Adipose Derived Stem Cells and Smooth Muscle Cells - Springer Link

Stem Cell Rev and Rep (2009) 5:256–265 DOI 10.1007/s12015-009-9084-y

Adipose Derived Stem Cells and Smooth Muscle Cells: Implications for Regenerative Medicine Jennifer Anne de Villiers & Nicolette Houreld & Heidi Abrahamse

Published online: 11 August 2009 # Humana Press 2009

Abstract The treatment of chronic wounds and other damaged tissues and organs remains a difficult task, in spite of greater adherence to recognised standards of care and a better understanding of pathophysiologic principles. Adipose derived stem cells (ADSCs), with their proliferative and impressive differentiation potential, may be used in the future in autologous cell therapy or grafting to replace damaged tissues. At this point in time, transplanted tissues are often rejected by the body. Autologous grafting would eliminate this problem. ADSCs are able to differentiate into a number of cells in vitro, for example smooth muscle cells (SMCs), when treated with lineage specific factors. SMCs play a key role in physiology and pathology as they form the principle layer of all SMC tissues. Smooth muscle biopsies are often impractical and morbid, and often lead to a low cell harvest. It has also been shown that SMCs derived from a diseased organ can lead to abnormal cells. Therefore, there is a great need for alternative sources of healthy SMCs. The use of ADSCs for cell-based tissue engineering (TE) represents a promising alternative for smooth muscle repair. This review discusses the potential uses of ADSCs and SMCs in regenerative medicine, and the potential of ADSCs to be differentiated into functional SMCs for TE and regenerative cellular therapies to repair diseased organs. Keywords Adipose derived stem cells . Smooth muscle cells . Tissue engineering . Cellular therapy . Differentiation

J. A. de Villiers : N. Houreld : H. Abrahamse (*) Laser Research Group, Faculty of Health Sciences, University of Johannesburg, P.O. Box 17011, Doornfontein 2028, South Africa e-mail: [email protected]

Footnotes and Abbreviations The material in this research paper submitted to stem cell reviews and reports has neither been published, nor is being considered elsewhere for publication ADSCs Adipose derived stem cells BAT Brown adipose tissue BDNF Brain derived neurotrophic factor BMSCs Bone marrow stem cells BMBone marrow-derived smooth muscle progenSMPC itor cells CBFA-1 Core binding factor alpha subunit 1 CNS Central nervous system DMD Duchenne muscular dystrophy ECM Extracellular matrix FM Fusion media GFAP Glial Fibrillary Acidic Protein GFP Green fluorescent protein hADSCs Human ADSCs HIBS Hardening injectable bone substitute ICH Intracerebral haemorrhage IL-5 Interleukin-5 MAP2 Microtubule-associated protein 2 MCAo Middle cerebral artery occlusion MHC Myosin heavy chain MyoD Myogenic determination OECs Olfactory ensheathing cells PLA Processed lipoaspirate PDGF Platelet-derived growth factor BB BB RT-PCR Reverse Transcriptase polymerase chain reaction SIS Small intestine submucosa SMCs Smooth muscle cells SMαSmooth muscle α-actin actin SMIM Smooth muscle induction medium

Stem Cell Rev and Rep (2009) 5:256–265

SMMHC SVF TE TGF β1

Smooth muscle myosin heavy chain Stromal vascular fraction Tissue engineering Transforming growth factor beta 1

Introduction The multidisciplinary science of regenerative medicine and tissue engineering (TE), which combines key elements such as biomaterials, stem cells and bioactive agents (e.g. growth factors), has evolved in parallel with recent biotechnological advances. These future cell-based therapies will benefit from a source of autologous stem cells that are pluripotential and easily accessible [1–4]. Over the past 25 years bone marrow stem cells (BMSCs) have been the focus of extensive research; however the clinical use of these cells has presented various quandaries such as low cell number upon harvest and pain and morbidity to the donor. Like bone marrow, adipose tissue is derived from the mesenchyme and contains an easily isolated supportive stroma containing stem cells, microvascular endothelial cells and smooth muscle cells (Table 1) [5, 6]. Mature adipocytes are easy to remove and separate from the stromal vascular fraction (SVF) by collagenase digestion and centrifugation, and the resulting cell population, termed ADSCs, are maintained in a non-inductive medium [3, 5, 7]. ADSCs are idyllic for cellular therapy applications due to various factors: they can be harvested, multiplied and handled easily, efficiently and non-invasively, they have a pluripotential and proliferative capacity comparable to bone marrow mesenchymal cells, and morbidity to donors is considerably less, requiring only local anaesthesia and a short wound healing time [7]. The most significant feature of adipose tissue as a cell source is that the relative expandability of this tissue allows for large quantities of stem cells to be obtained without difficulty and at minimal risk [8]. ADSCs have multipotential differentiation capacity along the classical mesenchymal lineages of adipogenesis, osteogenesis, chondrogenesis and myogenesis. Nonmesenchymal lineages have also been investigated and the transdifferentiation abilities of ADSCs confirmed, demonstrating that these cells can differentiate into bone, cartilage, fat, heart, nerve, liver and smooth muscle [5, 6, 8, 9] (Fig. 1). Cell therapies involving differentiation of SMCs may offer alternative treatment modalities for diseases that involve SMC pathology such as gastrointestinal disease, urinary incontinence, cardiovascular disease, bladder dysfunction, hypertension, asthma and many more [10, 11].

257 Table 1 Comparison of the characteristics of ADSCs and BM MSCs. ADSCs and BM MSCs have similar CD complements and differentiation potential Surface marker

ADSCs

BM MSCs

References

CD9 CD10 CD13 CD29 CD31 CD34 CD44 CD45 CD49d CD49e CD54 CD55 CD59 CD90 CD105

+ + + + – – + – + + + + + + +

+ + + + – – + – – + + + + + +

[5, [5, [5, [5, [5, [5, [5, [5, [5, [5, [5, [5, [5, [5, [5,

CD106 CD117 CD146

– + +

+ + +

[5, 6] [5] [5, 9]

CD166 STRO-1 Differentiation potential Adipogenesis Oesteogensis Chondrogensis Myogenesis Cardiomyogenesis Neurogenesis Endothelial differentiation Haematopoietic

+ +

+ +

[5, 9] [5, 6]

+ + + + + + + +

+ + + + + + + +

[1, 5, [1, 5, [1, 5, [1, 5, [1, 5, [1, 5, [1, 5, [9]

9] 9] 6, 9] 9] 6, 9] 6, 9] 6, 9] 6, 9] 9] 9] 9] 9] 9] 6, 9] 6, 9]

6, 9] 6, 9] 6, 9] 9] 9] 9] 9]

Adipose Derived Stem Cells in Regenerative Medicine ADSCs have been used for various cellular regeneration and TE studies to date, several of which will be discussed here. Neural Applications Unlike many other tissues, the nervous system has a limited capacity for self repair, with mature nerve cells lacking the capacity to regenerate, and neural stem cells have a limited capability to generate new functional neurons in response to injury [12]. Wang et al. (2007) [13] differentiated ADSCs into a neural cell lineage called olfactory ensheathing cells (OECs), using a co-culture of the stem cells with the OECs on three-dimensional scaffolds. The differentiated cells had

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Fig. 1 The multipotential differentiation capacity of ADSCs into bone, cartilage, fat (adipose tissue), heart (cardiac muscle), nerve (neurons) and smooth muscle

similar morphology and antigenic phenotypes (p75NTR+/ Nestin+/GFAP-) of OECs. Their results indicated that ADSCs had the potential to differentiate into OEC-like cells on three-dimensional scaffolds in vitro. These cells are believed to be important for the repair of damaged CNS (central nervous system), thus the transplantation of OECs is a promising potential therapy for spinal cord injury. ADSCs are known to secrete multiple growth factors and therefore have cytoprotective effects in various injury models [4]. A study was done that investigated the neuroprotective effects of ADSCs in a rat intracerebral haemorrhage (ICH) model and found that ADSC transplantation prompted functional recovery, reduced apoptosis and cerebral inflammation, and reduced brain atrophy and glial proliferation, however, the injected ADSCs had not differentiated into neuronal or glial lineages but the majority rather differentiated into endothelial lineages. In brief, the transplantation of ADSCs in the IHC model reduced chronic brain degradation and acute cerebral inflammation, and promoted long term functional recovery [4].

In addition, Kang et al. (2003) [12] reported that ADSCs have the ability to survive, migrate and improve functional recovery after stroke in rats. Neural differentiation was induced with azacytidine; the cells then exhibited expression of MAP2 (microtubule-associated protein 2) and GFAP (Glial Fibrillary Acidic Protein) and displayed neural morphology. Human ADSCs (hADSCs) were labelled either with LacZ (β-galactosidase) or brain derived neurotrophic factor (BDNF) adenoviruses. hADSCs labelled with LacZ adenovirus were injected into the lateral ventricle of the rat brain, and showed that the transplanted cells migrated to different parts of the brain, there was an increase in migration to the injured cortex in ischemic brain injury by middle cerebral artery occlusion (MCAo) and there were improved functional deficits. There was expression of MAP2 and GFAP in some of the transplanted cells. Intracerebral grafting of BDNF-transduced hADSCs drastically improved motor recovery of functional deficits in MCAo rats. This study demonstrates that genetically engineered hADSCs can express biologically active gene


products and as a result can function as useful vehicles for curative gene transfer to the brain. Cardiac Applications Congenital and acquired heart disease has emerged as a leading cause of mortality and morbidity world-wide [5, 14]. At present, the synthetic materials or bioprosthetic replacement devices for cardiovascular surgery are flawed and patients are subjected to various ongoing risks such as thrombosis, a need for re-operations and limited durability due to a lack of growth in children and young adults. Cellular therapy has transpired as a potential new therapeutic option to treat cardiovascular disease [15]. ADSCs are able to differentiate into cardiomyocytes, hence these autologous adult stem cells are emerging as a new source of cells for cardiovascular repair [16]. Yamada et al. (2006) [17] reported that CD29+ murine ADSCs from brown adipose tissue (BAT) could differentiate into cardiomyocytes with high efficiency. These findings were based on morphology, electrophysical parameters, and molecular and protein expression. They transplanted the CD29+ BAT-derived cells into the infarct border zone. Immunohistochemistry revealed that implanted cells expressed markers found in SMCs, endothelial cells and cardiomyocytes. A cardiac function test, measured by echocardiology, revealed that there was improved ventricular function and reduction of the infarction area due to the transplantation of the CD29+ BATderived cells. A recent article from America reports porcine bone marrow stem cells (BMSCs), as well as ADSCs engrafted in the infarct region, improved cardiac function and perfusion after intracoronary cell transplantation [18]. Myocardial infarction was induced and two million cultured autologous cells were intracoronary injected. Relative and absolute perfusion defects significantly decreased after 28± 3 days after BMSCs and ADSCs administration compared to carrier administration. There was a significant increase in left ventricular function, the thickness of the ventricular wall in the infarction area, and improved vascular density of the border zone after the administration of stem cells as compared to the carrier administration. Co-localisation of the grafted cells with von Willebrand factor and alphasmooth muscle actin was observed, with the incorporation into newly formed blood vessels. Musculoskeletal Applications Several million people are affected by bone diseases, such as osteoporosis and osteopenia, throughout the world. Ongoing studies for repairing bone defects are done in gene and cell therapies and pharmacology [19].

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In a recent article Elabd et al. (2007) [19] showed that multipotent human ADSCs can differentiate into osteocytelike cells, and can develop into mineralised woven bone after 4 weeks when loaded on a hardening injectable bone substitute (HIBS) biomaterial and injected subcutaneously into nude mice. The ADSCs were induced into oesteocytes in the presence of α-MEM (Eagle’s minimum essential medium) containing a hormonal cocktail. Oesteoblastic differentiation was established through staining with Alizarin Red (which indicates the formation of mineralised extracellular matrix) and expression of alkaline phosphatase, CBFA1 (core binding factor alpha subunit 1) and osteonectin. Six week old nude mice where anaesthetised and injected with the HIBS, with or without ADSCs, and after 4 weeks the HIBS/cell implants exhibited a hard consistency compared to the cell free/HIBS implants. Histological analysis showed that all implants were fully colonised with woven bone within the ceramic in the intergranular spaces, and several cuboidal-shaped osteoblasts were present on the surface of the biphasic calcium phosphate particles associated with numerous osteocytes. Many vessels that could support bone formation as well as multinucleated cells were extensively distributed. Duchenne muscular dystrophy (DMD) is an X-linked genetic disorder that is characterised by progressive muscle weakness and degeneration. Cell therapy is being pursued as a possible treatment modality for the repair of defective muscle in DMD [20]. Vieira et al. (2008) [20] showed that ADSCs participate in myotube formation resulting in the restoration of dystrophin. To differentiate hADSC’s into myogenic lineage stem cells were grown in growth media (DMEM-HG, 10% fetal bovine serum) supplemented with 0,1 µM dexamethasone, 50 µM hydrocortisone and 5% horse serum for 45 days. Immunofluorescence studies showed expression of alpha-actinin in the differentiated culture, and RT-PCR revealed expression of MyoD (myogenic determination), telethonin and dystrophin. Western blot confirmed the presence of dystrophin in early passage, high cell density cultures. Two different types of cocultures were tested. In the first equal amounts of GFP (green fluorescent protein)-negative myoblasts stained with DAPI and GFP-positive ADSCs were co-cultured. In the second culture the DMD myotubes were stained with DAPI and GFP-positive ADSCs were added at a ratio to 3:1. The cultures were then subjected to fusion media (FM) that induces myoblasts to coalesce and form multinucleated structures. The conclusion for the co-culture experiment was that ADSCs participate in the generation of human myotubes through cellular fusion as all GFP + syncytia presented at least one DAPI stained nuclei, and expressed dystrophin. ADSCs were plated with DMD myotubes, and it was revealed that ADSCs are able to fuse with DMD myotubes and restore dystrophin. Dystrophin expression

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was evaluated by RT-PCR and it was found that ADSCs, when differentiated into muscle cells, can express dystrophin at a level equal to that of normal myoblasts.

Smooth Muscle Cells Sheets of SMCs are contained in the walls of various organs and tubes in the body, for instance the blood vessels, intestines, bladder, stomach, airways, uterus, and the clitoral and penile cavernosal sinuses. Smooth muscle plays an active role in a multitude of physiological and pathological processes. These cells are involved in basic physiological functions such as reproduction, transport of nutrients and oxygen, and breathing [21–23]. SMCs contract to regulate the luminal diameter of hollow organs in normal tissues, and in disease states they serve as effectors of fibrosis, inflammation and muscle mass accumulation [24]. A great variation in size, morphology and biochemistry is displayed by SMCs. This variation is accredited to differences in the embryological origin and the state of differentiation of the cells. Associated with different functional properties, SMCs come in a variety of phenotypes, and are therefore distinctive among myogenic lineages in that they retain a multifunctional capacity for contraction, proliferation, migration, secretion of growth factors and synthesis of extracellular matrix (ECM) components. Unlike striated muscle types, SMCs can differentiate from a proliferative (synthetic) phenotype to a contractile phenotype (Fig. 2). This reversible process between a contractile and synthetic phenotype is called modulation and has been described as being a characteristic response of mature SMCs that are

Fig. 2 Phenotypic variation between SMCs. SMCs may undergo a reversible process called modulation between the contractile and synthetic phenotypes


derived from all visceral and vascular organs investigated and studied [22–27]. The reversion of primary cultured SMCs back to the contractile state is termed maturation and is marked by increased contractile apparatus-associated protein content and myofilament, decreased profusion of synthetic organelles, and reacquisition of archetypal pharmacological responsiveness [18]. Regardless of the tissue of origin, the primary role of SMCs is contraction and relaxation. In the body, the process of SMC contraction is regulated primarily by receptor and mechanical or stretch activation of actin, myosin and various other contractile proteins, and the expression of a large number of regulatory proteins that are present in precisely controlled concentrations [10, 21, 28]. A change in membrane potential brought about by the activation of stretch-dependent ion channels or by the firing of action potentials in the plasma membrane can trigger contraction [21]. Relaxation of SMCs occurs in one of two ways, active relaxation induced by activation of cyclic nucleotide-dependent signalling pathways in the continued presence of the contractile agent, or passive relaxation brought about by the removal of the contractile agent [29]. Some of the best characterised markers for contractile SMC include smooth muscle α-actin (SMα-actin), caldesmon, SM22, calponin, smooth muscle myosin heavy chain (SM-MHC) and smoothelin. SMα-actin is an early marker of developing SMCs, whereas the other markers are highly restricted to differentiated smooth muscle, particularly SMMHC and smoothelin. The latter two markers can be qualified as most SMC specific as they are exclusively expressed in contractile, and thus functional, SMCs. Smoothelin is the only marker that differentiates between SMCs and myofibroblasts [10, 22, 23, 26, 27, 30].


Smooth Muscle Cells in Regenerative Medicine As mentioned earlier, SMCs play a role in a variety of physiological and pathological conditions because they constitute the principal layer of all SMC tissues. They are known to play a critical role in a large number of major human diseases, such as arteriosclerosis, asthma, hypertension and cancer [31]. The following section will be dedicated to TE research done to date on several of the more prominent diseases involving SMC. Arteriosclerosis Arteriosclerosis encompasses spontaneous arteriosclerosis, autologous arterial or venous graft arteriosclerosis, restenosis after percutaneous transluminal coronary angioplasty and transplant arteriosclerosis. In all types of arteriosclerosis, accumulation of vascular SMC in the intima is a key event [32]. The SMCs of the arterial media play a predominant role in structural and functional alterations of the arterial wall. The switch from the contractile to the synthetic phenotype appears to be an early critical event in arteriosclerosis. This phenotypic modulation permits SMCs to migrate into the intima, proliferate and secrete extracellular matrix components [33].

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and caldesmon. When embedded in fibrin hydrogels, the BM-SMPCs contracted the matrix and displayed receptorand non-receptor-mediated contractility, which indicated that these cells can generate force in response to vasoreactive agonists. Tissue-engineered blood vessels were prepared from BM-SMPC and BM-derived endothelial cells and implanted into the jugular vein of lambs. Five weeks post-implantation, the grafted tissues showed a confluent endothelial layer over the medial layer in which BM-SMPC were aligned circumferentially and synthesised significant amounts of collagen, as well as high amounts of elastin organised in a fibrillar network very similar to native vessels. In conclusion, BM-SMPC may be of use in studying SMC differentiation and have potential for development of cell therapies for the treatment of cardiovascular disease. Asthma Chronic persistent asthma is characterised by poorly reversible airway obstruction, and shows marked inflammatory and architectural changes associated with airway wall thickening. Increased airway smooth muscle content is believed to be one of the principle contributors to airway wall thickening, and occurs as a result of hyperplastic and/ or hypertrophic growth [38].

Cardiovascular Applications of SMCs Respiratory System Applications of SMCs Cardiovascular disease is the leading cause of mortality around the world increasing the demand for small diameter blood vessels. Current treatments include stents, vascular grafts (currently the golden standard) and angioplasty [34, 35]. Venous grafts suffer from several major disadvantages: (i) pain and discomfort at the donor site; (ii) limited availability, especially for repeat grafting procedures; (iii) limited replicative capacity of cells from older donors, and (iv) high 10-year failure rate [35]. The aim of small diameter blood vessel TE is to create autologous replacement vessels for use in vascular replacement surgery, amongst others [36]. These conduits need to closely resemble the native vessel, not only in dimension but also in physiological responsiveness, cellular constituents, and mechanical properties as well as in its relaxing and antithrombotic factors released at the cellular level [37]. Liu et al. (2007) [34] developed a novel method for isolating SMCs from ovine bone marrow using a tissuespecific promoter and fluorescence-activated cell sorting, for potential use in cardiovascular tissue regeneration applications. When compared to vascular SMCs, bone marrow-derived smooth muscle progenitor cells (BMSMPC) showed higher proliferation, exhibited similar morphology and expressed several SMC markers such as myosin heavy chain, smoothelin, calponin, α-actin, SM22

Airway smooth muscle is present in the bronchial tree of most vertebrates. It encircles the entire airway beneath the level of the main bronchus, in a generally circular orientation, except at high lung volumes. These cells are present in the central and peripheral airways, more transverse in central airways and somewhat more prominent in the peripheral airways. Airway SMCs are arranged in a helical or geodesic pattern which is more apparent when the lung is fully inflated and in the peripheral airways [39]. Chakir et al. (2000) [40] evaluated the feasibility of an engineered human bronchial mucosa as a model to study cellular interactions in asthma. Human bronchial fibroblasts from normal and asthmatic donors were incorporated into collagen gel, and bronchial epithelial cells were seeded over this gel and cultured in an air-liquid interface in the presence or absence of T lymphocytes. Biopsy specimens from this engineered mucosa were taken for ultrastructural and structural analysis; T lymphocytes were harvested and IL-5 was localised. The histologic analysis demonstrated that engineered mucosa with normal bronchial cells presented a pseudostratified ciliated epithelium with the presence of mucus secretory cells. These features were confirmed with electron microscopy, and were comparable with those observed in normal bronchial tissues. However,

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in engineered mucosa from asthmatic subjects, the tissue structure was disorganised, principally the epithelial cell arrangement. The percentage of IL-5+ lymphocytes was significantly (P=.03) higher in engineered bronchial mucosa from asthmatic subjects (87%±2%) compared with mucosa from normal volunteers (2%±0.3%). In conclusion, the authors claim that using TE they produced an in vitro model of bronchial mucosa from normal and asthmatic subjects, which could be used in future to better comprehend the key mechanisms involved in the inflammation and repair of respiratory tissue. Bladder Disease A variety of injuries can lead to the damage or loss of the bladder, requiring the eventual replacement or repair of the organ [41]. Surgical repair of the urinary bladder using tissue augmentation is frequently required in the management of bladders damaged by spinal cord injuries, trauma, malignancies and inherited pediatric disorders such as bladder exstrophy and spinal bifida. Traditionally the surgery is performed using a pedicle flap of intestine to reconstruct a urinary reservoir. However, these reconstructive procedures are technically difficult and allied with numerous sequela including urinary calculi, intestinal adhesions, chronic infections, and secondary malignancies. The fields of materials science and TE have advanced, and in so doing interest has grown in the use of biomaterials synthetically designed for urinary storage to avoid the problems stated above. For these purposes bladder biomaterials have been used alone as well as with cell seeding. TE critics have pointed out that the conventional bladder cell harvest procedures are surgically invasive, and the prolonged cell expansion times are subject to contamination, expensive and are consequently unrealistic for everyday clinical use [42]. TE aims to regenerate urological organs and structures to a point where full physiological functions are restored. Stem cells can be differentiated into a variety of cells and are of great potential use in TE [43]. A recent article by Baumert et al. (2007) [44] describes a model using the omentum as an in vivo bioreactor for a previously seeded scaffold to prevent ischaemic fibrosis during tissue maturation. Bladder biopsies of five female pigs were taken, and both smooth muscle and urothelial cell cultures were made. These cultures were used to seed a sphere-shaped small intestine submucosa (SIS) matrix, which was then transferred into the omentum after 3 weeks of growth. The grafts were harvested 3 weeks later and immunohistochemical, histologic and functional studies were done. A highly vascularised tissue engineered construct was obtained which contracted in response to acetylcholine stimulation. An average wall thickness of 4 mm was seen. Immunostaining and histologic analysis


confirmed a multilayer urothelium on the luminal aspect and deeper facicles organised tissue composed of differentiated SMCs and mature fibroblasts with no evidence of necrosis or inflammation present. Large- and small-diameter blood vessels were clearly identified histologically. This study demonstrated that the omentum allowed in vivo maturation of seeded scaffolds with the development of a dense vascularisation that is anticipated to avoid a loss of contractility and prevent fibrosis. The authors concluded that this in vivo maturation into the omentum could be the first step before in situ implantation of the construct. Atala et al. (2006) [41] explored an alterative approach to cystoplasty as a treatment of patients with end-stage bladder disease using autologous engineered bladder tissues. Seven patients with myelomeningocele, with highpressure or poorly compliant bladders, were identified as candidates for cystoplasty and bladder biopsies were obtained. Muscle and urothelial cells were grown in culture, then seeded on a biodegradable bladder-shaped scaffold (collagen or collagen-polyglycolic acid). Seven weeks after the biopsy, the autologous engineered bladder constructs were used for reconstruction and implanted with or without an omental wrap. Cytograms, ultrasounds, bladder biopsies, serial urodynamics and serum analyses were then done. Follow up analyses were done in a mean range of 46 months. Post-operatively, the mean bladder leak point pressure decreased with capacity, and the compliance and volume increase was greatest in the composite engineered bladders with an omental wrap. After surgery, bowel function returned promptly. Metabolic consequences were not noted, no urinary calculi formed, renal function was preserved and mucus production was normal. An adequate structural architecture and phenotype was displayed by the engineered bladder biopsies. This research showed that engineered bladder tissues, created with autologous cells seeded on collagen-polyglycolic acid scaffolds, wrapped in omentum after implantation, can be used as an alternative to cystoplasty in patients with end stage bladder disease. Jack et al. (2009) [42] cultured hADSCs in smooth muscle inductive media (SMIM) (consisting of MCDB131 media (Sigma) supplemented with 1% FBS, 100 U/mL heparin) for 6 weeks at 37°C in 5% CO2. Successful differentiation was assessed by the presence of SMCspecific markers SMα-actin, caldesmon, and myosin heavy chain (MHC) as per immunofluorescence. Differentiated cells were then seeded into synthetic 85:15 poly-lacticglycolic acid (PLGA) bladder dome composites to tissue engineer bladder smooth muscle. Cell-seeded bladders expressed SMα-actin, MHC, calponinin, and caldesmon as determined by immunoflourescence and RT-PCR. Nude rats (n ¼ 45) underwent removal of half their bladder and repair using: (i) augmentation with the adipose stem cellseeded composites, (ii) augmentation with a matched


acellular composite, or (iii) suture closure. Twelve weeks post-implantation the animals were followed, and bladders were explanted serially. Results showed smooth muscle phenotype was maintained by the differentiated SMCs in the seeded scaffolds after 2 weeks in vitro in SMIM. RT-PCR demonstrated mRNA signal expression of SMα-actin, caldesmon, SM22, MHC, and smoothelin from the differentiated seeded constructs. MHC and smoothelin mRNA, both of which are specific to smooth muscle, were specific to the differentiated constructs and absent in the undifferentiated ADSC control constructs. Throughout the 12 weeks bladder compliance and capacity were maintained in the cell-seeded group, but deteriorated in the acellular scaffold group sequentially with time. By week 12, control animals repaired with sutures regained their baseline bladder capacities demonstrating a long-term limitation of this model. Histological analysis of explanted materials demonstrated viable ADSCs and an increase in smooth muscle mass in the cellseeded scaffolds as time ensued. Tissue bath stimulation showed smooth muscle contraction of the seeded implants but not the acellular implants after 12 weeks in vivo. This study demonstrates the feasibility and short term physical properties of ADSC-engineered bladder.

ADSCs Differentiated into SMCs There is a variety of diseases associated with SMC pathology. The use of stem cells for cell based TE provides a promising possible alternate to current treatment strategies for smooth muscle repair or regeneration. A limitation to date has been a reliable source of SMCs for these applications [10]. Rodriguez et al. (2006) [10] differentiated processed lipoaspirate (PLA) cells into functional (therefore contractile) SMCs. To induce differentiation, PLA cells were cultured in smooth muscle induction media (consisting of MCDB131 media (Sigma) supplemented with 1% FBS at varying concentrations of heparin), which brought about genetic expression, at a transcriptional and translation level, of smooth muscle alpha actin (SMαa), SM22, caldesmon, calponin, smoothelin and myosin heavy chain (MHC). Upon induction, PLA cells assumed typical SMC morphology. To examine functionality of the differentiated SMCs the cells were subjected to carbachol and atrophine. After 1 min exposure to 10−5M carbachol the cells began to contract, remaining in this state for up to 10 min, which shows similar contractile function as that of SMCs in vivo. The muscarinic antagonist atrophine (10−4M) was shown to block the effects mediated by carbachol. The cells exhibited SMC characteristics both phenotypically and functionally. Hence, ADSCs have the potential to differentiate into functional SMCs that

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could be used in therapeutic applications as a source of healthy SMCs. In a recent article by Kima et al. (2008) [45] it was demonstrated that the treatment of hADSCs with Angiotensin II (Ang II) increased the expression of smooth muscles specific genes, including SMα-actin, calponin, h-caldesmon, and SM-MHC, and also elicited the secretion of transforming growth factor β1 (TGF β1) and delayed phosphorylation of Smad2. The Ang II-induced expression of SMα-actin and delayed phosphorylation of Smad2 were blocked by pretreatment of the cells with a TGF β1 type I receptor kinase inhibitor, SB-431542, adenoviral expression of Smad7, and small interference RNA-mediated depletion of endogenous Smad2. In addition, the Ang II-induced TGF β1 secretion, SMα-actin expression, and delayed phosphorylation of Smad2 in hADSCs were abrogated by the MEK inhibitor U0126, suggesting a pivotal role of MEK/ERK pathway in the Ang II-induced activation of the TGF β1-Smad2 signaling pathway. The smooth muscle-like cells which were differentiated from hADSCs by Ang II treatment exhibited contraction in response to 60 mM KCl. These results suggest that Ang II induces differentiation of hADSCs into contractile smooth muscle-like cells through ERK-dependent activation of the autocrine TGF β1-Smad2 crosstalk pathway. Yang et al. (2008) [46] were interested in studying the feasibility of hADSCs induced into smooth muscle cells in vitro as seeding cells in vascular TE. When almost confluent, ADSCs were subcultured and platelet-derived growth factor (PDGF)-BB (50 ng/mL) and TGF-beta1 (5 ng/mL) were added to enhance the SMC phenotype. Cells were cultured under the inducing medium for 14 days. Morphologically induced cells exhibited “Hill and Valley” morphology, while the uninduced cells were similar to the passage zero (P0) ADSCs which had typical fibroblast-like morphology. Immunofluorescence results show that the induced cells expressed SMC-specific markers including SMα-actin, SMMHC and calponin. RT-PCR results revealed that the induced cells expressed SMα-actin, SM-MHC, calponin and SM-22α. These results indicate that hADSCs can be induced to express vascular smooth muscle markers, and they are a new potential source of vascular TE.

Conclusion Various complications are often seen in tissue and organ replacement surgeries because the human immune system identifies and antagonises extraneous organs and artificial prosthesis. Suppression of a patient’s immune system would solve this issue; however many other complications would arise. Thus, the aim of TE is to establish autologous tissue and organ transplants, i.e. from the patient’s own cells. This will diminish graft versus host reactions [47].

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Adult stem cells such as ADSCs offer medical researchers and scientists the chance to produce autologous de novo tissues ex vivo with few ethical dilemmas. ADSCs can be isolated from a patient’s own adipose tissue, cultured on specific matrices or scaffolds with lineage-specific growth factors that allow differentiation into a required tissue type, and ultimately the differentiated tissue could be grafted back into the same patient [48]. Studies have shown that ADSCs display similar immunoregulatory properties as human BMSC’s by inhibiting the proliferation and cytokine secretion of human primary T cells in response to mitogens and allogenic T cells [49]. ADSCs have emerged as a new and promising type of stem cell with two distinct advantages over previously used adult stem cells: (i) easy and repeatable access allows harvesting of high amounts of adipose tissue, and (ii) ADSCs have an increased potential to proliferate and expand themselves in culture [16]. Low level laser irradiation can positively affect human ADSCs by increasing cellular proliferation, viability and protein expression [50, 51] which highlights yet another potential advantage for the use of these cells in TE applications as the initial cell number could be increased before commencing differentiation leading to a higher yield of differentiated cells. Smooth muscle is an active component of the cardiovascular, reproductive, urinary and intestinal systems, and has been the subject of intense research in the field of regenerative medicine. A restriction to date has been a reliable source of healthy SMCs, as biopsies normally lead to low cell harvest that needs to be expanded at length before therapeutic use [10]. In addition, previous research has shown that SMCs that are obtained from diseased tissue can differ phenotypically and functionally from normal healthy SMCs, which consequently restricts their use [52, 53]. The findings of Rodriguez et al. (2006) [10], Jack et al. (2009) [42], Kima et al. (2008) [45] and Yang et al. (2008) [46] describe a source of cells to use for SMC applications, as the results show that ADSCs have the potential to differentiate into functional SMCs and consequently may prove a useful source of autologous cells for reconstruction of diseased human organs and tissues containing smooth muscle.

Author disclosure statement authors.

There is no conflict of interest for all

References 1. Gimble, J. M., Katz, A. J., & Bunnell, B. A. (2007). Adipose-derived stem cells for regenerative medicine. Circulation Research, 100, 1249–1260. 2. Sandor, G. K. B., & Suuronen, R. (2008). Combining adiposederived stem cells, resorbable scaffolds and growth factors: an overview of tissue engineering. JCDA, 74(2), 167–170.

3. Zuk, P. A., Zhu, M., Mizuno, H., Huang, J., Furtell, J. W., Katz, A. J., et al. (2001). Multilineage cells from human adipose tissue: implications for cell-based therapies. Tissue Engineering, 7(2), 211–228. 4. Kim, J. M., Lee, S., Chu, K., Jung, K., Song, E., Kim, S., et al. (2007). Systemic transplantation of human adipose stem cells attenuated cerebral inflammation and degeneration in a hemorrhagic stroke model. Brain Research, 1183, 43–50. 5. Strem, B. M., Hicok, K. C., Zhu, M., Wulur, I., Alfonso, Z., Schreiber, R. E., et al. (2005). Multipotential differentiation of adipose tissue-derived stem cells. Keio Journal of Medicine, 54 (3), 132–141. 6. Zuk, P. A., Zhu, M., Ashjian, P., De Ugarte, D. A., Huang, J. I., Mizuno, H., et al. (2002). Human adipose tissue is a source of multipotent stem cells. Molecular Biology of the Cell, 13, 4279–4295. 7. Ogawa, R. (2006). The importance of adipose-derived stem cells and vascularised tissue regeneration in the field of tissue transplantation. Current Stem Cell Research and Therapy, 1, 13–20. 8. Fraser, J. K., Wulur, I., Alfonso, Z., & Hedrick, M. H. (2006). Fat tissue: an underappreciated source of stem cells for biotechnology. TRENDS in Biotechnology, 24(4), 150–154. 9. Schäffler, A., & Büchler, C. (2008). Concise review: adipose tissue-derived stromal cells—basic and clinical implications for novel cell-based therapies. Stem Cells, 25, 818–827. 10. Rodriguez, L. V., Alfonso, Z., Zhang, R., Leung, J., Wu, B., & Ignarro, L. J. (2006). Clonogenic multipotent stem cells in human adipose tissue differentiate into functional smooth muscle cells. Proceeding of the National Academy of Sciences, 103(32), 12167– 12172. 11. Wong, J. Z., Woodcock-Mitchell, J., Mitchell, J., Rippetoe, P., White, S., Absher, M., et al. (1998). Smooth muscle actin and myosin expression in cultured airway smooth muscle cells. AJPLung Cellular & Molecular Physiology, 274, 786–792. 12. Kang, S. K., Lee, D. H., Bae, Y. C., Kim, H. K., Baik, S. Y., & Jung, J. S. (2003). Improvement of neurological deficits by intracerebral transplantation of human adipose tissue-derived stromal cells after cerebral ischemia in rats. Experimental Neurology, 183, 355–366. 13. Wang, B., Han, J., Gao, Y., Xiao, Z., Chen, B., Wang, X., et al. (2007). The differentiation of rat adipose-derived stem cells into OEC-like cells on collagen scaffolds by co-culturing with OECs. Neuroscience Letters, 421, 191–196. 14. Wu, K., Liu, Y. L., Cui, B., & Han, Z. (2006). Application of stem cells for cardiovascular grafts tissue engineering. Transplant Immunology, 16, 1–7. 15. Bai, X., Pinkernell, K., Song, Y., Nabzdyk, C., Reisser, J., & Alt, E. (2006). Genetically selected stem cells from human adipose tissue express cardiac markers. Biochemical and Biophysical Research Communications, 353, 665–671. 16. Sanz-Ruiz, R., Fernandez Santos, M. E., Dominguez Munoa, M., Ludwig Martin, I., Parma, R., Sanchez Fernandez, P. L., et al. (2008). Adipose tissue-derived stem cells: the friendly side of a classic cardiovascular foe. Journal of Cardiovascular Translational Research, 1(1), 55–63. 17. Yamada, Y., Wang, X., Yokayama, S., Fukuda, N., & Takakura, N. (2006). Cardiac progenitor cells in brown adipose tissue repaired damaged myocardium. Biochemical and Biophysical Research Communications, 342, 662–670. 18. Valina, C., Pinkernell, K., Song, Y., Bai, X., Sadat, S., Campeau, R. J., et al. (2007). Intracoronary administration of autologous adipose tissue-derived stem cells improves left ventricular function, perfusion, and remodelling after acute myocardial infarction. European Heart Journal, 28, 2667–2677. 19. Elabd, C., Chiellini, C., Massoudi, A., Cochet, O., Zaragosi, L., Trojani, C., et al. (2007). Human adipose tissue-derived multipotent stem cells differentiate in vitro and in vivo into osteocyte-like cells.


20.

21. 22.

23.

24.

25.

26. 27.

28.

29.

30.

31.

32. 33.

34.

35.

36.

Biochemical and Biophysical Research Communications, 361, 342– 348. Vieira, N. M., Brandalise, V., Zucconi, E., Jazedje, T., Secco, M., Nunes, V. A., et al. (2008). Human multipotent adipose derived stem cells restore dystrophin expression of Duchenne skeletal muscle cells in vitro. Biology of the Cell, 100(4), 231–241. Webb, R. C. (2003). Smooth muscle contraction and relaxation. Advances in Physiology Education, 27(4), 201–206. van Eys, G. J. J. M., Voller, M. C. W., Timmer, E. D. J., Wehrens, X. H. T., Small, J. V., Schalken, J. A., et al. (1997). Smoothelin expression characteristics development of a smooth muscle cell in vitro system and identification of a vascular varient. Cell Structure and Function, 22, 65–72. Suzuki, T., Nagai, R., & Yazaki, Y. (1998). Mechanisms of transcriptional regulation of gene expression in smooth muscle cells. Circulation Research, 82, 1238–1242. Halayko, A. J., Camoretti-Mercado, B., Forsythe, S. M., Vieria, J. E., Mitchell, R. W., Wylam, M. E., et al. (1999). Divergent differentiation paths in airway smooth muscle culture: induction of functionally contractile myocytes. American Physiology Society, 276, 197–206. Hollrigel, A., Puz, S., Al-Dubai, H., Kim, J. U., Capetanaki, Y., & Weitzer, G. (2002). Amino-terminally truncated desmin rescues fusion of des-/- myoblasts but negatively affects cardioyogenesis and smooth muscle development. FEBS Letters, 523, 229–233. van Eys, G. J., Niessen, P. M., & Rensen, S. S. (2007). Smoothelin in vascular smooth muscle cells. TCM, 17(1), 26–30. Halayko, A. J., & Solway, J. (2001). Molecular mechanisms of phenotypic plasticity in smooth muscle cells. Journal of Applied Physiology, 90, 358–368. Ma, X., Wang, Y., & Stephens, N. L. (1998). Serum deprivation induces a unique hypercontractile phenotype of cultured smooth muscle cells. AJP-Cell Physiology, 274, 1206–1214. Woodrum, D. A., & Brophy, C. M. (2001). The paradox of smooth muscle physiology. Molecular and Cellular Endocrinology, 177, 135–143. Owens, G. K., Kumar, M. S., & Wamhoff, B. R. (2004). Molecular regulation of vascular smooth muscle cell differentiation in development and disease. Physiological Reviews, 84, 767–801. Sinha, S., Wamhoff, B. R., Hoofnagle, M. H., Thomas, J., Neppi, R. L., Deering, T., et al. (2008). Assessment of contractility of purified smooth muscle cells derived from embryonic stem cells. Stem Cells, 24, 1678–1688. Mayr, M., & Xu, Q. (2001). Smooth muscle cell apoptosis in arteriosclerosis. Experimental Gerontology, 36, 969–987. Tukaj, C., Bohdanowicz, J., & Kubasik-Juraniec, J. (2004). The growth and differentiation of aortal smooth muscle cells with microtubule reorganisation—an in vitro study. Folia Morphologiica, 63(1), 51–57. Liu, J. Y., Swartz, D. D., Peng, H. F., Gugino, S. F., Russell, J. A., & Andreadis, S. T. (2007). Functional tissue-engineered blood vessels from bone marrow progenitor cells. Circulation Research, 75, 618–628. Wong, J. W., Velasco, A., Rajagopalan, P., & Pham, Q. (2003). Directed movement of vascular smooth muscle cells on gradientcompliant hydrogels. Langmuir, 19, 1908–1913. Krenning, G., Moonen, J. R. A. J., van Luyn, M. J. A., & Harmsen, M. C. (2008). Vascular smooth muscle cells for use in vascular tissue engineering obtained by endothelial-tomesenchymal transdifferentiation (EnMT) on collagen matrices. Biomaterials, 29, 3703–3711.

265 37. Campbell, J. H., Walker, P., Chue, W., Daly, C., Cong, H., Xiang, L., et al. (2004). Body cavities as bioreactors to grow arteries. International Congress Series, 1262, 118–121. 38. Hirst, S. L. (1996). Airway smooth muscle cell culture: application to studies of airway wall remodelling and phenotype plasticity in asthma. European Respiratory Journal, 9, 808–820. 39. James, A., & Carroll, N. (2000). Airway smooth muscle in health and disease; methods of measurement and relation to function. European Respiratory Journal, 15, 782–789. 40. Chakir, J., Pagé, N., Hamid, Q., Laviolette, M., Boulet, L. P., & Rouabhia, M. (2000). Bronchial mucosa produced by tissue engineering: a new tool to study cellular interactions in asthma. Journal of Allergy and Clinical Immunology, 107(1), 36–40. 41. Atala, A., Bauer, S., Soker, S., Yoo, J. J., & Retik, A. B. (2006). Tissue-engineered autologous bladders for patients needing cytoplasty. Lancet, 367, 1241–1246. 42. Jack, G. S., Zhang, R., Lee, M., Xu, Y., Wu, B. M., & Rodriguez, L. V. (2009). Urinary bladder smooth muscle engineered from adipose stem cells and a three dimensional synthetic composite. Biomaterials, 30, 3259–3270. 43. Sievert, K., Amend, B., & Stenzl, A. (2007). Tissue engineering for the lower urinary tract: a review of a state of the art approach. European Urology, 52, 1580–1589. 44. Baumert, H., Simon, P., Hekmati, M., Fromont, G., Levy, M., Balaton, A., et al. (2007). Development of a seeded scaffold in the great omentum: feasibility of an in vivo bioreactor for bladder tissue engineering. European Urology, 52, 884–892. 45. Kima, Y. M., Jeona, E. S., Kima, M. R., Jhob, S. K., Ryuc, S. W., & Kima, J. H. (2008). Angiotensin II-induced differentiation of adipose tissue-derived mesenchymal stem cells to smooth musclelike cells. The International Journal of Biochemistry & Cell Biology, 40, 2482–2491. 46. Yang, P., Yin, S., Cui, L., Li, H., Wu, Y., Liu, W., et al. (2008). Experiment of adipose derived stem cells induced into smooth muscle cells. Zhongguo Xiu Fu Chong Jian Wai Ke Za Zhi, 22(4), 481–486. 47. Ringe, J., Kaps, C., Burmester, G., & Sittinger, M. (2002). Stem cells for regenerative medicine: advances in the engineering of tissues and organs. Naturwissenschaften, 89, 338–351. 48. Moore, T. J. (2007). Stem cell Q and A—an introduction to stem cells and their role in scientific and medical research. Medical Technology SA, 21(1), 3–6. 49. Yanez, R., Lamana, L., Garcia-Castro, J., Colmenero, I., Ramirez, M., & Bueren, J. A. (2008). Adipose tissue-derived mesenchymal stem cells have in vivo immunosupressive properties applicable for the control of graft-versus-host disease. Stem Cells, 24, 2582– 2591. 50. Mvula, B., Mathope, T., Moore, T., & Abrahamse, H. (2007). The effect of low level laser therapy on adipose derived stem cells. Lasers in Medical Science, 23(3), 277–282. 51. Mvula, B., Moore, T. J., Abrahamse, H. (2009). Effect of low-level laser irradiation and epidermal growth factor on adult human adipose-derived stem cells. Lasers in Medical Science. doi:10.1007/ s10103-008-0636-1. 52. Dormorov, M. G., Kropp, B. P., Hurst, R. E., Cheng, E. Y., & Hsueh-Kung, L. (2007). Differentially expressed gene networks in cultured smooth muscle cells from normal and neuropathic bladder. Journal of Smooth Muscle Research, 43(2), 55–72. 53. Lin, H. K., Cowan, R., Moore, P., Zhang, Y., Yang, Q., Jr., Peterson, J. A., et al. (2004). Characterization of neuropathic bladder smooth muscle cells in culture. Journal of Urology, 171, 1348–1352.