Differentiation of Human Mesenchymal Stem/Stromal Cells into Myogenic Cells for Urethral Sphincter Muscle Engineering Sara Ferreira Martins Gomes Thesis to obtain the Master of Science Degree in Biotechnology Supervisors: Professor Cláudia Alexandra Martins Lobato da Silva Professor Joaquim Manuel Sampaio Cabral Examination Committee Chairperson: Professor Arsénio do Carmo Sales Mendes Fialho Supervisor: Professor Cláudia Alexandra Martins Lobato da Silva Member of the Committee: Professor Gabriel António Amaro Monteiro December 2015
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Differentiation of Human Mesenchymal Stem/Stromal Cells
into Myogenic Cells for Urethral Sphincter Muscle
Engineering
Sara Ferreira Martins Gomes
Thesis to obtain the Master of Science Degree in
Biotechnology
Supervisors: Professor Cláudia Alexandra Martins Lobato da Silva
Professor Joaquim Manuel Sampaio Cabral
Examination Committee
Chairperson: Professor Arsénio do Carmo Sales Mendes Fialho
Supervisor: Professor Cláudia Alexandra Martins Lobato da Silva
Member of the Committee: Professor Gabriel António Amaro Monteiro
December 2015
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I dedicate this thesis to the memory of my father, Henrique Martins Gomes (November 4th, 1946 - May 11th, 2015)
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Acknowledgments First, I would like to thank Professor Joaquim Cabral and Professor Cláudia Silva for making it
possible for me to develop my dissertation work at the Stem Cell Bioengineering and Regenerative
Medicine Laboratory (SCBL-RM), at IBB-BERG-IST, and for accepting to be my Supervisors. I am
truly grateful for the opportunity to work in this field of study, which I have been passionate about since
I was in high school. I also want to thank Professor Cláudia for all the guidance and encouragement
and for all the trust put into my work and my ideas.
Second, I owe a big thank you to Irina Simões, who guided me since the beginning, taught me
everything I needed to know about this project and was always accessible to help me even at
thousands of kilometres away in Zurich. I could not have achieved the results I present here without
her knowledge and direction, and I could not have been attributed a better mentor.
Third, I want to thank my colleagues at the SCBL-RM laboratory, who were always available
to help me and give me guidance when I needed: Ana Fernandes, Márcia Mata, Francisco Moreira,
Diogo Pinto, Raquel Cunha, Marta Costa, João Silva, Cláudia Miranda, Tiago Dias and Carlos
Rodrigues. You all had an important part in the development of this project and of my laboratory skills.
I want to give a warm thank you to Alexandra Salvado, Ângela Neves and Mafalda Cavalheiro
for being my partners in this journey, and for all the support and laughter that we shared over the last
two years. Without you, group assignments would have been extremely dull and definitely not as
productive as they were.
Last but definitely not least, I want to thank my family, especially the three most important
gentlemen in my life: my boyfriend/best friend, my brother, and my father. Rui, you are my rock: you
were there for me through the toughest times, holding me together and giving me a reason to fight for.
My dear Mano, I want to thank you for all the wise words, support and guidance, especially over the
last 6 months, and for encouraging me to always do my best. Dad, thank you for telling me how proud
of me you were; those words are what keep me going in the hardest moments. I hope I keep making
you proud. I deeply miss you.
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Abstract
Stress urinary incontinence (SUI) is a medical condition that requires novel alternative
therapies aiming to restore and maintain the integrity and function of the urethral sphincter, the muscle
layer responsible for the normal continence mechanism. This MSc project targeted the establishment
of effective myogenic differentiation protocols for Mesenchymal Stem/Stromal Cells (MSCs) for
urethral sphincter engineering, with particular focus on exploring the myogenic potential of the Stromal
Vascular Fraction (SVF) of the adipose tissue. The ability of MSCs to differentiate into smooth muscle
cells (SMCs) and skeletal myofibers, the main constituents of the sphincter, has already been
demonstrated in few in vivo and in vitro studies, but with little translation of this knowledge into clinical
settings.
The effects of 5-aza-2’-deoxycytidine (5-AZAd) and PD98059, chemical inducers of skeletal
muscle and smooth muscle differentiation in MSCs, respectively, were tested herein. MSC
differentiation into both cell types was evaluated by the detection of smooth and skeletal muscle
lineage-specific markers by flow cytometry and immunofluorescence techniques at different timepoints
in early cell passages (P<3). The expression of skeletal muscle markers was assessed in magnetically
sorted and unsorted SVF cells coated on distinct substrates.
Myogenesis-committed cells were identified in uncultured SVF, but no myoblast-like cells were
isolated. Still, SVF-derived cells were shown to possess intrinsic myogenic potential that was
enhanced when combined with culture substrates (in particular, gelatin coating), thus holding great
potential for skeletal muscle engineering applications. Conversely, 5-AZAd supplementation failed to
induce myogenesis and triggered severe cytotoxic effects, while PD98059 did not provide enough
stimuli to sustain smooth muscle differentiation, which likely requires the use of 3-D culture conditions
and/or biomechanical stimulation.
Keywords
Stress Urinary Incontinence
Mesenchymal Stem/Stromal Cells
Stromal Vascular Fraction
Myogenic Differentiation
Smooth Muscle Cells
Skeletal muscle Cells
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Resumo A incontinência urinária de esforço é uma condição que requer novas terapias que visem
restaurar e manter a integridade e função do esfíncter uretral, a camada de músculo responsável pelo
mecanismo de continência normal. Este Projeto de Mestrado teve como objectivo o desenvolvimento
de protocolos que permitam a diferenciação eficaz de células estaminais/estromais mesenquimais (ou
mesenquimatosas) - CEMs - em linhagens miogénicas para reconstituição do esfíncter uretral, com
especial foco no estudo do potencial miogénico da fracção vascular estromal (FVE) do tecido
adiposo.
Neste sentido, foi testado o efeito dos compostos 5-aza-2’-desoxicitidina (5-AZAd) e
PD98059, indutores químicos de diferenciação em músculo esquelético e liso, respectivamente, em
CEMs. A diferenciação de CEMs em ambos os tipos de músculo foi avaliada através da detecção de
marcadores específicos de cada linhagem, por técnicas de citometria de fluxo e imunofluorescência
em tempos de cultura distintos e em passagens celulares baixas (P<3). A expressão de marcadores
de músculo esquelético foi avaliada também em células da FVE minimamente processada e isoladas
magneticamente, plaqueadas sob superfícies revestidas com componentes da matriz extracelular
(gelatina e fibronectina).
Foram detectadas células miogénicas na FVE minimamente processada, no entanto não
foram isoladas células com morfologia mioblástica. Ainda assim, foi mostrado que as células obtidas
a partir da FVE são dotadas de um potencial miogénico intrínseco, reforçado na presença de
substratos, sendo potencialmente exequível a aplicação destas células em estratégias de
reconstituição de músculo esquelético. Por outro lado, a adição de 5-AZAd não levou à indução de
miogénese e conduziu a efeitos citotóxicos, enquanto que a adição de PD98059 não foi suficiente
para sustentar a diferenciação de CEMs em músculo liso, que provavelmente requer a
implementação de condições de cultura em 3-D e/ou estimulação biomecânica.
ACKNOWLEDGMENTS ......................................................................................................................... V
ABSTRACT ........................................................................................................................................... VII
RESUMO ................................................................................................................................................ IX
LIST OF FIGURES ............................................................................................................................... XIII
LIST OF TABLES ............................................................................................................................... XVI
ABBREVIATIONS LIST ..................................................................................................................... XVII
I. INTRODUCTION .............................................................................................................................. 1 I.1 BACKGROUND ............................................................................................................................... 1
I.2 MYOGENIC DIFFERENTIATION OF MSCS: CURRENT STATUS ......................................................... 18 I.2.1 Differentiation of MSCs into Skeletal Muscle Cells ........................................................... 18 I.2.2 Differentiation of MSCs into SMCs .................................................................................... 22
I.3 AIM OF STUDIES.......................................................................................................................... 26
II. MATERIALS AND METHODS ...................................................................................................... 27 II.1 HUMAN-DERIVED SAMPLES ......................................................................................................... 27 II.2 CULTURE MEDIA ........................................................................................................................ 27
II.2.1 Ex-vivo expansion of ADSCs, BM-MSCs and SVF-derived cells .................................... 27 II.2.2 Thawing and cryopreservation of SVF cells, ADSCs and BM-MSCs .............................. 27
II.5.1 Effect of 5-aza-2’-deoxycytidine on cell viability............................................................... 29 II.5.2 Effect of 5-aza-2’-deoxycytidine on cell apoptosis and morphology ................................ 29
II.6 CD34+ SVF-DERIVED CELL SORTING AND MYOGENESIS INDUCTION ................................................ 30 II.6.1 CD34/CD56 decay assessment in normal and ultralow attachment plates ..................... 30
III. RESULTS AND DISCUSSION .................................................................................................... 33 III.1 MESENCHYMAL STEM/STROMAL CELL ISOLATION....................................................................... 33
III.1.1 Mesenchymal Stem/Stromal Cell Characterization ........................................................ 35 III.2 DIFFERENTIATION OF MESENCHYMAL STEM/STROMAL CELLS INTO SKELETAL MUSCLE CELLS ..... 38
III.2.1 Myogenic induction using 5-aza-2’deoxycytidine ........................................................... 38 III.2.2 Myogenic induction of CD34+ cells from the Stromal Vascular Fraction ........................ 46
III.3 DIFFERENTIATION OF MESENCHYMAL STEM/STROMAL CELLS INTO SMOOTH MUSCLE CELLS ....... 63 III.4 DECELLULARIZATION OF PORCINE URETHRAS ............................................................................ 70
IV. FUTURE TRENDS AND CONCLUSIONS .................................................................................. 73
V. REFERENCES .............................................................................................................................. 75
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List of Figures
Figure I.1. The lower urinary tract in women and men. From Fry et al., 2009. ....................................... 1 Figure I.2. Cross-section of the upper portion of a female sphincter. From Heesakkers and Gerretsen,
2004. ............................................................................................................................................... 2 Figure I.3. The two main strategies involving stem cells in Regenerative Medicine-related clinical
applications. From Schmitt et al., 2012. ......................................................................................... 4 Figure I.4. The fertilized egg and its immediate progeny (until the 8-cell embryo) are totipotent cells.
The cells from the inner mass of the blastocyst are pluripotent, as they can only give rise to the 3 germ layers of the embryo. Once cultured in vitro, pluripotent stem cells are capable of differentiating into neural cells (ectoderm), cardiac muscle or blood cells (mesoderm), among others. From the website of Division of Biology and Medicine from Brown University (http://biomed.brown.edu).28 ........................................................................................................... 5
Figure I.5. Transcription factors involved in the regulation of myogenic lineage progression. While Pax3/7, Myf5 and MyoD are crucial for the initial specification and commitment of the progenitor cells, MyoG and Mrf4 are determinant for the early and late differentiation steps, leading to mature myotubes/myofibers. From Bentzinger et al., 2012. ........................................................... 9
Figure I.6. The Mesengenic Process: MSCs proliferate and their progeny can be induced to enter one of several mesenchymal lineage pathways, including marrow stroma, osteogenesis, chondrogenesis, tendogenesis, myogenesis and adipogenesis. Adapted from Caplan and Correa, 2011. ................................................................................................................................ 12
Figure I.7. The MSC perivascular niche hypothesis. Within the niche, O2 tension is variable. MSC in contact with blood vessels (BV) would interact with (1) various other differentiated cells (DC1, DC2, etc.) by means of cell-adhesion molecules, (2) ECM deposited by the niche cells (mediated by integrin receptors), and (3) signalling molecules. From Kolf et al., 2007................................. 16
Figure I.8. FACS vs. MACS. A) A fluorescence-activated sorter can be used as a preparative tool to separate fluorescently-labelled cells from a heterogeneous cell suspension. B) Through magnetic labelling, desired cells are firstly separated from contaminants and then eluted from the column. Left image from the website of Midlands Technical College (www.midlandstech.edu).116 Right image adapted from the website of Humboldt University of Berlin (http://edoc.hu-berlin.de/)117 . 20
Figure I.9. Structures of cytidine and its analogs, 5-AZA and 5-AZAd. R=ribose. dR=deoxyribose. Adapted from Christman, 2002. .................................................................................................... 21
Figure I.10. The ERK/MAPK signalling pathway. Its activation triggers the sequential phosphorylation of three kinases (MAPKKK,MAPKK and MAPK), resulting in the translocation of ERK to the nucleus and the activation of transcription factors (e.g. Elk-1 and c-Myc) which control the expression of genes that are required for cell growth, differentiation and survival. From Kim and Bar-Sagi, 2004. ............................................................................................................................. 24
Figure III.1. P0 SVF-derived cells cultured for 6 days in DMEM medium supplemented with 10% of (A) MSC-qualified FBS and (B) standard FBS. Arrow points to a pericyte-like cell. .......................... 34
Figure III.2. P0 SVF-derived cells cultured for 15 days in DMEM+3%FBS (A) and 11 days in DMEM+10%FBS (B). Some cells begin appearing more extended and flattened with increased culture time, possibly due to senescence. .................................................................................... 34
Figure III.3. Multilineage differentiation characterization of BM-MSCs, ADSCs and AT-SVF cells. BM-MSCs and ADSCs were isolated and plated in DMEM+10%FBS (MSC-qualified), while AT-SVF cells were isolated in DMEM media supplemented with 20%FBS (not MSC-qualified), dexamethasone and bFGF, before confluence was reached and the medium was replaced to the respective differentiation medium. Scale bar = 100 μm. .............................................................. 36
Figure III.4. Effect of 5-AZAd on cell morphology and viability. ADSCs were cultured for 7 days in DMEM+2%HS, harvested and placed on a haemocytometer for the trypan blue exclusion test. A) Control cells displayed typical round shapes; B) Cells exposed to 10 μM of 5-AZAd displayed a very thin elongated morphology and stained positively for trypan blue. ....................................... 39
Figure III.5. Cell viability assay for ADSCs cultured in DMEM+2%HS previously exposed to 5-AZAd. Cells were treated with various concentrations of 5-AZAd for 24 hours (2, 5, 7.5 and 10 μM) and cell viability was measured by counting cell numbers at 1, 3, 5, 7, 12 and 14 days thereafter. Two cell densities were tested: 3000 cells/cm2 (left) and 5000 cells/cm2 (right). No cell bars indicate zero cell viability, except for 5 μM. ................................................................................................ 39
Figure III.6 Effect of 5-AZAd on cell viability.......................................................................................... 41
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Figure III.7. Flow cytometry dot plot charts of ADSCs cultured for 2 days in DMEM+2%HS after 5-AZAd treatment. A) control ADSCs; B) ADSCs treated with 10 μM 5-AZAd. ............................... 41
Figure III.8. Alterations in cell morphology and confluence of ADSCs cultured in DMEM+2%HS after two weeks of 5-AZAd treatment. (A) Control ADSCs and ADSCs exposed to 2 μM (not shown) reached high confluence levels and maintained a fibroblast-like shape. (B) ADSCs exposed to 10 μM and 7.5 μM (not shown) of 5-AZAd achieved low confluence levels and two types of cell shapes could be discerned: thin elongated cells (black arrow) and broad irregular cells (white arrow). ........................................................................................................................................... 42
Figure III.9. Effect of 5-AZAd on cell morphology of ADSCs. Control ADSCs and ADSCs treated with 2 μM 5-AZAd were positive for phalloidin-stained F-actin (red), and DAPI-stained nuclei (blue) at days 2 (top), 7 (middle) and 14 (bottom). ADSCs treated with 10 μM 5-AZAd stained positively at day 2, but no fluorescence was detected thereafter. Scale bar = 100 μm. .................................. 43
Figure III.10. 5-AZAd effect on surface marker expression after 2, 7 and 14 days of treatment in ADSCs cultured in DMEM+2%HS. ............................................................................................... 44
Figure III.11. Evaluation of Pax7 expression in an uncultured SVF sample. (A) Gated flow cytometry dot plot chart in which 3 subpopulations could be discerned; (B) Pax7 expression values and number of Pax7-expressing cells in a total of 49.6x105 viable cells (C) Pax7 expression histograms for subpopulations 1, 2 and 3 (left to right). Red lines = control; Blue lines = sample. ...................................................................................................................................................... 46
Figure III.12. Flow cytometry dot plot charts of CD34-enriched (A) and CD34-depleted (B) fractions acquired after MACS for sample SVF-2. (C) Expression of CD34 and CD56 (in percentage and MFI) for both fractions. .................................................................................................................. 48
Figure III.13. Flow cytometry dot plot charts of unsorted SVF cells (A), and cells from the CD34-enriched (B) and CD34-depleted (C) fractions obtained after CD34 sorting for sample SVF-3. Two subpopulations (1 and 2) could be discerned in the CD34-enriched and CD34-depleted fractions. (D) Expression of CD34 and CD56 (in percentage and MFI). (E) Extracellular marker characterization of cells from the CD34-depleted fraction. ........................................................... 48
Figure III.14. Flow cytometry dot plot charts of unsorted SVF cells (A) CD34-enriched (B) and CD34-depleted (C) fractions obtained after CD34 sorting of a cell pool of SVF cells. Two subpopulations (1 and 2) could be discerned in the CD34-enriched fraction. (D) Expression of CD34 and CD56 (in percentage and MFI). ................................................................................... 49
Figure III.15. Evaluation of Pax7 expression by flow cytometry in an uncultured SVF sample (SVF-4) and in the enriched and depleted fractions obtained after CD34 MACS. (A) Gated flow cytometry charts; (B) Flow cytometry histograms of Pax7 expression in the unsorted SVF (left) and the CD34-enriched fraction (right); (C) Frequency, cell number and expression for each gated population and number of Pax7-expressing cells in a total of 48.6x105 viable SVF cells, 5.7x105 CD34 positive cells and 3.3x105 CD34 negative cells. ................................................................. 50
Figure III.16. Cells cultured on fibronectin-coated (A) and non-coated (B) plates, using LG DMEM media, derived from the CD34-enriched fraction of the SVF, acquired by MACS. Low confluence areas (left) and high confluence areas (right) could be obtained in all conditions, except for unsorted SVF cells, which were 100% confluent. Black arrows point to pericyte-like cells. ........ 51
Figure III.17. Flow cytometry analysis of CD34 and CD56 expression for SVF and cells cultured on non-coated plates, after 11 days in culture, and cells cultured on gelatin and fibronectin-coated plates, after 12 days in culture. Top: Flow cytometry dot plot charts. Bottom: Expression percentages and MFI values for CD34 and CD56 for each condition. MP = Main population (higher SSH and FSH gate), SP = Subpopulation (lower SSH and FSH gate). ........................... 52
Figure III.18. Expression of myogenic markers in P0 cells cultured on gelatin- and fibronectin-coated plates (HG) after 12 days in culture. Flow cytometry dot plot charts and histograms of Pax7, MyoD and myogenin (left to right) of the main population gated of cells cultured on fibronectin- (HG) (A) and gelatin-coated plates (B). (C) Myogenic marker expression (in percentage and MFI values) of the gated main populations and subpopulations of both conditions. ........................... 53
Figure III.19. Cells displaying distinct morphologies after 8 days of induction, detected by light microscopy (top) and fluorescence microscopy (bottom). (A) Cells cultured on fibronectin-coated plates (LG), in which aligned spindle-like and broad flattened cells can be discerned; (B) SVF cells, in which the development of several vacuoles is visible; (C) cells cultured on gelatin-coated plates and (D) cells cultured on non-coated plates; Broad flattened cells with enlarged nuclei (frequently binucleated) could be seen in all conditions, as well as smaller spindle-like cells. Nuclei were stained with DAPI and the cytoskeleton (F-actin filaments) was stained with phalloidin-TRITC. .......................................................................................................................... 54
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Figure III.20. Multinucleated cells (P1) present in a very low confluence area of a gelatin-coated well, cultured for 17 days, detected by DAPI and Phalloidin-TRITC staining. ...................................... 55
Figure III.21. MyoD expression assessed by immunofluorescence after 2, 4 and 8 days of medium replacement for all conditions (secondary antibody: goat anti-rabbit Alexa 546). From day 4 to day 8 a decrease in the fluorescence intensity and the number of fluorescent cells was seen. Phalloidin-TRITC and DAPI staining (bottom row) allowed the observation of aligned nuclei and some fused cells. No Pax7 and myogenin expression was observed through the assay for all the conditions. Scale bar = 50 μm. ..................................................................................................... 56
Figure III.22. Flow cytometry analysis of CD34 and CD56 expression after 2 and 3 days in culture for cells isolated from unsorted SVF and magnetically sorted cells from the CD34-enriched fraction. ...................................................................................................................................................... 58
Figure III.23. Aggregate adherence to gelatin-coated plates after 24 hours in culture. Left to right: unsorted SVF cells in expansion media; unsorted SVF cells in skeletal muscle cell growth media; CD34-enriched cell fraction in expansion medium; CD34-enriched cell fraction cell in skeletal muscle cell growth media. Scale bar = 100 μm. ........................................................................... 59
Figure III.24. Bladder SMCs (passaged 6) plated in bladder smooth muscle cell growth media after 8 days (A) and 21 days (B) of culture. ............................................................................................. 64
Figure III.25. Smooth muscle marker expression in cultured SMCs. A) Flow cytometry dot plot chart of bladder SMCs (P5) after 14 days in culture and B) percentage values of expression of smooth muscle markers. Secondary antibody: goat anti-mouse Alexa488. C) Immunofluorescence for markers α-SMA, SM-MHC, calponin and desmin detected in P6 bladder SMCs after 21 days in culture. Scale bar = 50 μm. Secondary antibodies: goat anti-mouse Alexa488 for α-SMA, calponin and SM-MHC; goat-anti rabbit Alexa 546 for desmin. ................................................... 64
Figure III.26. BM-MSCs (A) and SVF-derived cells (B) plated in DMEM+10%FBS, cultured for 3 and 9 days, respectively. ........................................................................................................................ 65
Figure III.27. Smooth muscle marker expression in AT-SVF cells and BM-MSCs after 2, 4 and 7 days of differentiation induction with PD98059, assessed by flow cytometry and immunofluorescence. A) Intracellular flow cytometry for α-SMA, calponin, SM-MHC and desmin. Secondary antibodies used: goat anti-mouse Alexa488 (in BM-MSCs assay); goat anti-mouse PE for α-SMA, calponin and SM-MHC and goat anti-rabbit Alexa 488 for desmin (in AT-SVF assay). B) Immunofluorescence after 7 days of induction for calponin (top) and desmin (bottom) in AT-SVF cells. C) Immunofluorescence after 7 days of induction for calponin in BM-MSCs. Secondary antibodies: goat anti-mouse Alexa488 for calponin and goat-anti rabbit Alexa 546 for desmin. Scale bar = 50 μm......................................................................................................................... 67
Figure III.28. (A) Schematics of the decellularization apparatus based on mechanochemical action. Briefly, the detergent solution (or distilled water) is placed inside the (a) vessel where the (b) cannulated urethra is submerged. The solution is pumped by a (c) peristaltic pump and re-circularized for 24 hours at a rate of 40 ml/min. Simultaneously, through a (d and e) magnetic stirrer plate the solution was agitated at 340 rpm. After 48 hours, the direction of perfusion was changed (from the black to the grey arrows). Adapted from Simões et al., 2015 (under revision). (B) Length and protocol duration for each decellularized urethra. (C) Main steps of the decellularization process and their outcomes for urethra #4. Biological tissue preparation includes AT and distal extremity removal, catheter placement and immobilization with sutures (all performed in day 0). Gradual whitening of the tissue over perfusion time occurred, indicating efficient cell removal. .................................................................................................................... 71
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List of Tables
Table II.1. Panel of mouse anti-human monoclonal antibodies used to characterize MSCs and SVF-derived cells, their commercial brands, conjugated fluorophores and isotypes. .......................... 29
Table II.2. Panel of anti-human primary monoclonal antibodies and respective fluorophore-conjugated secondary antibodies used to stain cells for markers relevant for myogenic differentiation assessment, dilution used, commercial brand and isotype. ......................................................... 31
Table III.1. Age and gender of the AT donors and the assays performed with each sample. .............. 33 Table III.2. Expression of the markers used in the identification of SVF-derived cells and MSCs,
according to Bourin et al. and Dominici et al., by several P0 SFV-derived plate-adherent populations cultured in distinct media, and by P0 BM-MSCs. For AT-SVF cells in expansion medium n=3 (except CD14 and HLA-DR, for which n=1). FBS*= MSC-qualified FBS ................ 35
Table III.3. Effect of 5-AZAd on cell viability. PI/Annexin V flow cytometry analysis for ADSCs cultured in DMEM+2%HS, treated with 2 and 10 μM of 5-AZAd, at days 2, 7 and 14 after treatment. The expression of annexin and PI served as a measure of cell viability, apoptosis and necrosis. ..... 41
Table III.4. 5-AZAd effect on surface marker expression after 2, 7 and 14 days of treatment in ADSCs cultured in DMEM+2%HS. ............................................................................................................ 44
Table III.5. Experiments performed with cells from different SVF donors. *Due to a shortage in SVF sample numbers, a cell pool combining samples SVF-4 and SVF-5 had to be performed. ......... 47
Table III.6. MACS yield for samples subjected to MACS. Total cell numbers of the SVF samples and the CD34 enriched and depleted fractions acquired after MACS were calculated by viable cell counts using the trypan blue exclusion test. The numbers of CD34+ cells within the samples and/or the positive fractions were calculated using values acquired by flow cytometry. .............. 48
Table III.7. Expression of myogenic markers (Pax7, MyoD, myogenin and skeletal MHC) in P1 cells after 2, 4 and 8 days of differentiation induction, assessed by flow cytometry. Secondary antibodies: Goat-anti mouse/rabbit Alexa 488 for myogenin (day 2) and MyoD (days 2 and 4), respectively; goat-anti mouse/rabbit PE for myogenin (days 4 and 8) and MyoD (day 8). .......... 55
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Abbreviations List
ADSCs Adipose derived-Stromal/Stem Cells
AT Adipose Tissue
bFGF Basic Fibroblast Growth Factor
BM Bone Marrow
BM-MSCs Bone Marrow-derived Mesenchymal Stem/Stromal Cells
CD Cluster of Differentiation
DMEM Dulbecco's modified Eagle's Medium
DNA Deoxyribonucleic acid
CpG C-phosphate-G
ECM Extracellular Matrix
EGF Epidermal Growth Factor
EMA European Medicines Agency
ERK Extracellular-signal-Regulated Kinase
FACS Fluorescence-Activated Cell Sorting
FDA Food and Drug Administration
FITC Fluorescein
FSH Forward Scatter
GSK-3β Glycogen Synthase Kinase-3β
GvHD Graft versus Host Disease
HDAC Histone Deacetylase
HG High Glucose
HGF Hepatocyte Growth Factor
HLA Human Leukocyte Antigen
HS Horse Serum
HSCs Hematopoietic Stem Cells
IGF Insulin-like Growth Factor
iPSCs Induced Pluripotent Stem Cells
ISCT International Society for Cellular Therapy
ITS Insulin-Transferrin-Selenium
LG Low Glucose
MACS Magnetic-Activated Cell Sorting
MAPK Mitogen-Activated Protein Kinase
MSCs Mesenchymal Stem/Stromal Cells
MHC Myosin Heavy Chain
MRFs Myogenic Regulatory Factors
Muse Multilineage-differentiating Stress-Enduring
MyoG Myogenin
PE Phyroerythrin
PCR Polymerase Chain Reaction
RNA Ribonucleic Acid
SDS Sodium Dodecyl Sulfate
SMCs Smooth Muscle Cells
SM-MHC Smooth Muscle Myosin Heavy Chain
SSH Side Scatter
SRF Serum Response Factor
SUI Stress Urinary Incontinence
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SVF Stromal Vascular Fraction
TGF-β Transforming Growth Factor-β
UCM-MSCs Umbilical Cord Matrix Stem Cells or Umbilical Cord-derived MSCs
UI Urinary Incontinence
3-D Three-Dimensional
5-AZA 5- azacytidine
5-AZAd 5-aza-2’-deoxycytidine
α-SMA Smooth Muscle Alpha Actin
1
I. INTRODUCTION I.1 Background
I.1.1 Urinary Incontinence
Urinary incontinence (UI) affects more than 200 million people worldwide.1,2 Although not
life-threatening, it is a common medical and social condition that dramatically reduces the quality of life
of patients. In the United States of America, the total annual cost of UI and associated conditions was
estimated to range up to 32 billion dollars in 2006.2,3 Stress urinary incontinence or SUI is the most
prevalent type of UI.4 According to the International Continence Society, SUI is the complaint of
involuntary leakage of urine from the urethra on effort or exertion, or on sneezing or coughing.1,5 SUI
affects between 16% and 35% of adult women and the occurrence of this condition rises with
increasing age.4,6,7 Although more prevalent in woman, SUI can affect men, mostly due to prostate
surgery.2
Still, many incontinent patients do not report symptoms for various reasons, including
embarrassment or fear of treatment, which might indicate that the prevalence of UI may be much
higher than documented.1,6 Disguised UI is the cause of decreased self-esteem, neuroses, depression
states and difficulties in sex life, leading to a profound impact on the psychological condition of the
patient.8 The risk factors for the development of SUI include age, vaginal deliveries, genetic load
Myogenin 84 6.8 90 9.0 0 - 0.30 2.0 Figure III.18. Expression of myogenic markers in P0 cells cultured on gelatin- and fibronectin-coated plates (HG) after 12 days
in culture. Flow cytometry dot plot charts and histograms of Pax7, MyoD and myogenin (left to right) of the main population
gated of cells cultured on fibronectin- (HG) (A) and gelatin-coated plates (B). (C) Myogenic marker expression (in percentage
and MFI values) of the gated main populations and subpopulations of both conditions.
After induction, the morphology of some cells was changed. These cells became broad and
flattened, and kept increasing in proportion with culture time. After 8 days of medium replacement,
three types of cells could be visualized: (a) spindle-like cells, usually arranged in an aligned fashion,
(b) spindle-like cells with several vacuoles and (c) broad, flattened cells with enlarged nuclei
(Figure III.19A). The presence of enlarged cells, with significantly decreased nucleocytoplasmic ratios
when compared to spindle-like cells, was confirmed by immunofluorescence (Figure III.19A and B).
Importantly, MyoD expression was reduced in cells with enlarged nuclei and was more intense around
confluent spindle-like cells, which suggests that flattened enlarged cells lost myogenic marker
expression. The accumulation of vacuolar structures similar to lipid droplets with increased culture
time might be related to spontaneous adipogenic differentiation, which has been correlated with
plating ADSCs and satellite cells in DMEM-HG (i.e. 4.5 g/l glucose).173,174 The frequency of these
structures depended on the plating conditions (approximately 40% for SVF cells, 25% for cells
cultured on non-coated plates, 10-15% on fibronectin-coated (LG), 5% on fibronectin-coated (HG) and
5-10% on gelatin-coated plates).
A
B
C
54
Figure III.19. Cells displaying distinct morphologies after 8 days of induction, detected by light microscopy (top) and
fluorescence microscopy (bottom). (A) Cells cultured on fibronectin-coated plates (LG), in which aligned spindle-like and broad
flattened cells can be discerned; (B) SVF cells, in which the development of several vacuoles is visible; (C) cells cultured on
gelatin-coated plates and (D) cells cultured on non-coated plates; Broad flattened cells with enlarged nuclei (frequently
binucleated) could be seen in all conditions, as well as smaller spindle-like cells. Nuclei were stained with DAPI and the
cytoskeleton (F-actin filaments) was stained with phalloidin-TRITC.
Similar dot plots were obtained for all conditions after medium replacement. However, the
marker expression results indicate that changing the plating medium to DMEM+2%HS induced drastic
alterations in myogenin and MyoD expression (Table III.7). At day 2, although MyoD expression was
maintained, myogenin expression decreased drastically for cells on gelatin and fibronectin-coated
(HG) plates when compared to the expression before induction (Figure III.18). These results were
confirmed by immunofluorescence, in which MyoD expression was detected in all tested conditions,
while myogenin and Pax7 were not. Surprisingly, MyoD was detected not only in the nucleus but also
in the cytoplasm of the cells (Figure III.20).
At day 4 after medium replacement, the results regarding marker expression were similar to
day 2. However, the flow cytometry results obtained for day 8 show a drastic reduction in MyoD
percentage levels. A reduction in MyoD-expressing cells was also observed in the
immunofluorescence results (Figure III.21). It should be noted that due to a stock rupture, at day 4 the
secondary antibody used for myogenin had to be switched (from Alexa 488 to PE), and at day 8 for
MyoD (also from Alexa 488 to PE). Therefore, it is possible that this switch in the fluorophore used
influenced the results.
At day 8 skeletal MHC was introduced in the panel in order to identify terminally differentiated
myoblasts. The results indicate that gelatin coating was the condition that promoted myogenesis the
most, since almost 34% of cells expressed skeletal MHC. Surprisingly, SVF cells cultured on
non-coated plates displayed the second highest level of MHC expression (6.5%), followed by
A B
C D
55
positively sorted cells cultured on uncoated plates (3.3%). In accordance, phalloidin staining allowed
the visualization of aligned nuclei and some fused cells (Figure III.21). Interestingly, round
multinucleated cells (P1) were identified by DAPI and phalloidin staining in a very low confluence area
of a gelatin-coated well. No Pax7 and myogenin expression were observed by immunofluorescence
throughout the assay.
Table III.7. Expression of myogenic markers (Pax7, MyoD, myogenin and skeletal MHC) in P1 cells after 2, 4 and 8 days of
differentiation induction, assessed by flow cytometry. Secondary antibodies: Goat-anti mouse/rabbit Alexa 488 for myogenin
(day 2) and MyoD (days 2 and 4), respectively; goat-anti mouse/rabbit PE for myogenin (days 4 and 8) and MyoD (day 8).
Day 2
SVF No coating Gelatin Fibronectin (LG) Fibronectin (HG)
% MFI % MFI % MFI % MFI % MFI
Pax7 3.6 1.3 0.70 1.3 1.8 1.4 0.20 1.2 0.7 1.4
MyoD 99 18 99 44 99 24 99 24 99 28
Myogenin 2.8 1.7 2.8 1.8 6.8 1.8 2.4 1.6 1.1 1.6
Day 4
SVF No coating Gelatin Fibronectin (LG) Fibronectin (HG)
% MFI % MFI % MFI % MFI % MFI
Pax7 0.50 1.3 0.10 1.1 0.70 1.2 0.70 1.3 2.4 1.4
MyoD 98 19 99 20 99 23 99 20 100 21
Myogenin 0.70 1.5 0.60 1.4 0.70 1.3 0.8 1.4 0 -
Day 8 (%)
SVF No coating Gelatin Fibronectin (LG) Fibronectin (HG)
Pax7 0 0.10 0 0.20 0.10
MyoD 10 7.4 33 6.8 35
Myogenin 1.0 0.10 0 0.10 0.60
Skeletal MHC 6.5 3.3 34 1.2 2.4
Figure III.20. Multinucleated cells (P1) present in a very low confluence area of a gelatin-coated well, cultured for 17 days,
detected by DAPI and Phalloidin-TRITC staining.
56
Figure III.21. MyoD expression assessed by immunofluorescence after 2, 4 and 8 days of medium replacement for all conditions (secondary antibody: goat anti-rabbit Alexa 546). From day 4 to day 8 a decrease in the fluorescence intensity and the number of fluorescent cells was seen. Phalloidin-TRITC and DAPI staining (bottom row) allowed the observation of aligned nuclei and some fused
cells. No Pax7 and myogenin expression was observed through the assay for all the conditions. Scale bar = 50 μm.
D4 M
yo
D
D8 M
yo
D
D2 M
yo
D
D8 F
-Acti
n
SVF No Coating Gelatin Fibronectin (LG) Fibronectin (HG)
57
III.2.2.4 CD34/CD56 decay
The results obtained for the experiments performed in the sections above suggest that both
unsorted SVF cells and CD34-enriched cells obtained by MACS tend to lose CD34 expression once
plated. It was hypothesized that this loss of expression was due to downregulation of CD34
expression rather than death of CD34+ cells. To confirm this hypothesis, cells from unsorted SVF and
magnetically-sorted CD34 cells were plated at 10 000 cells/cm2 in expansion medium and in a
commercially available formulation of skeletal muscle growth medium, to test if a skeletal-specific
medium would help the maintenance of a CD34+CD56+ phenotype. CD34 and CD56 expression was
then measured by flow cytometry 2 and 3 days after the cells were plated (Figure III.22). For this
assay, the number of sorted cells in the CD34 positive fraction was not sufficient to test both media,
therefore, the cells were only plated in expansion medium. At day 2, only 330 events in total were
obtained for CD34-sorted cells. Two populations could be discerned (Subpopulation 1: 98% CD34+,
0.84% CD56+; Subpopulation 2: 54% CD34+, 4.8% CD56+), however; this data was not taken into
consideration since it was not possible to perform a statistically relevant analysis.
At day 2 after plating, a population of cells placed at low FSH and SSH levels was obtained for
unsorted SVF cells. This population was positive for CD34 in both tested medias (over 89% and 86%
of cells were CD34+ when plated in skeletal and expansion medium, respectively). CD56 expression,
on the other hand, was more dependent on the plating medium, as 36% of the cells plated in skeletal
media were CD56+, while only 9.3% of cells plated in expansion medium were CD56+.
At day 3 after plating, the formation of two distinct cell populations was evident in the flow
cytometry dot plots for unsorted SVF cells (plated in both media) and sorted cells (plated in expansion
medium). Both populations presented different phenotypes especially in terms of CD56 expression.
The newly formed cell population, placed at higher FSH and SSH values (Population 1), was positive
for CD34 and presented some CD56 expression, while Population 2 presented lower CD34
expression and was negative for CD56 (0% expression for all conditions). It can be hypothesized that
between day 2 and 3 of culture the adhered cells started elongating, and thus a new population of
cells with higher complexity and size developed. Surprisingly, this new population of cells
(Population 1) presented increased CD34 expression when compared to cells plated for 2 days, for
unsorted SVF cells. At day 3, slight differences between cells plated in the two different media could
be detected, for cells belonging to this population. In comparison to cells plated in expansion medium,
cells cultured using skeletal muscle medium had lower CD34 expression and higher CD56 expression.
Cells derived from the CD34-enriched fraction had higher CD34 and CD56 expression percentages
than unsorted SVF cells plated in both media.
The role of CD34 in SVF cells and ADSCs is currently unknown.175 These results indicate that
SVF cells that initially adhere to tissue culture plates are predominantly CD34+. However, AT-SVF
cells seem to progressively lose CD34 expression when they start proliferating, or as a result of
increasing confluence and cell-to-cell contact. This hypothesis would explain why significantly lower
CD34 expression was detected for unsorted SVF cells after 11 days in culture (Figure III.17)
58
SVF – Day 3 CD34-enriched – Day 3
Skeletal Muscle Medium Expansion Medium Expansion Medium
for the generation of acellular urethral bioscaffold should be developed.
To this end, four urethras removed from euthanized female pigs were decellularized through a
mechanochemical process based on a protocol developed by Simões and colleagues (Simões et al.,
under revision 2015). A dynamic system was implemented in which cell removal was promoted by the
constant flow of a mild detergent solution inside of the urethral tissue through perfusion for 4 days.
Simultaneously, agitation (340 rpm) on the outer surface of the tissue was induced. Distilled water was
perfused for 24 hours in the beginning and at the end of the process, which lasted for 6 days
(Figure III.28).
The protocol adopted herein allowed efficient cell removal, which could be visualized by the
gradual whitening of the tissue. Cell removal was not completely successful in urethra #3, in which
some brown coloured tissue could be observed at the distal end (external extremity) of the urethra.
This was most likely caused by the urethra size, since the urethra was too long when compared to the
catheter to allow for successful perfusion in the mentioned extremity. The protocol duration was
exceptionally increased to 10 consecutive days in order to allow maximum cell removal without
compromising the tissue and ECM structure. In general, this protocol proved to be robust, reproducible
and inexpensive, since only one reagent (SDS) is required, and peristaltic pumps are common
equipment in research facilities that work with bioreactors.
Decellularization is an attractive technique for urethral scaffold generation not only because of
the possibility of retaining the architecture of the native tissue, including the vasculature, but also due
to the presence of biofactors and appropriate mechanical structure in the ECM that are required for
cell adhesion and proliferation.12,30 However, the field of urinary tract organ tissue engineering has
been poorly studied, in particular in areas such as the development of urethral decellularization
protocols for scaffold production, which are currently lacking in the literature. Still, bladder
decellularization strategies have been established for tissue engineering purposes204–206. These can
eventually be useful for clinical trials or studies aiming the treatment of bladder-related incontinence,
such as overactive bladder. SUI, in particular, requires the development of strategies aiming urethra -
and not bladder - decellularization, such as the protocol tested herein.
Severe incontinence cases, caused by damages in the urethral tissue, would greatly benefit from
studies performed in adequate in vitro urethral models, in particular for the study of the
rhabdosphincter, which is essential for the continence mechanism. In this sense, this protocol has
71
several applications, the most straightforward being the development of in vitro models of SUI for
studies involving drug testing, cell seeding strategies, cell-based therapies and urethral engineering.
Figure III.28. (A) Schematics of the decellularization apparatus based on mechanochemical action. Briefly, the detergent
solution (or distilled water) is placed inside the (a) vessel where the (b) cannulated urethra is submerged. The solution is
pumped by a (c) peristaltic pump and re-circularized for 24 hours at a rate of 40 ml/min. Simultaneously, through a (d and e)
magnetic stirrer plate the solution was agitated at 340 rpm. After 48 hours, the direction of perfusion was changed (from the
black to the grey arrows). Adapted from Simões et al., 2015 (under revision). (B) Length and protocol duration for each
decellularized urethra. (C) Main steps of the decellularization process and their outcomes for urethra #4. Biological tissue
preparation includes AT and distal extremity removal, catheter placement and immobilization with sutures (all performed in
day 0). Gradual whitening of the tissue over perfusion time occurred, indicating efficient cell removal.
Urethra # Length
(cm) Protocol
duration (days)
1 2 6
2 5 6
3 8.5 10
4 9 6
A B
C
72
73
IV. Future Trends and Conclusions
Skeletal muscle engineering has suffered from several setbacks that have precluded clinical
translation of research in this field. These setbacks arise from the inability to expand and maintain
skeletal muscle progenitors (satellite cells and myoblasts) in an undifferentiated state in vitro. This
MSc project targeted the establishment of effective myogenic differentiation protocols for MSCs for
urethral sphincter reconstruction, with particular focus on exploring the myogenic potential of the SVF
from AT.
AT-derived cells are the most promising autologous cell source candidate for SUI cell therapy
or tissue engineering approaches owing to the following reasons: (i) ability to proliferate in vitro,
allowing for clinically relevant cell numbers to be obtained; (ii) clinical safety, with no reports of tumour
formation in the literature; (iii) Abundance and ease of harvest, without the need of resorting to urinary
tract surgery/biopsies; (iv) Intrinsic potential for committing to myogenic lineages, as proven herein
and by few other studies.69,110–113
When reviewing the current literature on MSC myogenic differentiation induction, the most
frequently adopted approach is, by far, 5-AZA supplementation to culture medium. However, as shown
here, the cytotoxic effects caused by this DNA demethylating agent should exclude its application in
Regenerative Medicine approaches. Moreover, contrasting data on its effect on cell differentiation
have been published, and no myogenesis induction was detected with the means used herein. Thus,
research on this field should progress towards selecting and expanding cells with intrinsic myogenic
potential. The results obtained here contribute to the development of this field of research, as they
indicate that (i) myogenic-committed cells are present in uncultured SVF and (ii) even if this
subpopulation is not efficiently selected, plated AT-SVF cells show intrinsic myogenic potential in the
presence of adequate stimuli (e.g. dexamethasone and bFGF), that can be enhanced in the presence
of gelatin coating, as indicated by the higher skeletal MHC expression levels obtained. These results
should be repeated, to account for biological variability, and the following alterations should be
implemented: (i) marker expression should be assessed by mRNA transcript quantification by
RT-PCR, in order to confirm the results by an antibody-independent method; (ii) primary antibodies
conjugated to fluorophores should be preferred; (iii) skeletal MHC expression should be included in
the marker panel and be analysed earlier in culture; (iv) the formulation of the myogenic induction
medium used should be optimized (e.g. DMEM-LG supplemented with HS, dexamethasone and
hydrocortisone or DMEM-HG supplemented with HS, insulin and transferrin); (v) marker expression
should be evaluated at different cell confluence levels to establish a possible correlation between cell
confluence and myogenesis induction. Furthermore, to attempt the isolation of Pax7+ cells from the
SVF, FACS for CD56 should be employed, as already discussed. Finally, the use of substrates with
stiffness values similar to skeletal muscle (10 kPa) might also be an interesting approach to further
stimulate myogenesis without resorting to chemical supplementation or serum.
74
Regarding the results obtained for the smooth muscle differentiation assays, these indicate
that chemical induction with PD98059 was not sufficient to promote a consistent increase in smooth
muscle marker expression in AT-SVF cells and BM-MSCs. In fact, the observations made in these
assays suggest that 3-D-like spatial organization might have promoted desmin and calponin
expression even in non-chemically stimulated cells. Therefore, 3-D stimuli in the form of patterned
substrates and ECM mimicking through the presence of coating should be explored, as well as the
influence of biomechanical cues (e.g. mechanical forces involved in the functioning of the internal
smooth muscle layer of the urethral sphincter).
Although little is still known about smooth muscle progenitors/precursors and the mechanisms
that lead to the differentiation of bladder SMCs, TGF-β is assumed to be important in the embryonic
development of bladder SMCs.42 On the other hand, consistent data on the literature point to this
molecule as an effective smooth muscle differentiation inducer in MSCs.45,131–133,144,188 Hence, TGF-β
might be a more promising candidate to study in further studies.
Finally, by combining this knowledge with cell expansion, urethra decellularization and cell
seeding techniques, an in vitro model to study SUI can potentially be developed, which would serve as
an important connecting tool between in vitro and clinical research.
75
V. References 1. Norton, P. & Brubaker, L. Urinary incontinence in women. Lancet 367, 57–67 (2006).
2. Lin, C. & Lue, T. Stem cell therapy for stress urinary incontinence: a critical review. Stem Cells Dev. 21,
834–843 (2012).
3. Levy, R. & Muller, N. Urinary incontinence: economic burden and new choices in pharmaceutical treatment. Adv. Ther. 23, 556–573 (2006).
4. Cerruto, M., D’Elia, C., Aloisi, A., Fabrello, M. & Artibani, W. Prevalence, Incidence and Obstetric Factors’ Impact on Female Urinary Incontinence in Europe: A Systematic. Urol. Int. 90, 1–9 (2012).
5. Abrams, P. et al. The standardisation of terminology of lower urinary tract function: report from the Standardisation Sub-committee of the International Continence Society. Neurourol. Urodyn. 21, 167–178
(2002).
6. Thaker, H. & Sharma, A. K. Regenerative medicine based applications to combat stress urinary incontinence. World J. Stem Cells 5, 112–123 (2013).
7. Stothers, L. & Friedman, B. Risk factors for the development of stress urinary incontinence in women. Curr. Urol. Rep. 12, 363–369 (2011).
8. Iwanowicz-Palus, G., Stadnicka, G. & Włoszczak-Szubzda, A. Medical and psychosocial factors conditioning development of stress urinary incontinence (SUI). Ann. Agric. Environ. Med. 20, 135–139
(2012).
9. Fry, C. H., Meng, E. & Young, J. S. The physiological function of lower urinary tract smooth muscle. Auton. Neurosci. 154, 3–13 (2010).
10. Thor, K. B. & Donatucci, C. Central nervous system control of the lower urinary tract: new pharmacological approaches to stress urinary incontinence in women. J. Urol. 172, 27–33 (2004).
11. DeLancey, J. Why do women have stress urinary incontinence? Neurourol. Urodyn. 29, S13–S17 (2010).
12. Orabi, H. et al. Tissue engineering of urinary bladder and urethra: advances from bench to patients. Sci. World J. 2013, (2013).
13. Brading, a F. Spontaneous activity of lower urinary tract smooth muscles: correlation between ion channels and tissue function. J. Physiol. 570, 13–22 (2006).
14. Mitterberger, M. et al. Adult stem cell therapy of female stress urinary incontinence. Eur. Urol. 53, 169–75
(2008).
15. Heesakkers, J. P. F. a & Gerretsen, R. R. R. Urinary incontinence: sphincter functioning from a urological perspective. Digestion 69, 93–101 (2004).
16. Bae, J. H. & Yoo, J. J. Cell-based therapy for urinary incontinence. Korean J. Urol. 51, 1–7 (2010).
17. Shokeir, A. a, Harraz, A. M. & El-Din, A. B. S. Tissue engineering and stem cells: basic principles and applications in urology. Int. J. Urol. 17, 964–973 (2010).
18. Smythe, G. M. & Grounds, M. D. Exposure to tissue culture conditions can adversely affect myoblast behavior in vivo in whole muscle grafts: implications for myoblast transfer therapy. Cell Transpl. 9, 379–
393 (2000).
19. Kinebuchi, Y. et al. Autologous bone-marrow-derived mesenchymal stem cell transplantation into injured rat urethral sphincter. Int. J. Urol. 17, 359–368 (2010).
20. Gregorio, S., García-Arranz, M., Olmo, D. & Pedrero, F. Phase II Clinical Trial to Study the Feasibility and Safety of the Expanded Autologous Mesenchymal Stem Cells Use Derived From Adipose Tissue (ASC) for the Local Feminine Stress Urinary Incontinence. (2012).
21. Hussain, M., Greenwell, T. J., Venn, S. N. & Mundy, A. R. The current role of the artificial urinary sphincter for the treatment of urinary incontinence. J. Urol. 174, 418–424 (2005).
22. Kaiser, L. R. The future of multihospital systems. Top. Health Care Financ. 18, 32–45 (1992).
23. Schmitt, A., van Griensven, M., Imhoff, A. B. & Buchmann, S. Application of stem cells in orthopedics. Stem Cells Int. 2012, (2012).
24. Fortier, L. a. Stem Cells: Classifications, Controversies, and Clinical Applications. Vet. Surg. 34, 415–423
(2005).
25. Hui, H. et al. in Stem Cells in Clinic and Research (ed. Gholamrezanezhad, A.) 3–15 (InTech, 2011), Available at http://www.intechopen.com/books/stem-cells-in-clinic-and-research/stem-cells- general-features-and-characteristics (Accessed on January 2015)
26. Takahashi, K. & Yamanaka, S. Induction of pluripotent stem cells from mouse embryonic and adult fibroblast cultures by defined factors. Cell 126, 663–676 (2006).
76
27. Sanchez-Ramos, J. R. Neural cells derived from adult bone marrow and umbilical cord blood. J. Neurosci. Res. 69, 880–893 (2002).
28. Pluripotent Stem Cells. Division of Biology and Medicine, Brown University, Available at http://biomed.brown.edu/Courses/BI108/BI108_2005_Groups/07/PluripotentStemCells.jpg (Accessed on November 2015)
29. Ren, G., Chen, X., Dong, F. & Li, W. Concise Review : Mesenchymal Stem Cells and Translational Medicine : Emerging Issues. Stem Cells Transl. Med. 1, 51–58 (2012).
30. Hwang, N. S., Varghese, S. & Elisseeff, J. Controlled differentiation of stem cells. Adv. Drug Deliv. Rev. 60, 199–214 (2008).
31. Bharadwaj, S. et al. Multipotential differentiation of human urine-derived stem cells: potential for therapeutic applications in urology. Stem Cells 31, 1840–56 (2013).
32. Chao, Y., et al. Umbilical cord-derived mesenchymal stem cells for hematopoietic stem cell transplantation. J. Biomed. Biotechnol. 2012, (2012).
33. Prochymal. Available at http://www.osiris.com/therapeutics.php# (Accessed on November 2015)
34. Bakker, A. & Langer, C. Cell-based therapies - an innovative therapeutic option in ophthalmology : Treating corneal diseases with stem cells. Fed. Heal. J. – Heal. Res. – Heal. Prot. 58, 1259–1264 (2015).
35. Carpenter, M. K., Frey-Vasconcells, J. & Rao, M. S. Developing safe therapies from human pluripotent stem cells. Nat. Biotechnol. 27, 606–613 (2009).
36. Kirouac, D. C. & Zandstra, P. W. The systematic production of cells for cell therapies. Cell Stem Cell 3,
369–81 (2008).
37. Bravery, C. a. et al. Potency assay development for cellular therapy products: An ISCT* review of the requirements and experiences in the industry. Cytotherapy 15, 9–19 (2013).
38. Hansmann, J., Groeber, F., Kahlig, A., Kleinhans, C. & Walles, H. Bioreactors in tissue engineering - principles, applications and commercial constraints. Biotechnol. J. 8, 298–307 (2013).
39. Macchiarini, P. et al. Clinical transplantation of a tissue-engineered airway. Lancet 372, 2023–2030
(2008).
40. Ikada, Y. Challenges in tissue engineering. J. R. Soc. Interface 3, 589–601 (2006).
41. Haisch, A., Kläring, S., Gröger, A., Gebert, C. & Sittinger, M. A tissue-engineering model for the manufacture of auricular-shaped cartilage implants. Eur. Arch. Otorhinolaryngol. 3, 316–321 (2002).
42. Tasian, G., Cunha, G. & Baskin, L. Smooth muscle differentiation and patterning in the urinary bladder. Differentiation. 80, 106–117 (2010).
43. Oostrom, O., Fledderus, J., de Kleijn, D., Pasterkamp, G. & Verhaar, M. Smooth Muscle Progenitor Cells: Friend or Foe in Vascular Disease? Curr. Stem Cell Res. Ther. 4, 131–140 (2009).
44. Groot, A. G. Smooth muscle cell origin and its relation to heterogeneity in development and disease. Arterioscler. Thromb. Vasc. Biol. 19, 1589–1594 (1999).
45. Rzucidlo, E. M., Martin, K. A. & Powell, R. J. Regulation of vascular smooth muscle cell differentiation. J Vasc Surg. 45, A25–A32 (2007).
46. Ross, J., Hong, Z., Willenbring, B. & Zeng, L. Cytokine-induced differentiation of multipotent adult progenitor cells into functional smooth muscle cells. J Clin Invest. 116, 3139–3149 (2006).
47. Bentzinger, C. F., Wang, Y. X. & Rudnicki, M. a. Building muscle: molecular regulation of myogenesis. Cold Spring Harb. Perspect. Biol. 4, 1–16 (2012).
48. Tajbakhsh, S. Skeletal muscle stem cells in developmental versus regenerative myogenesis. J. Intern. Med. 266, 372–389 (2009).
49. Tajbakhsh, S., Borello, U. & Vivarelli, E. Differential activation of Myf5 and MyoD by different Wnts in explants of mouse paraxial mesoderm and the later activation of myogenesis in the absence of Myf5. Development. 125, 4155–4162 (1998).
50. Caplan, A. Mesenchymal stem cells. J. Orthop. Res. 9, 641–50 (1991).
51. Bentzinger, C. F., Wang, Y. X., von Maltzahn, J. & Rudnicki, M. a. The emerging biology of muscle stem cells: implications for cell-based therapies. Bioessays 35, 231–241 (2013).
52. Péault, B. et al. Stem and progenitor cells in skeletal muscle development, maintenance, and therapy. Mol. Ther. 15, 867–877 (2007).
53. Asakura, A., Komaki, M. & Rudnicki, M. Muscle satellite cells are multipotential stem cells that exhibit myogenic, osteogenic, and adipogenic differentiation. Differentiation 68, 245–253 (2001).
54. Boonen, K. J. M. & Post, M. J. The muscle stem cell niche: regulation of satellite cells during regeneration. Tissue Eng. Part B. Rev. 14, 419–431 (2008).
77
55. Beauchamp, J. R. et al. Expression of CD34 and Myf5 defines the majority of quiescent adult skeletal muscle satellite cells. J. Cell Biol. 151, 1221–1233 (2000).
56. Capkovic, K. L., Stevenson, S., Johnson, M. C., Thelen, J. J. & Cornelison, D. D. W. Neural cell adhesion molecule (NCAM) marks adult myogenic cells committed to differentiation. Exp. Cell Res. 314, 1553–
1565 (2008).
57. Lazarus, H. M. et al. Cotransplantation of HLA-identical sibling culture-expanded mesenchymal stem cells and hematopoietic stem cells in hematologic malignancy patients. Biol. Blood Marrow Transplant. 11,
389–398 (2005).
58. Farini, A., Razini, P., Erratico, S., Torrente, Y. & Meregalli, M. Cell based therapy for Duchenne muscular dystrophy. J. Cell. Physiol. 221, 526–534 (2009).
59. Deschênes, I., Chahine, M., Tremblay, J., Paulin, D. & Puymirat, J. Increase in the proliferative capacity of human myoblasts by using the T antigen under the vimentin promoter control. Muscle and Nerve 20,
437–445 (1997).
60. Bouchentouf, M., Benabdallah, B. F., Rousseau, J., Schwartz, L. M. & Tremblay, J. P. Induction of Anoikis following myoblast transplantation into SCID mouse muscles requires the Bit1 and FADD pathways. Am. J. Transplant. 7, 1491–1505 (2007).
61. Caplan, A. Mesenchymal stem cells. J. Orthop. Res. 9, 641–650 (1991).
62. Gimble, J. M. et al. Adipose-derived stromal/stem cells: a primer. Organogenesis 9, 3–10 (2013).
63. Friedenstein, A., Chailakhjan, R. & Lalykina, K. The development of fibroblast colonies in monolayer cultures of guinea-pig bone marrow and spleen cells. Cell Tissue Kinet. 3, 393–403 (1970).
64. Augello, A., Kurth, T. & Bari, C. De. Mesenchymal stem cells: a perspective from in vitro cultures to in vivo migration and niches. Eur Cell Mater 20, 121–133 (2010).
65. Dominici, M. et al. Minimal criteria for defining multipotent mesenchymal stromal cells. The International Society for Cellular Therapy position statement. Cytotherapy 8, 315–317 (2006).
66. Bianco, P., Cao, X., Frenette, P. & Mao, J. The meaning, the sense and the significance: Translating the science of mesenchymal stem cells into medicine. Nat. Med. 19, 35–42 (2013).
67. Wakitani, S., Saito, T. & Caplan, A. I. Myogenic cells derived from rat bone marrow mesenchymal stem cells exposed to 5-azacytidine. Muscle Nerve 18, 1417–1426 (1995).
68. Choi, Y., Vincent, L. G., Lee, A. R., Dobke, M. K. & Engler, A. J. Mechanical derivation of functional myotubes from adipose-derived stem cells. Biomaterials 33, 2482–2491 (2012).
69. Zuk, P., Zhu, M. & Mizuno, H. Multilineage cells from human adipose tissue: implications for cell-based therapies. Tissue Eng. 7, 211–228 (2001).
70. Kolf, C. M., Cho, E. & Tuan, R. S. Mesenchymal stromal cells. Biology of adult mesenchymal stem cells: regulation of niche, self-renewal and differentiation. Arthritis Res. Ther. 9, 204–214 (2007).
71. Kuhn, N. Z. & Tuan, R. S. Regulation of stemness and stem cell niche of mesenchymal stem cells: implications in tumorigenesis and metastasis. J. Cell. Physiol. 222, 268–677 (2010).
72. Deans, R. J. & Moseley, A. B. Mesenchymal stem cells : Biology and potential clinical uses. Exp. Hematol. 28, 875–884 (2000).
73. Chamberlain, G., Fox, J., Ashton, B. & Middleton, J. Concise review: mesenchymal stem cells: their phenotype, differentiation capacity, immunological features, and potential for homing. Stem Cells 25,
2739–49 (2007).
74. Ryan, J., Barry, F., Murphy, J. & Mahon, B. Mesenchymal stem cells avoid allogeneic rejection. J. Inflamm. 2, (2005).
75. Pittenger, M. F. Multilineage Potential of Adult Human Mesenchymal Stem Cells. Science (80-. ). 284,
143–147 (1999).
76. Sekiya, I., Larson, B. L., Smith, J. R. & Pochampally, R. Expansion of Human Adult Stem Cells from Bone Marrow Stroma: Conditions that Maximize the Yields of Early Progenitors and Evaluate Their Quality. Stem Cells 20, 530–541 (2002).
77. Yang, B., Zheng, J. & Zhang, Y. Myogenic differentiation of mesenchymal stem cells for muscle regeneration in urinary tract. Chin. Med. J. (Engl). 126, 2952–2959 (2013).
78. Lu, L., Liu, Y., Yang, S. & Zhao, Q. Isolation and characterization of human umbilical cord mesenchymal stem cells with hematopoiesis-supportive function and other potentials. Haematol. Hematol. J. 91, 1017–
1026 (2006).
79. Caplan, A. I. & Dennis, J. E. Mesenchymal stem cells as trophic mediators. J. Cell. Biochem. 98, 1076–84
(2006).
80. Caplan, A. & Correa, D. The MSC : An Injury Drugstore. Cell Stem Cell 9, 11–15 (2011).
78
81. Bronckaers, A. et al. Mesenchymal stem / stromal cells as a pharmacological and therapeutic approach to accelerate angiogenesis. Pharmacol. Ther. 143, 181–196 (2014).
82. Blanc, K. Le, Frassoni, F., Ball, L. & Locatelli, F. Mesenchymal stem cells for treatment of steroid-resistant, severe, acute graft-versus-host disease: a phase II study. Lancet 371, 1579–86 (2008).
83. Seo, S. & Na, K. Mesenchymal stem cell-based tissue engineering for chondrogenesis. J. Biomed. Biotechnol. 2011, (2011).
84. Ankrum, J. & Karp, J. M. Mesenchymal stem cell therapy: Two steps forward, one step back. Trends Mol. Med. 16, 203–209 (2010).
85. Kuroda, Y., Kitada, M., Wakao, S. & Nishikawa, K. Unique multipotent cells in adult human mesenchymal cell populations. 107, 8639–8643 (2010).
86. Heneidi, S. et al. Awakened by Cellular Stress : Isolation and Characterization of a Novel Population of Pluripotent Stem Cells Derived from Human Adipose Tissue. PLoS One 8, (2013).
87. Wakao, S. et al. Multilineage-differentiating stress-enduring (Muse) cells are a primary source of induced pluripotent stem cells in human fibroblasts. Proc. Natl. Acad. Sci. U. S. A. 108, 9875–9880 (2011).
88. Ogura, F. et al. Human adipose tissue possesses a unique population of pluripotent stem cells with nontumorigenic and low telomerase activities: potential implications in regenerative medicine. Stem Cells Dev. 23, 717–728 (2014).
89. Kuroda, Y. & Dezawa, M. Mesenchymal Stem Cells and Their Subpopulation , Pluripotent Muse Cells , in Basic Research and Regenerative Medicine. Anat. Rec. 110, 98–110 (2014).
90. Yamauchi, T. et al. Therapeutic Effects of Human Multilineage-Differentiating Stress Enduring (MUSE) Cell Transplantation into Infarct Brain of Mice. PLoS One 10, (2015).
91. Kinoshita, K., Kuno, S., Ishimine, H. & Aoi, N. Therapeutic Potential of Adipose-Derived SSEA-3-Positive Muse Cells for Treating Diabetic Skin Ulcers. Stem Cells Transl. Med. 4, 146–155 (2015).
92. Grayson, W., Zhao, F., Izadpanah, R., Bunnell, B. & Ma, T. Effects of Hypoxia on Human Mesenchymal Stem Cell Expansion and Plasticity in 3D Constructs. J. Cell. Physiol. 207, 331–339 (2006).
93. Dos Santos, F. et al. Ex vivo expansion of human mesenchymal stem cells: A more effective cell proliferation kinetics and metabolism under hypoxia. J. Cell. Physiol. 223, 27–35 (2010).
94. Crisan, M. et al. A perivascular origin for mesenchymal stem cells in multiple human organs. Cell Stem Cell 3, 301–13 (2008).
95. Lin, G. et al. Defining Stem and Progenitor Cells within Adipose Tissue. Stem Cells Dev. 17, 1053–1063
(2008).
96. Caplan, A. I. All MSC are Pericytes? Cell Stem Cell 3, 229–230 (2008).
97. Mohyeldin, A., Garzón-Muvdi, T. & Quiñones-Hinojosa, A. Oxygen in stem cell biology: A critical component of the stem cell niche. Cell Stem Cell 7, 150–161 (2010).
98. Orabi, H., AbouShwareb, T., Zhang, Y., Yoo, J. J. & Atala, A. Cell-seeded tubularized scaffolds for reconstruction of long urethral defects: a preclinical study. Eur. Urol. 63, 531–8 (2013).
99. Raya-Rivera, A. et al. Tissue-engineered autologous urethras for patients who need reconstruction: an observational study. Lancet 377, 1175–82 (2011).
100. Yamamoto, T. et al. Periurethral injection of autologous adipose-derived regenerative cells for the treatment of male stress urinary incontinence: Report of three initial cases. Int. J. Urol. 19, 652–659
(2012).
101. Aicher, W. et al. Towards a Treatment of Stress Urinary Incontinence: Application of Mesenchymal Stromal Cells for Regeneration of the Sphincter Muscle. J. Clin. Med. 3, 197–215 (2014).
102. Gang, E., Jeong, J., Hong, S. & Hwang, S. Skeletal Myogenic Differentiation of Mesenchymal Stem Cells Isolated from Human Umbilical Cord Blood. Stem Cells 22, 617–624 (2004).
103. Gunetti, M. et al. Myogenic potential of whole bone marrow mesenchymal stem cells in vitro and in vivo for usage in urinary incontinence. PLoS One 7, (2012).
104. Beier, J. P. et al. Myogenic differentiation of mesenchymal stem cells co-cultured with primary myoblasts. Cell Biol. Int. 35, 397–406 (2011).
105. Garza-Rodea, A. S. De et al. Myogenic Properties of Human Mesenchymal Stem Cells Derived From Three Different Sources. Cell Transplant. 21, 153–173 (2012).
106. Belanto, J. J. et al. Dexamethasone induces dysferlin in myoblasts and enhances their myogenic differentiation. Neuromuscul Disord. 20, 111–121 (2010).
107. Lee, E. a., Im, S.-G. & Hwang, N. S. Efficient myogenic commitment of human mesenchymal stem cells on biomimetic materials replicating myoblast topography. Biotechnol. J. 9, 1604–1612 (2014).
79
108. Bourin, P. et al. Stromal cells from the adipose tissue-derived stromal vascular fraction and culture expanded adipose tissue-derived stromal/stem cells: A joint statement of the International Federation for Adipose Therapeutics and Science (IFATS) and the International So. Cytotherapy 15, 641–648 (2013).
109. Bourin, P. et al. Myogenic Differentiation by Human Processed Lipoaspirate Cells. Cytotherapy 15, 641–
648 (2013).
110. Di Rocco, G. et al. Myogenic potential of adipose-tissue-derived cells. J. Cell Sci. 119, 2945–2952 (2006).
111. Shan, T. et al. Distinct populations of adipogenic and myogenic Myf5-lineage progenitors in white adipose tissues. J. Lipid Res. 54, 2214–24 (2013).
112. Mizuno, H. et al. Myogenic Differentiation by Human Processed Lipoaspirate Cells. Plast Reconstr Surg. 109, 199–209 (2002).
113. Dromard, C. et al. Mouse adipose tissue stromal cells give rise to skeletal and cardiomyogenic cell sub-populations. Front. cell Dev. Biol. 2, (2014).
114. Zhu, B. & Murthy, S. K. Stem cell separation technologies. Curr. Opin. Chem. Eng. 2, 3–7 (2013).
115. Diogo, M. M., da Silva, C. L. & Cabral, J. M. S. Separation technologies for stem cell bioprocessing. Biotechnol. Bioeng. 109, 2699–2709 (2012).
116. FACS. Midlands Technical College Webpage. Available at http://classes.midlandstech.edu/carterp/Courses/bio225/chap18/figure_18_12_labeled.jpg (Accessed on November 2015)
117. MACS. Humboldt University of Berlin Webpage. Available at http://edoc.hu-berlin.de/dissertationen/hajkova-petra-2002-09-16/HTML/hajkova_html_m271ca532.gif (Accessed on November 2015)
118. Miltenyi Biotec Web Page - CliniMACS® System. Available at http://www.miltenyibiotec.com/en/clinical-applications/clinimacs-system.aspx (Acessed on November 2015)
119. Grützkau, A. & Radbruch, A. Small but mighty: How the MACS1-technology based on nanosized superparamagnetic particles has helped to analyze the immune system within the last 20 years. Cytom. Part A 77, 643–647 (2010).
120. Pliml, J. & Šorm, F. Synthesis of a 2-deoxy-d-ribofuranosyl-s-azacytosine. Collect. Czechoslov. Chem. Commun. 29, 2576–2578 (1964).
121. Čihák, A., Veselý, J. & Šorm, F. Incorporation of 5-azacytidine into liver ribonucleic acids of leukemic mice sensitive and resistant to 5-azacytidine. Biochim. Biophys. Acta 108, 516–518 (1965).
122. Périgaud, C., Gosselin, G. & Imbach, J. L. Nucleosides and Nucleotides Nucleoside Analogues as Chemotherapeutic Agents : A Review. Nucleosides and Nucleotides 11, 903–945 (1992).
123. Christman, J. K. 5-Azacytidine and 5-aza-2 ’-deoxycytidine as inhibitors of DNA methylation : mechanistic studies and their implications for cancer therapy. Oncogene 21, 5483–5495 (2002).
124. Meligy, F. Y. et al. The efficiency of in vitro isolation and myogenic differentiation of MSCs derived from adipose connective tissue, bone marrow, and skeletal muscle tissue. Vitr. Cell. Dev. Biol. 48, 203–215
(2012).
125. Song, M., Kim, H., Choi, Y., Kim, K. & Chung, C. Skeletal myogenic differentiation of human periodontal ligament stromal cells isolated from orthodontically extracted premolars. Korean J. Orthod. 42, 249–254
(2012).
126. Drost, A. C. et al. In vitro Myogenic Differentiation of Human Bone Marrow – Derived Mesenchymal Stem Cells as a Potential Treatment for Urethral Sphincter Muscle Repair. Ann. N. Y. Acad. Sci. 1176, 135–143
(2009).
127. Tomita, S., Li, R., Weisel, R. D., Mickle, D. A. G. & Kim, E. Autologous Transplantation of Bone Marrow Cells Improves Damaged Heart Function. Circulation 100, 247–256 (1999).
128. Makino, S. et al. Cardiomyocytes can be generated from marrow stromal cells in vitro. J. Clin. Invest. 103,
697–705 (1999).
129. Zhang, Y., Chu, Y., Shen, W. & Dou, Z. Effect of 5-azacytidine induction duration on differentiation of human first-trimester fetal mesenchymal stem cells towards cardiomyocyte-like cells. Interact. Cardiovasc. Thorac. Surg. 9, 943–946 (2009).
130. Zhou, G.-S., Zhang, X.-L., Wu, J.-P. & Zhang, R.-P. 5-Azacytidine facilitates osteogenic gene expression and differentiation of mesenchymal stem cells by alteration in DNA methylation. Cytotechnology 60, 11–
22 (2009).
131. Fu, Q., Song, X.-F., Liao, G.-L., Deng, C.-L. & Cui, L. Myoblasts differentiated from adipose-derived stem cells to treat stress urinary incontinence. Urology 75, 718–723 (2010).
132. Rosca, A. & Burlacu, A. Effect of 5-Azacytidine : Evidence for Alteration of the Multipotent Ability of Mesenchymal Stem Cells. Stem Cells Dev. 20, 1213–1221 (2011).
80
133. Liu, Y. et al. Growth and differentiation of rat bone marrow stromal cells : does 5-azacytidine trigger their cardiomyogenic differentiation ? Cardiovasc. Res. 58, 460–468 (2003).
134. Martin-Rendon, E. et al. 5-Azacytidine-treated human mesenchymal stem/progenitor cells derived from umbilical cord, cord blood and bone marrow do not generate cardiomyocytes in vitro at high frequencies. Vox Sang. 95, 137–148 (2008).
135. McKee, J. A. et al. Human arteries engineered in vitro. EMBO Rep. 4, 633–638 (2003).
136. Galmiche, B. M. C. et al. Stromal Cells From Human Long-Term Marrow Cultures Are Mesenchymal Cells That Differentiate Following a Vascular Smooth Muscle Differentiation Pathway. Blood 82, 66–76 (1993).
137. Iwase, T. et al. Comparison of angiogenic potency between mesenchymal stem cells and mononuclear cells in a rat model of hindlimb ischemia. Cardiovasc. Res. 66, 543–551 (2005).
138. Sinha, S. et al. Transforming growth factor- beta 1 signaling contributes to development of smooth muscle cells from embryonic stem cells. Am. J. Physiol. cell Physiol. 22908, 1560–1568 (2004).
139. Kinner, B., Zaleskas, J. M. & Spector, M. Regulation of Smooth Muscle Actin Expression and Contraction in Adult Human Mesenchymal Stem Cells. Exp. Cell Res. 278, 72–83 (2002).
140. Tamama, K., Sen, C. K. & Wells, A. Differentiation of bone marrow mesenchymal stem cells into the smooth muscle lineage by blocking ERK/MAPK signaling pathway. Stem Cells Dev. 17, 897–908 (2008).
141. Gong, Z. & Niklason, L. E. Small-diameter human vessel wall engineered from bone marrow-derived mesenchymal stem cells (hMSCs). FASEB J. 22, 1635–48 (2008).
142. Tian, H. et al. Myogenic differentiation of human bone marrow mesenchymal stem cells on a 3D nano fibrous scaffold for bladder tissue engineering. Biomaterials 31, 870–877 (2010).
143. Wang, D. et al. Proteomic profiling of bone marrow mesenchymal stem cells upon transforming growth factor beta1 stimulation. J. Biol. Chem. 279, 43725–34 (2004).
144. Bajpai, V. K., Mistriotis, P., Loh, Y.-H., Daley, G. Q. & Andreadis, S. T. Functional vascular smooth muscle cells derived from human induced pluripotent stem cells via mesenchymal stem cell intermediates. Cardiovasc. Res. 96, 391–400 (2012).
145. Nishimoto, S. & Nishida, E. MAPK signalling: ERK5 versus ERK1/2. EMBO Rep. 7, 782–786 (2006).
146. Kim, H. J. & Bar-Sagi, D. Modulation of signalling by Sprouty: a developing story. Nat. Rev. Mol. Cell Biol. 5, 441–450 (2004).
147. Wang, Z. et al. Myocardin and ternary complex factors compete for SRF to control smooth muscle gene expression. Nature 428, 185–189 (2004).
148. Van Tuyn, J. et al. Activation of cardiac and smooth muscle-specific genes in primary human cells after forced expression of human myocardin. Cardiovasc. Res. 67, 245–255 (2005).
149. Alimperti, S., You, H., George, T., Agarwal, S. K. & Andreadis, S. T. Cadherin-11 regulates both mesenchymal stem cell differentiation into smooth muscle cells and the development of contractile function in vivo. J. Cell Sci. 127, 2627–38 (2014).
150. Wang, T., Xu, Z., Jiang, W. & Ma, A. Cell-to-cell contact induces mesenchymal stem cell to differentiate into cardiomyocyte and smooth muscle cell. Int. J. Cardiol. 109, 74–81 (2006).
151. Jang, J., et al. Combined effects of surface morphology and mechanical straining magnitudes on the differentiation of mesenchymal stem cells without using biochemical reagents. J. Biomed. Biotechnol. 2011, 1–9 (2011).
152. Liu, J., Wang, Y., Wu, Y., Ni, B. & Liang, Z. Sodium Butyrate Promotes the Differentiation of Rat Bone Marrow Mesenchymal Stem Cells to Smooth Muscle Cells through Histone Acetylation. PLoS One 9,
(2014).
153. Parmar, N., Ahmadi, R. & Day, R. M. A Novel Method for Differentiation of Human Mesenchymal Stem Cells into Smooth Muscle-Like Cells on Clinically Deliverable Thermally Induced Phase Separation Microspheres. Tissue Eng. Part C. Methods 00, 1–9 (2014).
154. Shah, F. S., Wu, X., Dietrich, M., Rood, J. & Gimble, J. M. A non-enzymatic method for isolating human adipose tissue-derived stromal stem cells. Cytotherapy 15, 979–985 (2013).
155. Butler, M. Animal Cell Culture and Technology. (Garland Science/BIOS Scientific Publishers, 2005).
156. Pisani, D. F. et al. Hierarchization of myogenic and adipogenic progenitors within human skeletal muscle. Stem Cells 28, 2182–2194 (2010).
157. Pisani, D. F. et al. Isolation of a Highly Myogenic CD34-Negative Subset of Human Skeletal Muscle Cells Free of Adipogenic Potential. Stem Cells 28, 753–764 (2010).
158. Pilz, G. a et al. Human mesenchymal stromal cells express CD14 cross-reactive epitopes. Cytometry. A 79, 635–45 (2011).
81
159. Basu, J. et al. Expansion of the human adipose-derived stromal vascular cell fraction yields a population of smooth muscle-like cells with markedly distinct phenotypic and functional properties relative to mesenchymal stem cells. Tissue Eng. Part C. Methods 17, 843–860 (2011).
160. Kundrotas, G. Surface markers distinguishing mesenchymal stem cells from fibroblasts. Acta medica Litu. 19, 75–79 (2012).
161. Christodoulou, I., Kolisis, F. N., Papaevangeliou, D. & Zoumpourlis, V. Comparative Evaluation of Human Mesenchymal Stem Cells of Fetal (Wharton’s Jelly) and Adult (Adipose Tissue) Origin during Prolonged In Vitro Expansion: Considerations for Cytotherapy. Stem Cells Int. 2013, (2013).
162. Neuhuber, B., Swanger, S. A., Howard, L., Mackay, A. & Fischer, I. Effects of plating density and culture time on bone marrow stromal cell characteristics. Exp. Hematol. 36, 1176–1185 (2008).
163. Vermes, I., Haanen, C., Steffens-Nakken, H. & Reutelingsperger, C. A novel assay for apoptosis. Flow cytometric detection of phosphatidylserine expression on early apoptotic cells using fluorescein labelled Annexin V. J. Immunol. Methods 184, 39–51 (1995).
164. Sinanan, A. C. M., Hunt, N. P. & Lewis, M. P. Human adult craniofacial muscle-derived cells: neural-cell adhesion-molecule (NCAM; CD56)-expressing cells appear to contain multipotential stem cells. Biotechnol. Appl. Biochem. 40, 25–34 (2004).
165. Yeh, S.-P. et al. Induction of CD45 expression on bone marrow-derived mesenchymal stem cells. Leukemia 20, 894–896 (2006).
166. Dupéré-Minier, G., Desharnais, P. & Bernier, J. Involvement of tyrosine phosphatase CD45 in apoptosis. Apoptosis 15, 1–13 (2010).
167. Konieczny, S. F. & Emerson, C. P. Induction of Stable Mesodermal Stem Cell Lineages from 10T1/2 Cells : Evidence for Regulatory Genes Controlling Determination. Cell 38, 791–800 (1984).
168. Bel, A. et al. Transplantation of autologous fresh bone marrow into infarcted myocardium: a word of caution. Circulation 108 Suppl , II247–52 (2003).
169. Andrade, P. Z., Lobato Da Silva, C., Dos Santos, F., Almeida-Porada, G. & Cabral, J. M. S. Initial CD34 + cell-enrichment of cord blood determines hematopoietic stem/progenitor cell yield upon ex vivo expansion. J. Cell. Biochem. 112, 1822–1831 (2011).
170. Zheng, B. et al. Prospective identification of myogenic endothelial cells in human skeletal muscle. Nat. Biotechnol. 25, 1025–1034 (2007).
171. Chen, C.-W., Corselli, M., Péault, B. & Huard, J. Human Blood-Vessel-Derived Stem Cells for Tissue Repair and Regeneration. J. Biomed. Biotechnol. 2012, 1–9 (2012).
172. Li, Y.-M. et al. Effects of high glucose on mesenchymal stem cell proliferation and differentiation. Biochem. Biophys. Res. Commun. 363, 209–215 (2007).
173. Aguiari, P. et al. High glucose induces adipogenic differentiation of muscle-derived stem cells. Proc. Natl. Acad. Sci. U. S. A. 105, 1226–31 (2008).
174. Vettor, R. et al. The origin of intermuscular adipose tissue and its pathophysiological implications. Am J Physiol Endocrinol Metab 987–998 (2009). doi:10.1152/ajpendo.00229.2009
175. Scherberich, A., Di Maggio, N. & McNagny, K. M. A familiar stranger: CD34 expression and putative functions in SVF cells of adipose tissue. World J Stem Cells 5, 1–8 (2013).
176. Bosnakovski, D., Xu, Z., Li, W. & Thet, S. Prospective Isolation of Skeletal Muscle Stem Cells with a Pax7 Reporter. Stem Cells 26, 3194–3204 (2008).
177. Sirabella, D., De Angelis, L. & Berghella, L. Sources for skeletal muscle repair: From satellite cells to reprogramming. J. Cachexia. Sarcopenia Muscle 4, 125–136 (2013).
178. Gimble, J. M., Katz, A. J. & Bunnell, B. A. Adipose-Derived Stem Cells for Regenerative Medicine. Circ. Res. 100, 1249–1260 (2007).
179. Baumgarth, N. & Roederer, M. A practical approach to multicolor flow cytometry for immunophenotyping. J. Immunol. Methods 243, 77–97 (2000).
180. Maumus, M. et al. Native human adipose stromal cells: localization, morphology and phenotype. Int. J. Obes. (Lond). 35, 1141–1153 (2011).
181. Grigoriadis, A. E., Heersche, J. N. M. & Aubin, J. E. Differentiation of muscle, fat, cartilage, and bone from progenitor cells present in a bone-derived clonal cell population: Effect of dexamethasone. J. Cell Biol. 106, 2139–2151 (1988).
182. Guerriero, V. & Florini, J. R. Dexamethasone effects on myoblast proliferation and differentiation. Endocrinology 106, 1198–1202 (1980).
183. Gates, G., Nandan, D., Brickenden, A. & Sanwal, B. Differentiation defective mutants of skeletal myoblasts altered in a gelatin-binding glycoprotein. Biochem Cell Biol. 65, 767–775 (1987).
82
184. Nusgens, B., Delain, D., Senechal, H. & Winand, R. Metabolic changes in the extracellular matrix during differentiation of myoblasts of the L6 line and of a myo- non-fusing mutant. Exp Cell Res. 161, 51–62
(1896).
185. Nandan, D., Clarke, E. P., Ball, E. & Sanwal, B. Ethyl-3,4-dihydroxybenzoate Inhibits Myoblast Differentiation: Evidence for an Essential Role of Collagen. Cell 110, 1673–1679 (1990).
186. Vaz, R., Martins, G. G., Thorsteinsdóttir, S. & Rodrigues, G. Fibronectin promotes migration, alignment and fusion in an in vitro myoblast cell model. Cell Tissue Res. 348, 569–578 (2012).
187. Chaturvedi, V. et al. Interactions between Skeletal Muscle Myoblasts and their Extracellular Matrix Revealed by a Serum Free Culture System. PLoS One 10, (2015).
188. Grzelkowska-Kowalczyk, K., Wieteska-Skrzeczyńska, W., Grabiec, K. & Tokarska, J. High glucose-mediated alterations of mechanisms important in myogenesis of mouse C2C12 myoblasts. Cell Biol. Int. 37, 29–35 (2013).
189. Munoz, J., Zhou, Y. & Jarrett, H. W. LG4-5 domains of laminin-211 binds alpha-dystroglycan to allow myotube attachment and prevent anoikis. J. Cell. Physiol. 222, 111–119 (2010).
190. Goudenege, S. et al. Enhancement of Myogenic and Muscle Repair Capacities of Human Adipose–derived Stem Cells With Forced Expression of MyoD. Mol. Ther. 17, 1064–1072 (2009).
191. Wang, C. et al. Small activating RNA induces myogenic differentiation of rat adipose-derived stem cells by upregulating MyoD. Int. braz j urol 41, 764–772 (2015).
192. Lingbeck, J. M., Trausch-Azar, J. S., Ciechanover, A. & Schwartz, A. L. Determinants of nuclear and cytoplasmic ubiquitin-mediated degradation of MyoD. J. Biol. Chem. 278, 1817–1823 (2003).
193. Choi, Y. S. et al. The alignment and fusion assembly of adipose-derived stem cells on mechanically patterned matrices. Biomaterials 33, 6943–51 (2012).
194. Schugar, R. C., Robbins, P. D. & Deasy, B. M. Small molecules in stem cell self-renewal and differentiation. Gene Ther. 15, 126–135 (2008).
195. Harris, L., Abdollahi, H., Zhang, P. & McIlhenny, S. Differentiation of Adult Stem Cells into Smooth Muscle for Vascular Tissue Engineering. J Surg Res 168, 306–314 (2011).
196. Zen-Bio Webpage. Available at http://www.zen-bio.com/products/cells/human-bladder-cells.php (Accessed on October 2015)
197. Beamish, J. A., He, P., Kottke-Marchant, K. & Marchant, R. E. Molecular Regulation of Contractile Smooth Muscle Cell Phenotype: Implications for Vascular Tissue Engineering. Tissue Eng. Part B Rev. 16, 467–491 (2010).
198. Rodríguez, L. V et al. Clonogenic multipotent stem cells in human adipose tissue differentiate into functional smooth muscle cells. Proc. Natl. Acad. Sci. U. S. A. 103, 12167–72 (2006).
199. Liu, Y., Deng, B., Zhao, Y., Xie, S. & Nie, R. Differentiated markers in undifferentiated cells: Expression of smooth muscle contractile proteins in multipotent bone marrow mesenchymal stem cells. Dev. Growth Differ. 55, 591–605 (2013).
200. Jaiswal, R., Jaiswal, N., Bruder, S. & Mbalaviele, G. Adult Human Mesenchymal Stem Cell Differentiation to the Osteogenic or Adipogenic Lineage Is Regulated by Mitogen-activated Protein Kinase. J. Cell. Physiol. 275, 9645–9652 (2000).
201. Kurpinski, K. et al. Transforming growth factor-beta and notch signaling mediate stem cell differentiation into smooth muscle cells. Stem Cells 28, 734–42 (2010).
202. Khani, M.-M., Tafazzoli-Shadpour, M., Goli-Malekabadi, Z. & Haghighipour, N. Mechanical characterization of human mesenchymal stem cells subjected to cyclic uniaxial strain and TGF-β1. J. Mech. Behav. Biomed. Mater. 43, 18–25 (2015).
203. Maul, T. M., Chew, D. W., Nieponice, A. & Vorp, D. A. Mechanical stimuli differentially control stem cell behavior: morphology, proliferation, and differentiation. Biomech. Model. Mechanobiol. 10, 939–953
(2011).
204. Rosario, D. J. et al. Decellularization and sterilization of porcine urinary bladder matrix for tissue engineering in the lower urinary tract. Regen. Med. 3, 145–156 (2008).
205. Consolo, F. et al. A dynamic distention protocol for whole-organ bladder decellularization: histological and biomechanical characterization of the acellular matrix. J. Tissue Eng. Regen. Med. 3, 145–156 (2013).
206. Liao, W. et al. Construction of ureteral grafts by seeding bone marrow mesenchymal stem cells and smooth muscle cells into bladder acellular matrix. Transplant. Proc. 45, 730–734 (2013).