factor (HGF), platelet-derived growth factor-homodimer polypeptide of B chain (BB) (PDGF-BB), and vascular endothelial growth factor (VEGF). Pharmacological inhibition on the canonical Wnt-GSK3β/β-catenin pathway significantly downregulated GSK3β phosphorylation and β-catenin activation, which in consequence reduced the augmented expression of the growth factors (including TGF-β1, HGF, PDGF-BB, and VEGF) as well as SMC markers (especially myosin) at a late stage of SMC differentiation. These results suggest that the canonical Wnt-GSK3β/β-catenin pathway contributes to DPSC differentiation into mature SMCs through the coordination of different growth factors.
A range of injuries or diseases including cancer, benign bladder contracture, and congenital anomalies (such as bladder exstrophy and myelomeningocele) can result in the damage or loss of the bladder [1–3]. Consequently, patients with those illnesses require bladder augmentation cystoplasty or replacement. However, current cystoplasty using gastrointestinal segments cannot completely restore the normal function of the Detrusor muscle, in some cases even leading to complications, such as metabolic disturbances, urolithiasis, infection, and malignant diseases [4, 5]. Therefore, bladder smooth muscle cell-based tissue engineering becomes one of the most promising remedies for restoring bladder organ function of the urinary system [6, 7]. However, due to the limited expansion of bladder tissue-derived SMCs and highly potential oncogenic risk, the progress of bladder tissue engineering is hampered by lack of a stable SMC source [8, 9]. Thus, search for an alternative cell source is essential for bladder tissue engineering, which could provide a new way to overcome the shortcomings of the aforementioned methods in the future.
Mesenchymal stem cells (MSCs) are one type of fibroblast-like cell population with potential extensive self-renewal and multilineage differentiation abilities . Compared with other MSCs derived from the bone marrow , adipose tissue , peripheral blood , and umbilical cord blood , MSCs derived from dental pulp tissues have marked advantages of easy access with the least invasive procedures and capacities of high proliferation, excellent regeneration, and multiple potential of differentiation along with little inherent immunogenicity, which make them particularly suitable for tissue engineering and gene therapy applications [15, 16]. Dental pulp stem cells (DPSCs) are normally isolated using a single colony method as DPSCs from pulp tissue are heterogeneous, which are expected to produce complex biological activities [17, 18]. For this reason, different clones of DPSCs exhibit different behaviours including various capacities of proliferation and differentiation according to the age of the donor, method of isolation, and conditions of the pulp tissue . DPSCs provide a potential source of progenitor cells for tissue engineering. However, the abilities of expansion and differentiation in vitro should be fully explored before use. In this study, we have isolated three DPSC clones from different patients. The clones were investigated by comparing their proliferation rates and potential to differentiate into three mesenchymal lineages (namely, osteogenic, adipogenic, and chondrogenic), to determine the best clone as the candidate cell source for further tissue engineering research.
We have recently reported the feasibility of using human DPSCs as bladder SMC progenitors for the regeneration of SMCs . Although the capacity of DPSC differentiation into SMCs has been demonstrated, whether they can form a smooth muscle layer and its underlying molecular mechanisms remains largely unknown. The Wnt signaling pathway is an ancient and evolutionarily conserved pathway which orchestrates a range of biological processes, such as cell fate determination during embryonic development, cell proliferation, cell cycle arrest, differentiation, and apoptosis, as well as tissue homeostasis . β-Catenin is a plasma membrane-associated protein that acts as an intracellular signaling transducer in the Wnt signaling pathway . It has been demonstrated to induce the myogenic differentiation of rat MSCs through upregulation of myogenic regulatory factors . Glycogen synthase kinase 3β (GSK3β) is a proline-directed serine-threonine kinase, and its phosphorylation appears to be a critical step in directing β-catenin to the nucleus . Myogenic growth factors, including platelet-derived growth factor-homodimer polypeptide of B chain (PDGF-BB), transforming growth factor-β1 (TGF-β1), and vascular endothelial growth factor (VEGF), as well as hepatocyte growth factor (HGF), have been proven to play a vital role in SMC differentiation in vivo . Therefore, the aim of this study is to analyse the mechanisms of the Wnt signaling pathway and the expression of myogenic growth factors involved in the regulation of differentiation of DPSCs toward bladder SMCs using the in vitro model we established before.
2. Materials and Methods
2.1. Human DPSC Clones and SMC Isolation
The pulp tissues were obtained from third molars (donors aged from 17 to 20 years) with the patient’s informed consent and ethical approval by the South East Wales Research Ethics Committee of the National Research Ethics Service (permission number: 07/WESE04/84). The clonal populations of DPSCs were isolated using a fibronectin-based selection protocol as described previously [20, 25] after ethical approval and patient consent (permission number: 07/WESE04/84). Following 12 days of culture, single cell-derived clones were isolated using cloning rings and accutase digestion and then expanded. Three clones were selected, named as A11, B11, and A32. The level of population doublings (PDs) during expansion culture was monitored to measure the proliferation rate of the three clones . Then, the three clones were induced to differentiate into three mesenchymal lineages (including osteogenic, adipogenic, and chondrogenic) in appropriate differentiation condition in vitro to compare their capacities of differentiation.
Human SMCs were obtained as reported previously from the bladder of patients who underwent open procedures for their bladder, after patient consent and ethical approval by the South East Wales Research Ethics Committee of the National Research Ethics Service (permission number: 07/WESE04/84) . Briefly, bladder muscle tissue was minced into 1 × 1 mm pieces and digested in collagenase type IV enzyme (Sigma-Aldrich) for 30 minutes at 37°C. The digested muscle tissues were plated in Dulbecco’s modified Eagle’s medium (DMEM) with 10% FBS for establishing the primary culture.
2.2. Differentiation of Human DPSC Clone A32 and Wnt Pathway Inhibition Assay
Differentiation of the A32 was induced by using conditioned medium (CM) collected from bladder SMC culture, supplemented with transforming growth factor beta 1 (TGF-β1), as previously described .
The Wnt pathway was analysed by using the inhibitors including XAV939 (Wnt/β-catenin signaling inhibitor, Sigma), SB216763 (GSK-3 inhibitor, Sigma), and LiCl (GSK-3 inhibitor, Sigma), as previously described [26–28]. Briefly, cells were seeded into a 6-well plate. At 80% confluence of cells, the culturing medium was changed into the bladder SMC differentiation induction medium with the XAV939 (5 μM), SB216763 (20 μM), or LiCl (2.5 mM). An equivalent amount of dimethylsulfoxide was added to the control wells. After 14 days of incubation, the mRNA and protein levels were compared to the control group using qPCR and western blotting methods, respectively.
The cells were fixed with 4% PFA for 30 min and then incubated in 0.1% Triton X-100 for 10 min on ice and then blocked with bovine serum albumin (BSA) for 60 min at 37°C. After the blocking step, the cells were incubated with antimyosin (1 : 50), anti-α-SMA (1 : 100), and antidesmin (1 : 50) at 4°C, overnight; PBS was used as the negative control. The cells were then washed with PBS 3 times and incubated with anti-mouse IgG Alexa Fluor-488 or Alexa Fluor-594 secondary antibodies for 1 h at room temperature. The nuclei were counterstained with DAPI (VectorLabs). The protein expressions were observed with a fluorescent microscope and analysed with ImageJ software.
2.4. Flow Cytometry
The cells were washed and resuspended in PBS supplemented with 3% FBS that contained saturating concentrations (1 : 100 dilution) of the following reagents: FITC-conjugated anti-human monoclonal antibodies, anti-CD29-phycoerythrin (PE), anti-CD90-PE, anti-CD34-PE, anti-CD45-PE, anti-CD146-PE, or anti-STRO-1-allophycocyanin (APC) for 1 h at room temperature in the dark. As a negative control, PE- and APC-conjugated nonspecific mouse IgG1 were substituted for the primary antibodies. The cell suspensions were washed twice, resuspended in 3% FBS/PBS, and analysed with a flow cytometry cell-sorting Vantage cell sorter (Becton Dickinson). The data were analysed with a Mod-Fit 2.0 cell cycle analysis program (Becton Dickinson).
2.5. Real-Time Quantitative PCR (qPCR)
Total RNA was extracted from the cells using an RNeasy Mini Kit (QIAGEN) according to the manufacturer’s directions. The total yield of RNA per extraction was calculated using a NanoVue spectrophotometer (GE Healthcare) to measure the absorbance at 260 nm. A260/A280 ratios of 1.9-2.1 indicated extraction of high-quality RNA. cDNA was synthesised with 2000 ng RNA using MMLV reverse transcriptase (Promega). For qPCR, three separate cDNA samples were used and each measured in triplicate. Target-specific primers (Table 1) were added to each cDNA sample together with Precision MasterMix with ROX and SYBR green (PrimerDesign). The PCR reaction was run by the ABI Prism fast 7500 qPCR system (Advanced Biosystems) under the following cycling conditions: an initial denaturation step of 95°C for 2 minutes followed by 40 cycles of 15 seconds denaturation (95°C) and 1 minute annealing/elongation at 60°C. The relative amount or fold change of the target gene expression was normalized relative to the level of D-glyceraldehyde-3-phosphate dehydrogenase (GAPDH) and relative to a control sample (noninduced cells).
2.6. Western Blot Analysis
The total protein was extracted from the cells with lysis buffer containing protease inhibitors (Roche, UK). The protein concentration was measured by a BCA-200 protein assay kit (Pierce, USA). Equal amounts of proteins were separated by 4-12% sodium dodecyl sulfate/polyacrylamide gel electrophoresis and transferred to a polyvinylidene fluoride (PVDF) membrane. The membrane was blocked in TRIS-buffered saline with Tween (TBST) containing 5% fat-free milk for 2 h and probed with primary antibodies, p-GSK3β (1 : 1000; Cell Signaling), t-GSK3β (1 : 1000; Cell Signaling), active β-catenin (1 : 1000; Cell Signaling), myosin (1 : 500; Sigma), α-SMA (1 : 500; Sigma), desmin (1 : 500, Sigma), and GAPDH (1 : 1000; Cell Signaling) overnight at 4°C and then incubated for 2 h with a horseradish-peroxidase-conjugated anti-mouse IgG antibody or anti-rabbit IgG diluted 1 : 2000 (Cell signaling). Protein bands were visualized on an X-ray film by using an enhanced chemiluminescence system (GE Healthcare, Buckinghamshire, UK). The relative protein expression intensities were quantified by densitometry using Quantity One analysis software.
2.7. Statistical Analysis
Each experiment was performed at least three times, unless otherwise indicated. Data are reported as the (standard deviation) deviation from three independent experiments. The significance of the differences between the experimental and the control groups was determined by using one-way analysis of variance; indicated statistical significance.
3.1. The Proliferation and Differentiation Ability of Three Clones of Human Dental Pulp Stem Cells (DPSCs) (A11, B11, and A32) and Characterization of A32
Dental pulp cells were isolated from pulp tissue of extracted third molars from patients. Three clones of cells that adhered to fibronectin were selected, noted as A11, B11, and A32. The proliferation rate and differentiation potential of the three clones were analysed. A32 demonstrated a high proliferation capacity extending beyond 80PDs, whilst the other two clones (A11 and B11) exhibited less than 36PDs (Figure 1(a)). Compared to A11 and B11 clones, A32 showed the best differentiation capacity into three mesenchymal lineages including osteogenic, adipogenic, and chondrogenic competency (Figure 1(b), B, F, J). The clone A32 was further characterized by flow cytometric analysis, which revealed that A32 was negative for CD34 and CD45. The culture population contained 99.8% CD29-positive cells, 100% CD90-positive cells, 64.4% CD146-positive cells, and 27.2% STRO-1-positive cells (Figure 1(c)).
3.2. The Smooth Muscle Layer-Like Phenotype Generated by the DPSC Clone A32 after SMC Induction
Our previous study proved that the DPSC clone A32 displayed the potential to differentiate into SMCs by induction of CM from bladder SMCs in combination with TGF-β1. In this study, we evaluate whether DPSCs can generate a smooth muscle layer after an extended differentiation duration. Noninduced A32 were found to express α-SMA (Figure 2(a)) and desmin (Figure 2(i)) already, but none of these cells stained positive for myosin (Figure 2(e)). The induced cells from clone A32 formed a monolayer structure and also generated the smooth muscle-like phenotype which were shown to be positive for the SMC markers such as α-SMA (Figures 2(b)–2(d)), myosin (Figures 2(f)–2(h)), and desmin (Figures 2(j)–2(l)) after a longer period of differentiation (up to 20 days).
3.3. The Involvement of Wnt-Mediated GSK3β/β-Catenin in the SMC Differentiation from DPSC Clone A32
The Wnt-GSK3β/β-catenin signaling pathway has been previously reported to play an important role in TGF-β1-induced MSC differentiation. To investigate whether this canonical Wnt signaling is also involved in the bladder SMC differentiation from DPSCs, we evaluated the protein level of phosphorylation GSK3β (p-GSK3β), total GSK3β (t-GSK3β), and active β-catenin in A32 in response to the induction of differentiation medium by western blotting. The protein level of p-GSK3β increased and maintained at the peak through 11 to 14 days (Figures 3(a) and 3(b)), while the protein level of t-GSK3β did not change (Figure 3(a)). As for the expression of active β-catenin, a significant upregulation was detected since day 8 of induction, which reached to the peak expression at day 11 (Figures 3(c) and 3(d)).
3.4. Inhibition of the Wnt-GSK3β/β-Catenin Signaling Pathway Prohibits the SMC-Specific Markers in the DPSC Clone A32 following SMC Induction
To further investigate whether canonical Wnt signaling is required for the SMC differentiation of cell clone A32, we assessed the SMC-specific markers, including α-SMA, myosin, and desmin, in the presence or absence of the Wnt/β-catenin-specific inhibitor, XAV939 and GSK3 inhibitors, SB216763, and LiCl. Pharmacological inhibition of Wnt and GSK3 significantly downregulated the protein expression of p-GSK3β (Figures 4(a) and 4(c)) and active β-catenin (Figures 4(b) and 4(d)) but had no effect on t-GSK3β (Figure 4(a)). The inhibition triggered a further reduction on the expression of SMC-specific markers including α-SMA, myosin, and desmin, at both gene and protein levels (Figures 4(e)–4(l)), compared to those of the non-inhibitor-treated control group of A32 following an SMC induction protocol for 14 days.
3.5. Growth Factor Analysis in SMC Differentiation from A32
The mRNA expression of growth factors, including bFGF, EGF, TGF-β1, HGF, VEGF, and PDGF-BB, in the process of SMC differentiation from DPSC clone A32 were detected by qPCR. The mRNA expression of TGF-β1 was notably increased through 5 to 14 days of differentiation (Figure 5(a)); the HGF and VEGF expression both increased since day 8 of differentiation and reached its peak level at day 14 (Figures 5(b) and 5(c)); the PDGF-BB expression appeared to go up after 11 days of differentiation and reached the highest level at day 14 (Figure 5(d)); the bFGF expression rose to its peak at day 5 but subsequently dropped with 14-day induction (Figure 5(e)); the EGF expression level was increased and maintained through 11 days of differentiation and then downregulated by day 14 (Figure 5(f)).
3.6. Wnt-GSK3β/β-Catenin Signaling Pathway Promotes the DPSC Clone A32 to Differentiate into Mature SMC via Regulating the Expression of Growth Factors
We have demonstrated that Wnt-GSK3β/β-catenin signaling as well as several key growth factors was required in the process of human DPSCs differentiating into human bladder SMCs. To further confirm the involvement of Wnt signaling in the process of SMC differentiation from clone A32 by regulating the expression of growth factors, pharmacological inhibitors of the Wnt pathway were used to evaluate the change of gene expression of several key growth factors at the latter stage of SMC differentiation. After 14 days of differentiation, the gene expression of several growth factors including TGF-β1, HGF, PDGF-BB, HGF, and VEGF markedly upregulated (Figures 6(a)–6(d)), whilst either Wnt/β-catenin-specific inhibitor (XAV939) or GSK3 inhibitors (SB216763 and LiCl) significantly downregulated the expression levels of these growth factors (Figures 6(a)–6(d)), which suggests a promoting effect of Wnt-mediated GSK3β/β-catenin signaling on SMC differentiation of clone A32 by regulating the release of growth factors including TGF-β1, HGF, PDGF-BB, and VEGF.
SMC-based tissue engineering provides a potential therapy for SMC pathology, including cardiovascular diseases, gastrointestinal diseases, urinary incontinence, and bladder dysfunction [29–31]. A reliable cell source of healthy SMCs that can be easily obtained and safely expanded plays a vital role in promoting the progress of smooth muscle tissue engineering. DPSCs have demonstrated the advantages of easy access with the least invasive procedures and little inherent immunogenicity but without any ethical issues, which make them a promising cell population for developing tissue engineering and regenerative medicine . In order to use DPSCs for clinical therapy, in vitro expansion and differentiation ability should be taken into account for the success of clinical applications in tissue engineering. In this study, three clones of DPSCs from two patients were isolated, which were named as A11 and A32 (from patient A) and B11 (from patient B). Investigation of the proliferation rate using PDs as an indicator demonstrated that two of the clones (A11 and B11) showed less than 36PDs and senesced within 85 days, while A32 had a high proliferation rate with over 85PDs that only started senescing after more than 300 days. Compared with the previous studies about the analysis of proliferation rate of MSCs from bone marrow [32–34], stem cell clone A32 displayed a better proliferation capacity, giving this clone a potential advantage. It has been reported that only a small percentage of the overall clonal population of DPSCs, which is less than 5% of the total population, has the ability to differentiate into three mesenchymal lineages, including osteoblast, chondrocytes, and adipocytes [35–37], whilst our data demonstrated that all the three clones (A32, A11, and B11) displayed good ability to differentiate into the osteogenic phenotype, among which A32 was able to differentiate into three different lineages. This result is consistent with data presented by Halleux et al., in which 24 designated clones differentiated into osteogenic lines, while 17/24 and 18/24 clones differentiated into chondrogenic and adipogenic lineages, respectively . The niche of these located clones and the different stages of their development may be accountable for their different growth kinetic profiles and differentiation abilities. Therefore, DPSC clone A32 which has the best proliferation and differentiation ability was selected for further bladder tissue engineering research. We characterized clone A32 by using flow cytometric analysis. The cell populations were positive for a range of mesenchymal stem cell markers including CD29, CD90, CD146, and STRO-1 but negative for hematopoietic stem cell markers, such as CD34 and CD45, which demonstrated that the clone A32 is sourced from the mesenchymal stem cells, not hematopoietic stem cells.
The urinary bladder wall is mainly composed of the Detrusor smooth muscle layer which is made of smooth muscle fibers arranged in spiral, longitudinal, and circular bundles . The Detrusor smooth muscle is mainly responsible for storing urine under low pressure and contraction for voiding, lined by a layer of transitional cells that provide a barrier to absorption. Therefore, it is a crucial step to regenerate the Detrusor smooth muscle layer as a tissue-specific effort for the tissue engineering of the urinary bladder. A previous study in our lab has demonstrated that α-SMA and desmin were already present in noninduced human DPSC clone A32 at a low basal level, which indicates A32 may be more suitable to be induced into SMCs for bladder tissue regeneration. Additionally, this clone has the potential to differentiate into bladder SMC by the CM from cultured bladder SMC in combination with TGF-β1 . In this study, we further proved that A32 could generate a smooth muscle layer-like phenotype after an extended differentiation duration. Further functional analysis would be required using 3D cell culture by seeding the DPSC-derived smooth muscle layer-like structure into synthetic bladder composites and transplanting them into nude rats which underwent removal of half the bladder, in order to evaluate its regeneration capacity of the Detrusor smooth cells in vivo.
Canonical Wnt/β-catenin signaling plays a primary role in the regulation of proliferation and differentiation of stem cells [21, 39]. GSK3 exists as two highly homologous isoforms encoded by distinct genes known as GSK3α and GSK3β. GSK3β originally isolated from muscles as a kinase promotes glycogen assimilation by phosphorylating and inactivating glycogen synthase. It is also involved in the maintenance and plasticity of the skeletal muscle mass, as well as playing an important role in skeletal muscle atrophy in vivo [40, 41]. β-Catenin is a multifunctional protein which is located in both the nucleus and cytoplasm. It leads to myogenic specification and prevents adipogenic differentiation in adult stem cells [42–44]. GSK3β and β-catenin are key factors in the canonical Wnt pathway. Activation of the Wnt signaling promotes the stabilization and accumulation of β-catenin in the cytoplasm by triggering GSK3β phosphorylation which prevents subsequent GSK3β-mediated β-catenin phosphorylation in association with Axin and adenomatous polyposis coli (APC) . The stabilized β-catenin enters the nucleus and induces activation of target genes by binding with members of the T-cell factor (TCF) and lymphoid enhancer factor (LEF) transcription factor family . It has been reported that Wnt-GSK3β/β-catenin contributes to the progressive nature of smooth muscle tissue remodelling  and also acts as an indispensable regulator for myogenesis in embryogenesis and postnatal muscle regeneration , as well as inducing myogenic differentiation in stem cells during muscle regeneration . In this study, we found that Wnt-GSK3β/β-catenin signaling is involved in regulating the process of bladder SMC differentiation from human DPSCs by the SMC induction protocol. To confirm the effect of Wnt/β-catenin signaling and GSK3β on SMC differentiation, two types of pharmacological inhibitors were used. One is XAV939 which is a tankyrase inhibitor for inhibiting Wnt/β-catenin signaling . The others are SB216763 and LiCl which block GSK3 activation [26, 27]. SB216763 is a small molecule that competes with ATP and potently inhibits the activity of α and β isozymes of GSK3. Considering the high degree of sequence similarity, GSK3α and GSK3β share some similar functions. For example, the single loss of either GSK3α or GSK3β in mouse embryonic stem cells (ESC) did not negatively alter Wnt/β-catenin signaling, whereas GSK3α/β double knockout ESCs displayed hyperactivated Wnt/β-catenin signaling, resulting in dramatically skewed cell differentiation . Nevertheless, the two isoforms were shown to have opposite effects on the transcriptional activation of certain transcription factors . GSK3α or GSK3β plays distinct roles in cardiomyocyte differentiation and cardiovascular development in mice [49–51]. Therefore, it appears that GSK3α and GSK3β have both common and nonoverlapping cellular functions, largely depending on the physiological context and the cell type studied. It has been reported GSK3β protein expression in human and mouse muscles was found to be three to four times higher than GSK3α, suggesting that GSK3β, rather than GSK3α, may be the predominant GSK3 isoform in the muscle . Two types of inhibitors significantly suppressed the activation of p-GSK3β as well as active β-catenin and also downregulated the expression of SMC markers (especially myosin which is only expressed in contractile SMCs) at the latter stage of differentiation at both mRNA and protein levels. It suggests that Wnt-GSK3β/β-catenin signaling promotes the human DPSCs to differentiate into bladder SMCs. Further analysis would be required by downregulation of GSK3α or GSK3β to investigate the role of GSK3α in regulating the process of DPSC differentiation into SMC.
Previous studies have identified that a variety of stem cells, including those from the embryo [52, 53], bone morrow , adipose tissue , and dental pulp tissue  can be induced to differentiate into bladder SMCs by using differentiation agents, such as CM from SMC culture in addition to myogenic growth factors, or by indirectly coculturing with the target cells. The CM or indirect coculture system contains cytokine growth factors secreted by the target cells that can be used to induce cell differentiation . Here, we showed that several growth factors, including TGF-β1, HGF, VEGF, PDGF-BB, bFGF, and EGF, play different roles in the process of SMC differentiation. It has been demonstrated that TGF-β1 evokes an important signal that induces expression of vascular SMC markers in a range of nonsmooth muscle precursor cell types, including multipotent embryonic fibroblast , neural crest cells , and MSCs . We here found that the expression of TGF-β1 upregulated during the whole process of SMC differentiation, indicating that TGF-β1 also plays an important role in the bladder SMC differentiation from DPSCs. It has been shown previously that an increased expression of bFGF was associated consistently with SMC proliferation of the neointima formation  and atheromatous lesions  in vivo. Additionally, experimental inhibition of bFGF activation leads to the SMC marker expression in the model of BMMSC differentiation into SMC . As one of the most important mitogenic growth factors, bFGF is known to promote proliferation of marrow cells , which is consistent with our data, because the phenomenon of DPSC proliferation was observed when the expression of bFGF was upregulated during 5 days of SMC differentiation (data not shown). PDGF-BB, which is previously associated with proliferation and differentiation of SMCs [62, 63], was upregulated after 11 days of differentiation in our study, indicating that this growth factor is mainly for promoting bladder SMC differentiation from DPSCs. As for the expression of myogenic growth factors, such as HGF and VEGF, they were both upregulated after 8 days of differentiation, which is consistent with the previous study of BMMSC differentiation into bladder SMCs . The expression levels of the growth factors, including TGF-β1, HGF, VEGF, and PDGF-BB, were all upregulated at 14 days of differentiation, indicating that these growth factors may induce the maturity of SMCs at the latter stage of bladder SMC differentiation from DPSCs. In order to further investigate whether Wnt-GSK3β/β-catenin signaling is involved in the mature process of SMC differentiation, we evaluated the expression of the growth factors above with or without two types of inhibitors at a later stage of SMC differentiation. We noticed that the expression of these growth factors was significantly downregulated with the Wnt/β-catenin inhibitor and GSK3β inhibitors at that stage, indicating that the Wnt-GSK3β/β-catenin signaling promotes the human DPSC differentiation into mature bladder SMCs by regulating the expression of these myogenic growth factors.
As summarized in Figure 7, we hypothesize that the canonical Wnt which is activated by CM from SMCs in combination with TGF-β1 triggers phosphorylation of GSK3β, thus disrupting the APC/Axin/GSK3β/β-catenin complex. The stabilization and accumulation of β-catenin in cytoplasm translocates into the nucleus and enhances the expression of growth factors, including TGF-β1, HGF, VEGF, and PDGF-BB, which subsequently promote the SMC differentiation from DPSCs, for instance, clone A32.
In this study we characterized one of our DPSC clones (clone A32) with the best proliferation and differentiation abilities and demonstrated that the clone A32 possesses a potential to differentiate into a bladder smooth muscle cell layer-like phenotype in vitro using the bladder SMC induction protocol we established before. Additionally, we have found that the Wnt-GSK3β/β-catenin signaling is involved in SMC differentiation. Several growth factors (including TGF-β1, HGF, VEGF, PDGF-BB, and bFGF) regulated by Wnt-GSK3β/β-catenin signaling promote the proliferation/differentiation during the process of bladder SMC differentiation from human DPSCs.
The data used to support the findings of this study are included within the article.
Conflicts of Interest
The authors declare that there is no conflict of interests regarding the publication of this paper.
Wenkai Jiang and Diya Wang contributed equally to the paper.
This research was supported through funding from the National Natural Science Foundation of China (81700951 and 81803186), the Shaanxi Young Talent Lift Project (20170403 and 20180303), the Shaanxi Provincial Natural Science Basic Research Foundation of China (2018JM7123, 2017SF-203, and 2017SF-118), the European Research Council StG grant 243261, and the Royal Society URF award UF051616.
- K. I. Glassberg, “Current issues regarding posterior urethral valves,” The Urologic Clinics of North America, vol. 12, no. 1, pp. 175–185, 1985.
- D. A. Kiddoo, M. C. Carr, S. Dulczak, and D. A. Canning, “Initial management of complex urological disorders: bladder exstrophy,” Urologic Clinics of North America, vol. 31, no. 3, pp. 417–426, 2004.
- W. T. Snodgrass and R. Adams, “Initial urologic management of myelomeningocele,” Urologic Clinics of North America, vol. 31, no. 3, pp. 427–434, 2004.
- W. S. McDougal, “Metabolic complications of urinary intestinal diversion,” The Journal of Urology, vol. 147, no. 5, pp. 1199–1208, 1992.
- T. M. Soergel, M. P. Cain, R. Misseri, T. A. Gardner, M. O. Koch, and R. C. Rink, “Transitional cell carcinoma of the bladder following augmentation cystoplasty for the neuropathic bladder,” The Journal of Urology, vol. 172, no. 4, pp. 1649–1652, 2004.
- J. Y. Lai, C. Y. Yoon, J. J. Yoo, T. Wulf, and A. Atala, “Phenotypic and functional characterization of in vivo tissue engineered smooth muscle from normal and pathological bladders,” The Journal of Urology, vol. 168, no. 4, pp. 1853–1858, 2002.
- C. Danielsson, S. Ruault, A. Basset-Dardare, and P. Frey, “Modified collagen fleece, a scaffold for transplantation of human bladder smooth muscle cells,” Biomaterials, vol. 27, no. 7, pp. 1054–1060, 2006.
- A. Kanematsu, S. Yamamoto, E. Iwai-Kanai et al., “Induction of smooth muscle cell-like phenotype in marrow-derived cells among regenerating urinary bladder smooth muscle cells,” The American Journal of Pathology, vol. 166, no. 2, pp. 565–573, 2005.
- Z. Gong, G. Calkins, E. C. Cheng, D. Krause, and L. E. Niklason, “Influence of culture medium on smooth muscle cell differentiation from human bone marrow-derived mesenchymal stem cells,” Tissue Engineering Part A, vol. 15, no. 2, pp. 319–330, 2009.
- A. I. Caplan, “Mesenchymal stem cells,” Journal of Orthopaedic Research, vol. 9, no. 5, pp. 641–650, 1991.
- M. F. Pittenger, A. M. Mackay, S. C. Beck et al., “Multilineage potential of adult human mesenchymal stem cells,” Science, vol. 284, no. 5411, pp. 143–147, 1999.
- B. M. Strem, K. C. Hicok, M. Zhu et al., “Multipotential differentiation of adipose tissue-derived stem cells,” The Keio Journal of Medicine, vol. 54, no. 3, pp. 132–141, 2005.
- W. A. Robinson, J. E. Kurnick, and B. L. Pike, “Colony growth of human leukemic peripheral blood cells in vitro,” Blood, vol. 38, no. 4, pp. 500–508, 1971.
- K. Bieback, S. Kern, H. Kluter, and H. Eichler, “Critical parameters for the isolation of mesenchymal stem cells from umbilical cord blood,” Stem Cells, vol. 22, no. 4, pp. 625–634, 2004.
- S. Gronthos, M. Mankani, J. Brahim, P. G. Robey, and S. Shi, “Postnatal human dental pulp stem cells (DPSCs) in vitro and in vivo,” Proceedings of the National Academy of Sciences of the United States of America, vol. 97, no. 25, pp. 13625–13630, 2000.
- W. J. Longxing Ni, “Tissue engineering and regenerative medicine, from and beyond the dentistry,” Dentistry, vol. 5, no. 6, 2015.
- H. O. Trowbridge, “Pulp biology: progress during the past 25 years,” Australian Endodontic Journal, vol. 29, no. 1, pp. 5–12, 2003.
- A. Karystinou, F. Dell'Accio, T. B. A. Kurth et al., “Distinct mesenchymal progenitor cell subsets in the adult human synovium,” Rheumatology, vol. 48, no. 9, pp. 1057–1064, 2009.
- R. J. Waddington, S. J. Youde, C. P. Lee, and A. J. Sloan, “Isolation of distinct progenitor stem cell populations from dental pulp,” Cells, Tissues, Organs, vol. 189, no. 1–4, pp. 268–274, 2008.
- B. Song, W. Jiang, A. Alraies et al., “Bladder smooth muscle cells differentiation from dental pulp stem cells: future potential for bladder tissue engineering,” Stem Cells International, vol. 2016, 11 pages, 2016.
- J. L. Teo and M. Kahn, “The Wnt signaling pathway in cellular proliferation and differentiation: a tale of two coactivators,” Advanced Drug Delivery Reviews, vol. 62, no. 12, pp. 1149–1155, 2010.
- R. Gosens, H. Meurs, and M. Schmidt, “The GSK-3/β-catenin-signalling axis in smooth muscle and its relationship with remodelling,” Naunyn-Schmiedeberg's Archives of Pharmacology, vol. 378, no. 2, pp. 185–191, 2008.
- Y. C. Shang, C. Zhang, S. H. Wang et al., “Activated β-catenin induces myogenesis and inhibits adipogenesis in BM-derived mesenchymal stromal cells,” Cytotherapy, vol. 9, no. 7, pp. 667–681, 2007.
- Y. Zhang, H. K. Lin, D. Frimberger, R. B. Epstein, and B. P. Kropp, “Growth of bone marrow stromal cells on small intestinal submucosa: an alternative cell source for tissue engineered bladder,” BJU International, vol. 96, no. 7, pp. 1120–1125, 2005.
- W. Jiang, H. Lv, H. Wang et al., “Activation of the NLRP3/caspase-1 inflammasome in human dental pulp tissue and human dental pulp fibroblasts,” Cell and Tissue Research, vol. 361, no. 2, pp. 541–555, 2015.
- S. Guo, K. Arai, M. F. Stins, D. M. Chuang, and E. H. Lo, “Lithium upregulates vascular endothelial growth factor in brain endothelial cells and astrocytes,” Stroke, vol. 40, no. 2, pp. 652–655, 2009.
- J. Liu, B. Zhu, G. Zhang et al., “Electric signals regulate directional migration of ventral midbrain derived dopaminergic neural progenitor cells via Wnt/GSK3β signaling,” Experimental Neurology, vol. 263, pp. 113–121, 2015.
- S.-M. A. Huang, Y. M. Mishina, S. Liu et al., “Tankyrase inhibition stabilizes axin and antagonizes Wnt signalling,” Nature, vol. 461, no. 7264, pp. 614–620, 2009.
- Z. Gong and L. E. Niklason, “Small-diameter human vessel wall engineered from bone marrow-derived mesenchymal stem cells (hMSCs),” The FASEB Journal, vol. 22, no. 6, pp. 1635–1648, 2008.
- C. R. Chapple, N. I. Osman, A. Mangera et al., “Application of tissue engineering to pelvic organ prolapse and stress urinary incontinence,” LUTS: Low Urinary Tract Symptoms, vol. 7, no. 2, pp. 63–70, 2015.
- A. Atala, S. B. Bauer, S. Soker, J. J. Yoo, and A. B. Retik, “Tissue-engineered autologous bladders for patients needing cystoplasty,” The Lancet, vol. 367, no. 9518, pp. 1241–1246, 2006.
- G. T.-J. Huang, S. Gronthos, and S. Shi, “Mesenchymal stem cells derived from dental tissues vs. those from other sources: their biology and role in regenerative medicine,” Journal of Dental Research, vol. 88, no. 9, pp. 792–806, 2009.
- Y. Tamaki, T. Nakahara, H. Ishikawa, and S. Sato, “In vitro analysis of mesenchymal stem cells derived from human teeth and bone marrow,” Odontology, vol. 101, no. 2, pp. 121–132, 2013.
- D. L. Alge, D. Zhou, L. L. Adams et al., “Donor-matched comparison of dental pulp stem cells and bone marrow-derived mesenchymal stem cells in a rat model,” Journal of Tissue Engineering and Regenerative Medicine, vol. 4, no. 1, pp. 73–81, 2010.
- C. Halleux, V. Sottile, J. A. Gasser, and K. Seuwen, “Multi-lineage potential of human mesenchymal stem cells following clonal expansion,” Journal of Musculoskeletal & Neuronal Interactions, vol. 2, no. 1, pp. 71–76, 2001.
- T. Okamoto, T. Aoyama, T. Nakayama et al., “Clonal heterogeneity in differentiation potential of immortalized human mesenchymal stem cells,” Biochemical and Biophysical Research Communications, vol. 295, no. 2, pp. 354–361, 2002.
- K. C. Russell, D. G. Phinney, M. R. Lacey, B. L. Barrilleaux, K. E. Meyertholen, and K. C. O'Connor, “In vitro high-capacity assay to quantify the clonal heterogeneity in trilineage potential of mesenchymal stem cells reveals a complex hierarchy of lineage commitment,” Stem Cells, vol. 28, no. 4, pp. 788–798, 2010.
- Y. de Jong, J. H. F. M. Pinckaers, R. M. ten Brinck, A. A. B. Lycklama à Nijeholt, and O. M. Dekkers, “Urinating standing versus sitting: position is of influence in men with prostate enlargement. A systematic review and meta-analysis,” PLoS One, vol. 9, no. 7, article e101320, 2014.
- L. Ling, V. Nurcombe, and S. M. Cool, “Wnt signaling controls the fate of mesenchymal stem cells,” Gene, vol. 433, no. 1-2, pp. 1–7, 2009.
- N. Embi, D. B. Rylatt, and P. Cohen, “Glycogen synthase kinase-3 from rabbit skeletal muscle. Separation from cyclic-AMP-dependent protein kinase and phosphorylase kinase,” European Journal of Biochemistry, vol. 107, no. 2, pp. 519–527, 1980.
- J. R. Woodgett and P. Cohen, “Multisite phosphorylation of glycogen synthase. Molecular basis for the substrate specificity of glycogen synthase kinase-3 and casein kinase-II (glycogen synthase kinase-5),” Biochimica et Biophysica Acta (BBA) - Protein Structure and Molecular Enzymology, vol. 788, no. 3, pp. 339–347, 1984.
- A. Polesskaya, P. Seale, and M. A. Rudnicki, “Wnt signaling induces the myogenic specification of resident CD45+ adult stem cells during muscle regeneration,” Cell, vol. 113, no. 7, pp. 841–852, 2003.
- J. De Boer, H. J. Wang, and C. Van Blitterswijk, “Effects of Wnt signaling on proliferation and differentiation of human mesenchymal stem cells,” Tissue Engineering, vol. 10, no. 3-4, pp. 393–401, 2004.
- F. Belema Bedada, A. Technau, H. Ebelt, M. Schulze, and T. Braun, “Activation of myogenic differentiation pathways in adult bone marrow-derived stem cells,” Molecular and Cellular Biology, vol. 25, no. 21, pp. 9509–9519, 2005.
- L. Li, H. Yuan, W. Xie et al., “Dishevelled proteins lead to two signaling pathways. Regulation of LEF-1 and c-Jun N-terminal kinase in mammalian cells,” The Journal of Biological Chemistry, vol. 274, no. 1, pp. 129–134, 1999.
- M. Ikeya and S. Takada, “Wnt signaling from the dorsal neural tube is required for the formation of the medial dermomyotome,” Development, vol. 125, no. 24, pp. 4969–4976, 1998.
- B. W. Doble, S. Patel, G. A. Wood, L. K. Kockeritz, and J. R. Woodgett, “Functional redundancy of GSK-3α and GSK-3β in Wnt/β-catenin signaling shown by using an allelic series of embryonic stem cell lines,” Developmental Cell, vol. 12, no. 6, pp. 957–971, 2007.
- M. H. Liang and D. M. Chuang, “Differential roles of glycogen synthase kinase-3 isoforms in the regulation of transcriptional activation,” The Journal of Biological Chemistry, vol. 281, no. 41, pp. 30479–30484, 2006.
- T. Force and J. R. Woodgett, “Unique and overlapping functions of GSK-3 isoforms in cell differentiation and proliferation and cardiovascular development,” The Journal of Biological Chemistry, vol. 284, no. 15, pp. 9643–9647, 2009.
- R. Kerkela, L. Kockeritz, K. Macaulay et al., “Deletion of GSK-3β in mice leads to hypertrophic cardiomyopathy secondary to cardiomyoblast hyperproliferation,” The Journal of Clinical Investigation, vol. 118, no. 11, pp. 3609–3618, 2008.
- J. Cho, P. Rameshwar, and J. Sadoshima, “Distinct roles of glycogen synthase kinase (GSK)-3α and GSK-3β in mediating cardiomyocyte differentiation in murine bone marrow-derived mesenchymal stem cells,” The Journal of Biological Chemistry, vol. 284, no. 52, pp. 36647–36658, 2009.
- H. Huang, X. Zhao, L. Chen et al., “Differentiation of human embryonic stem cells into smooth muscle cells in adherent monolayer culture,” Biochemical and Biophysical Research Communications, vol. 351, no. 2, pp. 321–327, 2006.
- A. L. Firth and J. X.-J. Yuan, “Human models for smooth muscle cell differentiation. Focus on “a novel in vitro model system for smooth muscle differentiation from human embryonic stem cell-derived mesenchymal cells”,” American Journal of Physiology Cell Physiology, vol. 304, no. 4, pp. C287–C288, 2013.
- H. Tian, S. Bharadwaj, Y. Liu, P. X. Ma, A. Atala, and Y. Zhang, “Differentiation of human bone marrow mesenchymal stem cells into bladder cells: potential for urological tissue engineering,” Tissue Engineering Part A, vol. 16, no. 5, pp. 1769–1779, 2010.
- L. V. Rodriguez, Z. Alfonso, R. Zhang, J. Leung, B. Wu, and L. J. Ignarro, “Clonogenic multipotent stem cells in human adipose tissue differentiate into functional smooth muscle cells,” Proceedings of the National Academy of Sciences of the United States of America, vol. 103, no. 32, pp. 12167–12172, 2006.
- S. Fukuhara, S. Tomita, S. Yamashiro et al., “Direct cell-cell interaction of cardiomyocytes is key for bone marrow stromal cells to go into cardiac lineage in vitro,” The Journal of Thoracic and Cardiovascular Surgery, vol. 125, no. 6, pp. 1470–1479, 2003.
- K. K. Hirschi, S. A. Rohovsky, and P. A. D'Amore, “PDGF, TGF-β, and heterotypic cell-cell interactions mediate endothelial cell-induced recruitment of 10T1/2 cells and their differentiation to a smooth muscle fate,” The Journal of Cell Biology, vol. 141, no. 3, pp. 805–814, 1998.
- N. M. Shah, A. K. Groves, and D. J. Anderson, “Alternative neural crest cell fates are instructively promoted by TGFβ superfamily members,” Cell, vol. 85, no. 3, pp. 331–343, 1996.
- V. Lindner and M. A. Reidy, “Proliferation of smooth muscle cells after vascular injury is inhibited by an antibody against basic fibroblast growth factor,” Proceedings of the National Academy of Sciences of the United States of America, vol. 88, no. 9, pp. 3739–3743, 1991.
- S. E. Hughes, D. Crossman, and P. A. Hall, “Expression of basic and acidic fibroblast growth factors and their receptor in normal and atherosclerotic human arteries,” Cardiovascular Research, vol. 27, no. 7, pp. 1214–1219, 1993.
- D. Orlic, J. Kajstura, S. Chimenti et al., “Mobilized bone marrow cells repair the infarcted heart, improving function and survival,” Proceedings of the National Academy of Sciences of the United States of America, vol. 98, no. 18, pp. 10344–10349, 2001.
- J. Yamashita, H. Itoh, M. Hirashima et al., “Flk1-positive cells derived from embryonic stem cells serve as vascular progenitors,” Nature, vol. 408, no. 6808, pp. 92–96, 2000.
- M. Stehr, R. M. Adam, J. Khoury et al., “Platelet derived growth factor-BB is a potent mitogen for rat ureteral and human bladder smooth muscle cells: dependence on lipid rafts for cell signaling,” The Journal of Urology, vol. 169, no. 3, pp. 1165–1170, 2003.
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