Skip to main content
NCBI home page
NIHPA Author Manuscripts logoLink to NIHPA Author Manuscripts
. Author manuscript; available in PMC: 2013 Jul 1.
Published in final edited form as: J Tissue Eng Regen Med. 2011 Jul 14;6(7):505–511. doi: 10.1002/term.451

Phenotypic changes in cultured smooth muscle cells: limitation or opportunity to tissue engineering of hollow organs?

Alexander Huber 1, Stephen F Badylak 1,✉
PMCID: PMC3237740  NIHMSID: NIHMS307355  PMID: 21755602

Abstract

Smooth muscle cells (SMCs) are typically used as a cell source for the reconstruction of hollow organs by conventional tissue engineering techniques. However, the necessity and advantage for the use of tissue specific SMCs is unknown. The present study investigated the phenotypic changes that occur following isolation and in vitro expansion of rat SMC populations isolated from three different tissues: the aorta, esophagus, and urinary bladder. rSMCs were isolated by enzymatic dispersion and expanded by conventional cell culture techniques yielding microscopically homogeneous populations. SMC phenotypes were monitored according to their expression of marker proteins during the first two passages. Two of the three SMC populations (rSMC-a and rSMC-e) showed a marked change in their marker protein profiles during the first two passages which resulted in a homogenous phenotype that was neither fully contractile nor fully synthetic. SMCs from the urinary bladder did not show such a shift. Differences between the three rSMC populations were observed with regard to proliferative activity and gene expression patterns suggesting the retention of some tissue-specific cell characteristics. In summary, phenotypic changes in SMCs occur as a result of conventional cell isolation and expansion techniques, thus questioning the necessity of a tissue specific cell source for regenerative medicine applications.

1. Introduction

The therapeutic replacement or regeneration of functional tissue following loss due to trauma, neoplasia, or congenital defects is challenging at best. The field of tissue engineering/regenerative medicine has emerged as a potential strategy for the replacement of various tissues including osteochondral (Frohlich et al., 2008, Kandel et al., 2008, Soucacos et al., 2008), urogenital (Atala, 2009), cardiovascular (Habib et al., 2008, Yuasa and Fukuda, 2008), respiratory (Nichols and Cortiella, 2008), hepatic (Fiegel et al., 2009), and lymphatic (Hitchcock and Niklason, 2008) tissue among others. Conventional tissue engineering approaches include the use of structurally and functionally defined scaffold materials of natural or synthetic origin in combination with allogeneic or autogeneic cells of a single or a small number of different cell types. As such, the construction of even small-scale tissue engineered devices for clinical application requires the isolation and expansion of large numbers of cells while maintaining cellular phenotype and physiology without inducing events of cellular senescence (Hayflick and Moorhead, 1961) and dedifferentiation (Shi et al., 2008, Yu et al., 2007).

Smooth muscle cells (SMCs) play an important role in the functionality of numerous tissues and organs, including the vasculature, gastrointestinal tract, urinary bladder, respiratory tract, and the reproductive tract. Several cellular tissue engineering approaches have investigated the use of SMCs in the reconstruction of hollow organs including the esophagus (Nakase et al., 2008, Saxena et al., 2009), intestine (Nakase et al., 2006), internal anal sphincter (Somara et al., 2009), and blood vessels (Zhang et al., 2009). There is surprising heterogeneity with regard to the developmental origin and phenotypic characteristics of SMCs (Owens et al., 2004, Rensen et al., 2007). SMCs have been characterized as showing either a proliferative “synthetic” phenotype or a biomechanically active, but quiescent, “contractile” phenotype. Phenotypic plasticity (“phenotypic switching”) can occur reversibly and transiently between these two forms in vitro and in vivo (Aikawa et al., 1997, Thyberg et al., 1995, Thyberg et al., 1997) and has been observed in response to biochemical and mechanical cues (Beamish et al., Shen et al., 2006).

The use of SMCs, or for that matter any cell type, as part of a tissue engineering strategy for functional tissue replacement requires the reliable expansion of cells in vitro while minimizing the processes of dedifferentiation and/or controlling the processes of cellular redifferentiation towards the desired cellular phenotype. The events associated with SMCs' plasticity have mostly been studied in vascular SMCs. However, the actual process of cellular dedifferentiation, phenotypic changes, and senescence associated with in vitro cell expansion have been largely unexplored. Similarly, the methods to either prevent or proactively induce such changes by biochemical or biomechanical means are unknown.

The present study investigated the phenotypic changes that occur in rat smooth muscle cells harvested from three different tissues (aorta, esophagus, and urinary bladder), during the processes of initial cell isolation and early stages of cell maintenance and expansion using conventional cell culture techniques.

2. Material and Methods

2.1 rSMC Isolation and in vitro Cell Culture

Rat Smooth Muscle Cells (rSMCs) were isolated under sterile conditions from three different tissues; aorta, esophagus, and urinary bladder by enzymatic dispersion technique (Sreejayan and Yang, 2007). Briefly, following tissue isolation, all connective, adipose tissue and the adventitia were removed. The tissues were opened longitudinally (esophagus: along the greater curvature side) and the luminal surface layers (aorta: endothelium; esophagus/urinary bladder: epithelium, mucosa, and submucosa) were removed mechanically by scraping with a scalpel blade. The tissues were washed three times in Krebs Ringer solution (Sigma, St. Louise, MT, USA containing 0.015 mM NaHCO3, 3.0 mM CaCl2, 100 U/ml Pen-Strep). The tissues were minced and digested 30 min at 37°C under agitation in Krebs Ringer solution containing 15 U/ml elastase (Calbiochem), 20 U/ml papain (Sigma, St. Louise, MT, USA), 200 U/ml collagenase type II (Gibco), 0.4 mg/ml trypsin inhibitor (Soybean) (Sigma, St. Louise, MT, USA), and 1.7 mg/ml of bovine serum albumin (Sigma, St. Louise, MT, USA). Single cell suspensions were prepared by trituration through a glass pipette. Undigested tissue fragments were removed by centrifugation. Cells were plated at 2.5 × 104 cells/cm2 in complete culture medium (RPMI 1640 + 10% fetal bovine serum + ab/am) in a T25 tissue culture-treated cell culture flask in a humidified incubator, 37°C, 5% CO2 in air. Culture media was replaced after 24 hrs to remove dead cells and grown until 90% confluent changing the media every 2-3 days.

At near confluency, cells were collected by trypsinization and expanded further in a T75 tissue culture flask. Again at near confluency, cells were collected by trypsinization and expanded further in a T175 tissue culture flask.

Cells were cultured for two passages following isolation. At each subculturing step, a subset of cells (5.0 × 104 cells) was subcultured on collagen type I-coated glass coverslips (diameter: 2.0 cm) for immunocytochemical evaluation.

2.2 Immunohistochemical Staining

The expression of marker proteins for contractile and synthetic phenotypes of SMC was evaluated by conventional immunocytochemical techniques for cells grown on collagen type I-coated glass coverslips. All samples were processed in parallel to enable comparable processing and staining times between samples. Cells were fixed and permeabilized in ice-cold methanol for 5 min and rehydrated in phosphate buffer solution. Samples were blocked with solution consisting of PBS and 10% of the appropriate serum at room temperature for 30 min. Endogenous peroxidase activity was quenched with 0.3% hydrogen peroxide solution. Slides were then incubated sequentially with primary antibody (Ab) at 4°C overnight. Primary Abs against smooth muscle actin (1:200; Dako Cytomation, Carpinteria CA, USA), Smoothelin (1:100; Santa Cruz Biotech Inc., Santa Cruz, CA, USA), and cellular retinol binding protein 1 (1:50; Santa Cruz Biotech Inc., Santa Cruz, CA, USA) were applied and followed by biotinylated anti-goat, anti-mouse, or anti-rabbit Ab, respectively. After washing, slides were incubated with avidin–biotin conjugate of horseradish peroxidase (ABC; Vector Labs, Burlingame, CA, USA) in PBS for 30 min at room temperature. To identify the resulting peroxidase activity, the slides were treated with 3,3′-diaminobenzidine (DAB) solution (Vector Labs, Burlingame, CA, USA). The slides were counterstained with hematoxylin and mounted on glass slides with mounting medium (Vector). The cells were visualized and photographed by conventional light microscopy (Nikon E600, Nikon Instr. Inc., Melville, NY, USA).

2.3 Cell Proliferation Assay

The rate of rSMC proliferation was determined as described previously (Rago et al., 1990). Briefly, following trypsinization, cells were suspended in complete growth media. Cell standards were prepared by serial dilutions (1-125.0× 104 cells in 100 μl) on a 96 well cell culture plate (Corning, Inc., Corning, NY, USA). Cells were left to adhere for 8-10 hrs in the incubator (37°C, 5% CO2 in air) before further processing. Experimental plates were prepared by culturing 5.0× 104 cells in 100 μl complete culture medium in a 96 well plate. Cells were incubated for 1-3 days. At each endpoint, the culture medium was removed and the cells were frozen at -80 °C.

On the day of the assay, 100 μl dH2O was added to each well and incubated at room temperature for 60 min. The plates were frozen at -80 °C and thawed to reach room temperature before adding 100 μl of 20 μg/ml Hoechst 33258 in TNE buffer (10.0 mM Tris, 1.0 mM EDTA, 2.0 M NaCl, pH 7.4). Sample emissions (460nm) were measured on a microplate reader (SpectraMax M2, Molecular Devices, Inc., Sunnyvale, CA, USA) at an excitation of 350 nm. Readings were normalized to medium alone.

2.4 Gene Expression Analysis by RealTime-PCR

The gene expression activity of marker proteins for contractile and synthetic phenotypes of SMC was evaluated by qPCR in a two-step protocol. Total RNA was isolated from the explanted cells using RT TRI-Reagent (Molecular Research Center Inc., Cincinnati, OH, USA) according to the manufacturer's instructions. The isolated total RNA was purified using an RNAeasy Mini Kit (Qiagen, Valencia, CA, USA). cDNA was synthesized from 1.0 μg of total RNA in a volume of 20 μl using a Superscript RT III kit (Invitrogen, Carlsbad, CA, USA) according to the manufacturer's instructions.

Quantitative real-time PCR was performed using primers specific for genes of contractile or synthetic marker proteins (Huber Table1). Gene primer sequences were custom designed using Beacon Designer 7.2 primer design software (PREMIER Biosoft International, Palo Alto, CA) or as described elsewhere. Primers for the housekeeping gene Glyceraldehyde 3-phosphate dehydrogenase (gapdh) were purchased as part of a housekeeping gene primer kit (Rat Housekeeping Gene Primer Set, Real Time Primers, Elkins Park, PA, USA), and gapdh expression was used to normalize reactions. 1.0 μg of cDNA was mixed with the appropriate primers and 2× SYBR Green Master Mix (BioRad, Hercules, CA, USA) in a total volume of 25.0 μl. All reactions were performed in triplicate and were monitored using an iQ5 Real-Time Detection System (BioRad, Hercules, CA, USA). Absolute gene expression levels normalized to the housekeeping gene were expressed in their respective threshold cycles for each gene of interest.

Table 1.

Primer Sequences for RT-PCR.

gene GenID forward primer (5′-3′) reverse primer (5′-3′) reference
acta2 81633 AGCCAGTCGCCATCAGGAAC TCATCACCAGCAAAGCCCG Bouchez et al., 2008
col1a1 29393 ATCAGCCCAAACCCCAAGGAGA CGCAGGAAGGTCAGCTGGATAG Heinemeier et al., 2007
col1a2 84352 TTCCCGGTGAATTCGGTCTC ACCTCGGATTCCAATAGGACCAG Nozaki et al., 2005
col3a1 84032 TGGGATCCAATGAGGGAGAA CTCATGGCCTTGCGTGTTTG Petrick et al., 2009
eln 25043 CCTGTCCCTGACTCCCATTA CAGTGTGAGAAGTCGTCGGA Gray et al., 2008
cnn1 65204 GCCAGGTGTATGATCCCAAGT ATTGTGGGTGGGCTCG Hendriks-Balk et al., 2008
myh11 24582 GAAGCAGCCAGCATCAAG CAGCACAGAAGCCTCATTG Beacon
rbp-1 25056 AGGCATAGATGACCGCAAGT TCATCACCCTCAATCCACTG Mezaki et al., 2007
smtn 289734 CCAGAGGCTCCTCTAACACTAAGAG TTGGCTCTTGATTTTGGGTTGGCTG Ross et al., 2006
Open in a new tab

3. Results and Discussion

Cell based tissue engineering approaches require the reliable isolation and expansion of cells in vitro, and the maintenance of the cells' tissue-specific phenotype while preventing undesirable events such as cellular senescence and de- and transdifferentiation. A variety of cell types have been used for the reconstruction of complex tissues and organs as part of a conventional cellular tissue engineering approach. For instance, SMCs have been suggested as a useful cell type for the reconstruction of hollow organs including the esophagus (Nakase et al., 2008, Saxena et al., 2009), intestine (Nakase et al., 2006), internal anal sphincter (Somara et al., 2009), and blood vessels (Zhang et al., 2009). However, previous research has demonstrated the phenotypic plasticity of SMCs to transiently switch between a contractile (functional) and synthetic (proliferative) phenotype as indicated through the gradual expression of several marker proteins (Beamish et al., Rensen et al., 2007).

The present study evaluated the effects of conventional cell isolation and expansion techniques on the phenotype of rat SMCs isolated from three different tissues (aorta, esophagus, and urinary bladder) during early stages of cell expansion. Conventional cell isolation and pre-plating techniques resulted in the successful establishment of in vitro cell cultures of SMCs from rat aorta (rSMC-a), esophagus (rSMC-e), and urinary bladder (rSMC-ub). The isolated cell cultures were homogeneous in their microscopic appearance (Huber Figure1).

Figure 1.

Figure 1

Open in a new tab

Microscopic Appearance of Rat Smooth Muscle Cells grown in cell cultures in vitro as isolated by enzymatic dispersion technique from aorta (rSMC-a, A), esophagus (rSMC-e, B), and urinary bladder (rSMC-ub, C); scale bar = 150 μm.

These cells dedifferentiated within two passages on tissue culture-treated culture flasks towards phenotypically homogeneous cell populations showing a distinct alteration in the expression of SMC marker proteins as determined by immunocytochemical staining (Huber Figure2). Cells from all three tissues stained positive for the presence of alpha smooth muscle actin at the early and later stage of cell expansion. No SMC population stained positive for the presence of the CRBP-1, a synthetic marker, nor for the expression of smoothelin, a late contractile marker protein. While urinary bladder SMCs showed a low level of calponin expression, esophageal SMCs and aortic SMCs showed increasing levels of calponin between passages (rSMC-a: none → moderate; rSMC-e: weak → moderate). While a clear distinction between the synthetic and contractile phenotype is difficult due to the absence of a set of clear-cut markers and the nature of transient marker gene expression, the synthetic marker cellular retinol binding protein-1 (CRBP-1), and contractile markers such as alpha-smooth muscle actin (Aikawa et al., 1997), calponin, and smoothelin (Christen et al., 1999) have been described as a good indicator for the phenotypic state of SMCs. As a result, the expanded rSMC populations isolated as part of this study were determined to be of neither a fully synthetic, nor fully contractile phenotype, as demonstrated by lack of CRBP-1 and smoothelin, respectively.

Figure 2.

Figure 2

Open in a new tab

Immunocytochemical Staining for the Presence of Marker Proteins. Cultures of rSMCs grown on collagen-coated glass coverslips were stained immunocytochemically for the presence of alpha-smooth muscle actin (α-SM actin), calponin, cellular retinol binding protein-1 (CRBP-1), and smoothelin in cell cultures following the first and second passage after cell isolation. positive controls (pos. ctrl.) from rat aorta (a), esophagus (b), and urinary bladder (c). Scale bars = 150 μm.

In spite of showing a remarkable level of similarity between the three rSMC populations with respect to their microscopic appearance and the presence of marker proteins, tissue-specific differences between the three rSMC populations were observed with regard to the cells' proliferative activity and gene expression patterns. rSMC-a showed an almost fourfold greater rate of cell proliferation, duplicating every 33.1 hours, in comparison with both rSMC-e and rSMC-ub cell populations that duplicated every 128.2 and 123.7 hours, respectively (Huber Figure3).

Figure 3.

Figure 3

Open in a new tab

Rates of Proliferation: 5.0× 104 rSMCs were grown on tissue culture-treated multi-well plates. The level of cell proliferation was assessed over a period of three days by Hoechst 33258 staining of DNA and expressed in relative change to day 1 (n = 2).

In addition, gene expression of structural proteins as well as marker proteins associated with phenotypic differentiation of SMCs, including collagens type I and III, elastin, smooth muscle actin, calponin, smooth muscle myosin heavy chain, and smoothelin, as determined by quantitative RT-PCR showed distinct tissue-specific differences between the three SMC populations (Huber Figure4). rSMC-ub showed the highest level of gene expression for alpha actin (acta2), while rSMC-a had the lowest levels, with rSMC-e having an intermediate level of alpha actin gene expression. Similar expression levels were observed for collagen type I (col1a1, col1a2), collagen type III (col3a2), retinol binding protein 1 (rbp-1), smoothelin (smtn), calponin (cnn1), and smooth muscle myosin heavy chain (myh11). The expression of elastin was the highest in rSMC-a cultures, the lowest in rSMC-ub and intermediate in rSMC-e cultures. Stated differently, each type of SMC showed a distinct gene expression profile.

Figure 4.

Figure 4

Open in a new tab

Expressional Profiling of rSMCs grown in vitro. Expanded rSMCs were evaluated for their expression of structural and functional gene associated with SMC phenotype including collagen I (col1a1 and col1a2), collagen II (col3a2), elastin (eln), alpha-smooth muscle actin (acta2), calponin (cnn1), and smooth muscle myosin heavy chain (myh11), retinol binding protein-1 (rbp-1), and smoothelin (smtn). Results presented as absolute threshold levels (Ct), normalised to gapdh, n = 3.

This study focused only on a small subset of SMC-specific marker genes and proteins for the evaluation of a phenotypic shift during SMC expansion in vitro. Although these have previously been described as reliable markers for the evaluation of SMC phenotypic populations, a more detailed understanding in the molecular changes affecting these during SMC dedifferentiation in culture is desirable to evaluate SMC cell states. Such a study would also allow for a more detailed analysis of the effects of the environmental cues (i.e. the microenvironmental niche) upon cell phenotype outlined above.

The phenotypic switch observed in SMC populations may be used to control the tissue-specific phenotype of cells as part of a cellular tissue engineering approach by controlling environmental conditions during the cell expansion, tissue assembly, and neotissue maturation. Methods to maintain cell phenotype may include soluble factors (e.g. growth factors), cell culture environments (e.g. synthetic and natural scaffold materials), and biomechanical stimulation (e.g. flow or mechanical stretch). A more detailed understanding of the effects of these and other environmental cues on SMC phenotype is needed to allow for optimization of tissue engineering/regenerative medicine strategies for hollow organ reconstruction. Future experiments will use various environmental conditions to control and regulate tissue-specific SMC phenotypes.

Previous experiments in the tissue engineering of hollow organs including blood vessels (Furukawa et al., 2002, Nelson et al., 2008, Yang et al., 2005, Zhang et al., 2009), esophagus (Marzaro et al., 2006, Nakase et al., 2008, Saxena et al., 2009, Zhu et al., 2005, Zhu et al., 2006) and urinary bladder (Azzarello et al., 2009, Baumert et al., 2007, Gabouev et al., 2003, McManus et al., 2007, Nakanishi et al., 2003, Pariente et al., 2002, Pattison et al., 2005, Zhang et al., 2004, Zhang et al., 2000) following the conventional cellular tissue engineering paradigm have employed the use of SMCs from their respective tissue-specific cell source. However, the controlled regulation of SMC phenotype may allow the use of SMCs from one tissue, e.g. bladder SMCs, for the in vitro reconstruction of another tissue, e.g. esophagus. Therefore, the source of SMCs for tissue engineering applications may be irrelevant if a cell expansion phase is included as part of the strategy for creating a replacement organ. Experimental findings in the use of intestinal and urinary bladder SMCs in urinary bladder wall replacements in the rabbit seem to support this view (Lai et al., 2006). Consequentially, the phenotypic plasticity of certain cells, in particular the plasticity of SMCs, may be exploited as part of a tissue engineering approach by using cells from an easily obtainable tissue location, e.g. via a skin biopsy, without the need to isolate cells from the functionally compromised tissue that is to be reconstructed.

The extent of phenotypic changes within a particular cell type are of great importance with respect to the paradigm of cellular tissue engineering. While a certain, yet reversible, degree of dedifferentiation may be tolerable, it is fundamental that tissue engineered constructs are structurally and functionally complementary to the target host tissue. Such tissue specificity will require the strict control of cellular phenotype of each cell type used in a tissue engineering approach throughout the tissue engineering process, from initial cell isolation, expansion, and seeding, to neotissue assembly, maturation, and transplantation. The careful design of the appropriate, tissue-specific cell culture conditions including the application of biochemical stimuli (i.e. growth factors and cell culture supplements) and biomechanical stimuli will need to be taken into consideration for any tissue engineering approach. This must not be seen as an insurmountable obstacle but as an essential control mechanism ensuring the quality of engineered tissues and organs.

Acknowledgments

This study was supported in part by the National Institute of Health (NIH, 5R01EB008051) and the Armed Forces Institute for Regenerative Medicine (AFIRM, W81XWH-08-2-0032).

Footnotes

Disclosure Statement: No competing financial interests exist.

Contributor Information

Alexander Huber, Email: hubera@upmc.edu.

Stephen F. Badylak, Email: badylaks@upmc.edu.

References

  1. Aikawa M, Sakomura Y, Ueda M, et al. Redifferentiation of smooth muscle cells after coronary angioplasty determined via myosin heavy chain expression. Circulation. 1997;96:82–90. doi: 10.1161/01.cir.96.1.82. [DOI] [PubMed] [Google Scholar]
  2. Atala A. Engineering organs. Curr Opin Biotechnol. 2009;20:575–592. doi: 10.1016/j.copbio.2009.10.003. [DOI] [PubMed] [Google Scholar]
  3. Azzarello J, Kropp BP, Fung KM, et al. Age-dependent vascular endothelial growth factor expression and angiogenic capability of bladder smooth muscle cells: implications for cell-seeded technology in bladder tissue engineering. J Tissue Eng Regen Med. 2009;3:579–589. doi: 10.1002/term.199. [DOI] [PubMed] [Google Scholar]
  4. Baumert H, Simon P, Hekmati M, et al. Development of a seeded scaffold in the great omentum: feasibility of an in vivo bioreactor for bladder tissue engineering. Eur Urol. 2007;52:884–890. doi: 10.1016/j.eururo.2006.11.044. [DOI] [PubMed] [Google Scholar]
  5. Beamish JA, He P, Kottke-Marchant K, et al. Molecular Regulation of Contractile Smooth Muscle Cell Phenotype: Implications for Vascular Tissue Engineering. Tissue Eng Part B Rev. doi: 10.1089/ten.teb.2009.0630. [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Bouchez G, Sensebe L, Vourc'h P, et al. Partial recovery of dopaminergic pathway after graft of adult mesenchymal stem cells in a rat model of Parkinson's disease. Neurochem Int. 2008;52:1332–1342. doi: 10.1016/j.neuint.2008.02.003. [DOI] [PubMed] [Google Scholar]
  7. Christen T, Bochaton-Piallat ML, Neuville P, et al. Cultured porcine coronary artery smooth muscle cells - A new model with advanced differentiation. Circulation Research. 1999;85:99–107. doi: 10.1161/01.res.85.1.99. [DOI] [PubMed] [Google Scholar]
  8. Fiegel HC, Kneser U, Kluth D, et al. Development of hepatic tissue engineering. Pediatr Surg Int. 2009;25:667–673. doi: 10.1007/s00383-009-2389-8. [DOI] [PubMed] [Google Scholar]
  9. Frohlich M, Grayson WL, Wan LQ, et al. Tissue engineered bone grafts: biological requirements, tissue culture and clinical relevance. Curr Stem Cell Res Ther. 2008;3:254–264. doi: 10.2174/157488808786733962. [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Furukawa KS, Ushida T, Toita K, et al. Hybrid of gel-cultured smooth muscle cells with PLLA sponge as a scaffold towards blood vessel regeneration. Cell Transplant. 2002;11:475–480. [PubMed] [Google Scholar]
  11. Gabouev AI, Schultheiss D, Mertsching H, et al. In vitro construction of urinary bladder wall using porcine primary cells reseeded on acellularized bladder matrix and small intestinal submucosa. Int J Artif Organs. 2003;26:935–942. doi: 10.1177/039139880302601011. [DOI] [PubMed] [Google Scholar]
  12. Gray MA, Wang CC, Sacks MS, et al. Time-dependent alterations of select genes in streptozotocin-induced diabetic rat bladder. Urology. 2008;71:1214–1219. doi: 10.1016/j.urology.2007.11.054. [DOI] [PubMed] [Google Scholar]
  13. Habib M, Caspi O, Gepstein L. Human embryonic stem cells for cardiomyogenesis. J Mol Cell Cardiol. 2008;45:462–474. doi: 10.1016/j.yjmcc.2008.08.008. [DOI] [PubMed] [Google Scholar]
  14. Hayflick L, Moorhead PS. The serial cultivation of human diploid cell strains. Exp Cell Res. 1961;25:585–621. doi: 10.1016/0014-4827(61)90192-6. [DOI] [PubMed] [Google Scholar]
  15. Heinemeier KM, Olesen JL, Haddad F, et al. Expression of collagen and related growth factors in rat tendon and skeletal muscle in response to specific contraction types. J Physiol. 2007;582:1303–1316. doi: 10.1113/jphysiol.2007.127639. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Hendriks-Balk MC, van Loenen PB, Hajji N, et al. S1P receptor signalling and RGS proteins; expression and function in vascular smooth muscle cells and transfected CHO cells. Eur J Pharmacol. 2008;600:1–9. doi: 10.1016/j.ejphar.2008.09.041. [DOI] [PubMed] [Google Scholar]
  17. Hitchcock T, Niklason L. Lymphatic tissue engineering: progress and prospects. Ann N Y Acad Sci. 2008;1131:44–49. doi: 10.1196/annals.1413.004. [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Kandel R, Roberts S, Urban JP. Tissue engineering and the intervertebral disc: the challenges. Eur Spine J. 2008;17(4):480–491. doi: 10.1007/s00586-008-0746-2. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Lai JY, Chang PY, Lin JN. A comparison of engineered urinary bladder and intestinal smooth muscle for urinary bladder wall replacement in a rabbit model. J Pediatr Surg. 2006;41:2090–2094. doi: 10.1016/j.jpedsurg.2006.08.013. [DOI] [PubMed] [Google Scholar]
  20. Marzaro M, Vigolo S, Oselladore B, et al. In vitro and in vivo proposal of an artificial esophagus. J Biomed Mater Res A. 2006;77:795–801. doi: 10.1002/jbm.a.30666. [DOI] [PubMed] [Google Scholar]
  21. McManus M, Boland E, Sell S, et al. Electrospun nanofibre fibrinogen for urinary tract tissue reconstruction. Biomed mater. 2007;2:257–262. doi: 10.1088/1748-6041/2/4/008. [DOI] [PubMed] [Google Scholar]
  22. Mezaki Y, Yoshikawa K, Yamaguchi N, et al. Rat hepatic stellate cells acquire retinoid responsiveness after activation in vitro by post-transcriptional regulation of retinoic acid receptor alpha gene expression. Archives of Biochemistry and Biophysics. 2007;465:370–379. doi: 10.1016/j.abb.2007.06.024. [DOI] [PubMed] [Google Scholar]
  23. Nakanishi Y, Chen G, Komuro H, et al. Tissue-engineered urinary bladder wall using PLGA mesh-collagen hybrid scaffolds: a comparison study of collagen sponge and gel as a scaffold. Journal of Pediatric Surgery. 2003;38:1781–1784. doi: 10.1016/j.jpedsurg.2003.08.034. [DOI] [PubMed] [Google Scholar]
  24. Nakase Y, Hagiwara A, Nakamura T, et al. Tissue engineering of small intestinal tissue using collagen sponge scaffolds seeded with smooth muscle cells. Tissue Eng. 2006;12:403–412. doi: 10.1089/ten.2006.12.403. [DOI] [PubMed] [Google Scholar]
  25. Nakase Y, Nakamura T, Kin S, et al. Intrathoracic esophageal replacement by in situ tissue-engineered esophagus. J Thorac Cardiovasc Surg. 2008;136:850–859. doi: 10.1016/j.jtcvs.2008.05.027. [DOI] [PubMed] [Google Scholar]
  26. Nelson GN, Mirensky T, Brennan MP, et al. Functional small-diameter human tissue-engineered arterial grafts in an immunodeficient mouse model: preliminary findings. Arch Surg. 2008;143:488–494. doi: 10.1001/archsurg.143.5.488. [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. Nichols JE, Cortiella J. Engineering of a complex organ: progress toward development of a tissue-engineered lung. Proc Am Thoracic Soc. 2008;5:723–730. doi: 10.1513/pats.200802-022AW. [DOI] [PubMed] [Google Scholar]
  28. Nozaki T, Takeyasu M, Hirao A, et al. Differentiation of Rat Dental Pulp-derived Cells into an Osteoblastic Lineage. Oral Science International. 2005;2 [Google Scholar]
  29. Owens GK, Kumar MS, Wamhoff BR. Molecular regulation of vascular smooth muscle cell differentiation in development and disease. Physiol Rev. 2004;84:767–801. doi: 10.1152/physrev.00041.2003. [DOI] [PubMed] [Google Scholar]
  30. Pariente JL, Kim BS, Atala A. In vitro biocompatibility evaluation of naturally derived and synthetic biomaterials using normal human bladder smooth muscle cells. J Urol. 2002;167:1867–1871. [PubMed] [Google Scholar]
  31. Pattison MA, Wurster S, Webster TJ, et al. Three-dimensional, nano-structured PLGA scaffolds for bladder tissue replacement applications. Biomaterials. 2005;26:2491–2500. doi: 10.1016/j.biomaterials.2004.07.011. [DOI] [PubMed] [Google Scholar]
  32. Petrick JS, Blachere FM, Selmin O, et al. Inorganic arsenic as a developmental toxicant: In utero exposure and alterations in the developing rat lungs. Mol Nutr Food Res. 2009;53:583–591. doi: 10.1002/mnfr.200800019. [DOI] [PMC free article] [PubMed] [Google Scholar]
  33. Rago R, Mitchen J, Wilding G. DNA fluorometric assay in 96-well tissue culture plates using Hoechst 33258 after cell lysis by freezing in distilled water. Anal Biochem. 1990;191:31–34. doi: 10.1016/0003-2697(90)90382-j. [DOI] [PubMed] [Google Scholar]
  34. Rensen SS, Doevendans PA, van Eys GJ. Regulation and characteristics of vascular smooth muscle cell phenotypic diversity. Neth Heart J. 2007;15:100–108. doi: 10.1007/BF03085963. [DOI] [PMC free article] [PubMed] [Google Scholar]
  35. Ross JJ, Hong Z, Willenbring B, et al. Cytokine-induced differentiation of multipotent adult progenitor cells into functional smooth muscle cells. J Clin Invest. 2006;116:3139–3149. doi: 10.1172/JCI28184. [DOI] [PMC free article] [PubMed] [Google Scholar]
  36. Saxena AK, Kofler K, Ainodhofer H, et al. Esophagus tissue engineering: hybrid approach with esophageal epithelium and unidirectional smooth muscle tissue component generation in vitro. J Gastrointest Surg. 2009;13:1037–1043. doi: 10.1007/s11605-009-0836-4. [DOI] [PubMed] [Google Scholar]
  37. Shen JY, Chan-Park MB, He B, et al. Three-dimensional microchannels in biodegradable polymeric films for control orientation and phenotype of vascular smooth muscle cells. Tissue Engineering. 2006;12:2229–2240. doi: 10.1089/ten.2006.12.2229. [DOI] [PubMed] [Google Scholar]
  38. Shi Y, Desponts C, Do JT, et al. Induction of pluripotent stem cells from mouse embryonic fibroblasts by Oct4 and Klf4 with small-molecule compounds. Cell Stem Cell. 2008;3:568–574. doi: 10.1016/j.stem.2008.10.004. [DOI] [PubMed] [Google Scholar]
  39. Somara S, Gilmont RR, Dennis RG, et al. Bioengineered internal anal sphincter derived from isolated human internal anal sphincter smooth muscle cells. Gastroenterology. 2009;137:53–61. doi: 10.1053/j.gastro.2009.03.036. [DOI] [PubMed] [Google Scholar]
  40. Soucacos PN, Johnson EO, Babis G. An update on recent advances in bone regeneration. Injury. 2008;39(2):S1–4. doi: 10.1016/S0020-1383(08)70009-3. [DOI] [PubMed] [Google Scholar]
  41. Sreejayan N, Yang X. Isolation and Functional Studies of Rat Aortic Smooth Muscle Cells. In: S N, R J, editors. Vascular Biology Protocols. Totowa, NJ: Humana Press Inc; 2007. pp. 283–292. [DOI] [PubMed] [Google Scholar]
  42. Thyberg J, Blomgren K, Hedin U, et al. Phenotypic modulation of smooth muscle cells during the formation of neointimal thickenings in the rat carotid artery after balloon injury: an electron-microscopic and stereological study. Cell Tissue Res. 1995;281:421–433. doi: 10.1007/BF00417860. [DOI] [PubMed] [Google Scholar]
  43. Thyberg J, Blomgren K, Roy J, et al. Phenotypic modulation of smooth muscle cells after arterial injury is associated with changes in the distribution of laminin and fibronectin. J Histochem Cytochem. 1997;45:837–846. doi: 10.1177/002215549704500608. [DOI] [PubMed] [Google Scholar]
  44. Yang J, Motlagh D, Webb AR, et al. Novel biphasic elastomeric scaffold for small-diameter blood vessel tissue engineering. Tissue Engineering. 2005;11:1876–1886. doi: 10.1089/ten.2005.11.1876. [DOI] [PubMed] [Google Scholar]
  45. Yu J, Vodyanik MA, Smuga-Otto K, et al. Induced pluripotent stem cell lines derived from human somatic cells. Science. 2007;318:1917–1920. doi: 10.1126/science.1151526. [DOI] [PubMed] [Google Scholar]
  46. Yuasa S, Fukuda K. Cardiac regenerative medicine. Circ J. 2008;72(A):A49–55. doi: 10.1253/circj.cj-08-0378. [DOI] [PubMed] [Google Scholar]
  47. Zhang X, Wang X, Keshav V, et al. Dynamic culture conditions to generate silk-based tissue-engineered vascular grafts. Biomaterials. 2009;30:3213–3223. doi: 10.1016/j.biomaterials.2009.02.002. [DOI] [PMC free article] [PubMed] [Google Scholar]
  48. Zhang Y, Kropp BP, Lin HK, et al. Bladder regeneration with cell-seeded small intestinal submucosa. Tissue Engineering. 2004;10:181–187. doi: 10.1089/107632704322791835. [DOI] [PubMed] [Google Scholar]
  49. Zhang Y, Kropp BP, Moore P, et al. Coculture of bladder urothelial and smooth muscle cells on small intestinal submucosa: potential applications for tissue engineering technology. J Urol. 2000;164:928–934. doi: 10.1097/00005392-200009020-00004. discussion 934-925. [DOI] [PubMed] [Google Scholar]
  50. Zhu Y, Chan-Park MB, Sin Chian K. The growth improvement of porcine esophageal smooth muscle cells on collagen-grafted poly(DL-lactide-co-glycolide) membrane. J Biomed Mater Res B Appl Biomater. 2005;75:193–199. doi: 10.1002/jbm.b.30305. [DOI] [PubMed] [Google Scholar]
  51. Zhu Y, Chian KS, Chan-Park MB, et al. Protein bonding on biodegradable poly(L-lactide-co-caprolactone) membrane for esophageal tissue engineering. Biomaterials. 2006;27:68–78. doi: 10.1016/j.biomaterials.2005.05.069. [DOI] [PubMed] [Google Scholar]

RESOURCES