Differentiation of Human Mesenchymal Stem/Stromal Cells … · Differentiation of Human Mesenchymal...

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.

Palavras-Chave

Incontinência Urinária de Esforço

Células Estaminais/Estromais Mesenquimais (ou Mesenquimatosas)

Fração Vascular Estromal

Diferenciação Miogénica

Células do Músculo Liso

Células do Músculo Esquelético

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Index

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.1.1 Urinary Incontinence ........................................................................................................... 1 I.1.2 Regenerative Medicine ........................................................................................................ 3 I.1.3 Muscle Cells ........................................................................................................................ 8 I.1.4 Mesenchymal Stem/Stromal Cells .................................................................................... 11 I.1.5 Tissue Engineering-derived Urethral Constructs in Clinical Trials .................................... 17

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.3 MULTILINEAGE DIFFERENTIATION ASSAYS.................................................................................... 28 II.4 IMMUNOPHENOTYPE CHARACTERIZATION .................................................................................... 28 II.5 5-AZA-2’-DEOXYCYTIDINE TREATMENT ........................................................................................ 29

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

II.7 SMOOTH MUSCLE DIFFERENTIATION INDUCTION ........................................................................... 31 II.8 IMMUNOFLUORESCENCE STAINING .............................................................................................. 31 II.9 INTRACELLULAR STAINING FOR FLOW CYTOMETRY ....................................................................... 32 II.10 DECELLULARIZATION OF PORCINE URETHRAS ........................................................................... 32

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

(collagen deficiency), estrogens deficiency, diabetes mellitus, pelvic surgery, chronic constipation,

chronic cough, obesity and heavy physical work.7,8

UI emerges from a malfunction of the lower urinary tract. The lower urinary tract comprises

two regions: the urinary bladder and the outflow tract (bladder neck, or base, and urethra - Figure I.1).9

Figure I.1. The lower urinary tract in women and men. From Fry et al., 2009.

Urine is produced in the kidneys and stored in the bladder while the lumen of the urethra is

tightly shut, occasionally opening for the voluntary elimination of urine through the urethra.2

Structurally, the bladder is composed mainly by smooth muscle and connective tissue.10 During urine

2

storage, the smooth muscle - detrusor - is relaxed, making the bladder a low pressure, high volume

reservoir. The reverse happens during micturition, when the muscle layer becomes active, making the

bladder a high pressure, contracting vesicle for urine expulsion.10

The urethra plays a key role in the continence mechanism. The urethra is a multilayered

tubular structure that consists of smooth muscle with intrafascicular connective tissue, a rich

submucosa (or lamina propria) with collagen fibers and microvascularization, and a lining epithelium

(or urothelium).11,12 Additionally, the presence of a circular omega-shaped striated muscle layer that

surrounds the smooth muscle, named rhabdosphincter or external urethral sphincter, in the

mid-urethra in humans, allows the normal urethral closure mechanism (Figure I.2).11

Figure I.2. Cross-section of the upper portion of a female sphincter. From Heesakkers and Gerretsen, 2004.

The combined actions of the mentioned tissues serve to create wall tension that compresses

the lumen closed, though the rhabdosphincter contributes the most for the continence mechanism.2,11

During urine storage, this layer is tonically active, becoming dynamically active during increases in

abdominal pressure and inactive during normal micturition.10 Similarly, urethral smooth muscle cells

(SMCs) exhibit tonic spontaneous contractile activity that allows urine to remain in the bladder against

the forces of gravity, while micturition is assured by the relaxation of the urethral smooth muscle.13

SUI may arise from malfunctions of the rhabdosphincter, when bladder pressure exceeds

urethral pressure, in the setting of sudden increases of intra-abdominal forces.1 In this way, leakage of

urine from the urethra is caused by functional and morphological defects of the rhabdosphincter (e.g. a

decrease in the density of the sphincter), which can arise, as mentioned before, from age-dependent

spontaneous apoptosis of the striated muscle cells or from childbirth, maternal injury, or surgical

interventions.14

The current treatments for SUI include non-invasive measures such as lifestyle changes (fluid

intake management), pelvic floor muscle training and pharmaceutical agents.11 However, since SUI is

fundamentally an anatomical problem, none of these measures fully corrects the underlying pathology,

nor are these satisfactorily efficient. Moreover, the unwanted side-effects of some drugs

3

(e.g. cardiovascular issues and orthostatic hypotension) are not adequate for the elderly population

and the expenses of new drugs often lead to discontinuation.1,6 On the other hand, the surgical

approach to SUI involves suspension operations, placement of slings to support the urethra and

periurethral and transurethral injection therapy, by injecting biomaterials (bulking agents) to increase

urethral coaptation and outlet obstruction.15 Still, these procedures give rise to several complications

and do not always lead to the correction of UI as they do not treat its pathophysiologic causes.6,14,16

Due to the limitations of current treatments, novel alternative approaches have been

considered, such as Regenerative Medicine strategies, with the objective of restoring and maintaining

the integrity and function of damaged/insufficient urethral muscle tissue.17 Furthermore, one of the

objectives of Regenerative Medicine therapy for SUI is the reconstruction of the sphincter muscle

itself, by resorting to stem cells.16 Clinical trials using injectable fibroblasts and muscle cells have been

somewhat successful, with the purpose of treating atrophy of the urethral submucosa and defects of

the rhabdosphincter, respectively.14 However, fibroblasts only act as a tissue bulking agent, through

the production of extracellular matrix (ECM), and the ex vivo preparation of muscle precursor cells

drastically reduces their myogenic potential in vivo.16,18 Animal studies suggest that mesenchymal

stem/stromal cells (MSCs) – focused in Section I.1.4 – display the potential to regenerate both muscle

and ganglion components of the sphincter, thus holding great potential for sphincter reconstruction.6,19

Importantly, a Phase II clinical trial aiming to test the therapeutic effects of the intralesional application

of ex-vivo expanded adipose-derived cells for female SUI was already initiated.20

Artificial sphincters have been tested on patients; however, these devices are associated with

a multitude of complications, including urethral edema and atrophy and erosion of the sphincter.6,21

Therefore, a combined strategy using cell therapy and proper scaffolds should be the ideal approach

to reconstruct a functional/contractile urethral sphincter. Apart from its therapeutic potential, such

construct would serve as a representative in vitro model of SUI, allowing researchers to test novel

therapies, including drugs (i.e. drug screening) or cell therapy, and their effects in terms of toxicology

and efficacy, before proceeding into clinical trials. Additionally, it would be important to better

understand how specific damages in urethral support give rise to improper functioning of the urethral

sphincter, consequently leading to incontinence, and the development of such research tool would

allow to answer several questions that remain unanswered about SUI.11

I.1.2 Regenerative Medicine

In 1992, the term “Regenerative Medicine” was introduced by Leland Kaiser, who predicted

that a new branch of medicine would develop to attempt to change the course of chronic diseases and

in many instances would regenerate tired and failing organ systems.22 Regenerative Medicine

comprises two main branches: cell therapy and tissue engineering (Figure I.3). Cell therapy aims to treat

diseases or regenerate injured tissues through the injection of a specific cell population.23 The cells

can either be isolated from the patient (autologous setting) or from donors (allogeneic setting). On the

other hand, tissue engineering aims to construct tissues or organs for implantation, through the

4

combined use of stem/progenitor/adult cells, three-dimensional (3-D) scaffolds and/or soluble

factors.23

Figure I.3. The two main strategies involving stem cells in Regenerative Medicine-related clinical applications. From Schmitt

et al., 2012.

I.1.2.1 Stem Cells

Stem cells are undifferentiated cells that are defined by their functional capacity to self-renew

as well as generate differentiated progeny.24,25 One example of a truly totipotent cell, capable of

generating all cell types of a complete organism (including the extraembryonic tissues) is the fertilized

egg, but also the cells that result from its first divisions.24 As the fertilized egg starts to asymmetrically

divide, giving rise to the blastocyst, the cells from the epyblast (the inner mass of the blastocyst) are

no longer totipotent, as they can only differentiate into cells from the three germ layers: mesoderm,

ectoderm and endoderm (Figure I.4).24 These embryonic stem cells are classified as pluripotent,

together with other cell types such as embryonic germ cells and embryonal carcinoma cells (isolated

from teratocarcinomas) and, more recently, induced pluripotent stem cells (iPSCs).25 As their

denomination suggests, iPSCs are obtained from committed cells that were artificially reprogrammed

into a pluripotent state. The original method, developed by Yamanaka and co-workers, consisted in

the transduction of skin cells with four genes (the pluripotency keepers Oct 3/4 and Sox2 and the

oncogenes c-Myc and Klf4).26

The following term in the cell potency hierarchy is “multipotent”, and it is attributed to cells

capable of differentiating into a limited range of cell types, within the same germ layer

(e.g. hematopoietic stem cells or HSCs).24,25 These are also denominated as progenitor cells. Lastly,

precursor cells such as angioblasts, hepatoblasts or myoblasts, are regarded as unipotent, as they

can only sustain one cell lineage or cell type.25 When referring to stem cells obtained from any

postnatal organism, the term “adult stem cell” is commonly applied in order to distinguish them from

embryonic stem cells.24 For instances, MSCs (further discussed in Section I.1.4) are multipotent cells

included in the adult stem cell group.

5

Stem cells in adult animals are extremely important to maintain tissue and organ mass during

normal cellular turnover, with each tissue having a resident population of stem cells.24 It is still not

clear if adult stem cells retain some plasticity and are able to go through a phenomenon called

“transdifferentiation”, which is described to occur when stem cells derived from one adult tissue

change into cellular phenotypes not normally found in that tissue.27 Although the legitimacy of this

phenomenon has been highly debated, some authors claim that adult stem cells from mature tissues

can be reprogrammed (“transdifferentiated”) to change into a completely distinct phenotype from that

normally found in that tissue.27

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

I.1.2.2 (Stem) Cell Therapy

Stem cells represent a promising cell therapy tool for Regenerative Medicine largely because

of their ability to self-renew and differentiate into many functional cell types.29 Ideally, a stem cell

therapy approach would comprise validated methods of stem cell isolation, proliferation and

differentiation. Once isolated, and under the appropriate experimental conditions, stem cells could

proliferate in order to attain clinically relevant numbers of undifferentiated cells.30 Subsequently, these

undifferentiated cells could be also induced into lineage-specific differentiation through the

implementation of defined in vitro differentiation protocols, generating the desired cell type.30 This

approach allows the use of autologous stem cells, which is advantageous in the clinical setting, as

6

they do not induce immune responses and/or rejection.31 However, tissue-resident stem cells are a

very small subpopulation of cells and they are difficult to isolate from most differentiated somatic cells

present in postnatal tissues and organs.31

Currently, the use of stem cells for cell therapy as a well established and validated procedure

is only performed in the treatment of hemato-oncological diseases. The transplantation of HSCs, one

of the most successful examples of cell therapy, has been practiced for decades for this purpose.23 In

bone marrow (BM) transplantation, HSCs are injected into the blood circulation of the recipient and

find their way to the BM by a cytokine-mediated phenomenon termed “homing”.23 In the last decade,

the use of cells derived from the umbilical cord blood ascended as a potential alternative cell source to

BM transplantation due to advantages such as painless extraction procedure and lower incidence of

graft-versus-host disease (GvHD).32

Besides BM transplantation that is fully implemented worldwide, some examples of approved

cell therapy-based products include Prochymal (human adult MSCs expanded ex-vivo) and Holoclar

(limbal epithelial stem cells). Prochymal was approved in Canada and New Zealand for paediatric use

in acute GvHD33 - discussed in Section I.1.4.4 -, while Holoclar was approved by the European

Medicines Agency (EMA) for the treatment of corneal diseases34.

Aside from ethical issues, the main reason why embryonic stem cell-derived therapy products

have not been approved by regulatory agencies such as the United States Food and Drug

Administration (FDA) is due to concerns related with the safety of transplanting embryonic stem

cell-derived products.35 Cell products derived from pluripotent stem cells entail the possibility of

spontaneous teratoma formation (i.e. residual undifferentiated stem cells) and the presence of

unwanted cell types.35 Furthermore, (stem) cell therapy implicates other risks such as biological or

donor-to-donor variability, lack of genetic, epigenetic and overall stability (a major hurdle when

ensuring lot-to-lot consistency), microbiological contamination and immunological responses to

alloantigens.35,36 Ultimately, until standards and methodologies for preclinical safety and efficacy

evaluation - namely potency assays -37, product characterization, and process validation and control

are fully developed, the approval and implementation of (stem) cell therapy is hindered.36

I.1.2.3 Tissue Engineering

Cell therapy alone is not sufficient to regenerate large tissue defects or even replace whole

organs. In this regard, the approach of “tissue engineering” is the more promising strategy.23 Two main

approaches are used in tissue engineering. The acellular approach involves the use of natural or

synthetic matrices (often termed scaffolds) alone, while the cellular approach involves the use of cells,

which are processed and seeded onto a scaffold (cell-seeded scaffold approach) before

implantation.17

7

Tissue generation is a complex procedure comprising several steps, beginning with the

isolation of cells from a biopsy, the expansion/differentiation of cells in vitro, the generation and

maturation of a 3-D construct, and subsequently, the use of the construct as a test system or graft.38

One of the first successful cases of a tissue-engineered organ transplant occurred in 2008, in which a

patient from Spain received a re-engineered trachea. Briefly, a donor trachea was first decellularized

using a detergent (without denaturing the native ECM) and the resulting scaffold was re-cellularized in

a rotating bioreactor using autologous epithelial cells and mesenchymal stem cell-derived

chondrocytes, where cells were exposed to both air and liquid phases to mimic the environment of the

organ.39

3-D shaped biomaterials - scaffolds - are of extreme importance for tissue engineering

applications, since the properties of the substrate on which the cells are seeded influence cellular

responses and cell fate.30 Furthermore, scaffolds should present various requirements, such as the

presence of interconnected micropores (allowing cell seeding and proliferation, as well as vascular

formation and waste transport), optimal porosity with adequate surface area and mechanical

strength.40 Although synthetic/biosynthetic biomaterials and, most recently, nano-biomaterials, are

used for stem cell differentiation, naturally-derived scaffolds are the most appropriate for developing

tissue engineering constructs, mainly due to their biocompatibility. For instances, biomaterials like

agarose, alginate, hyaluronic acid, gelatin (porous denaturated collagen), fibrin glue, collagen

derivatives, and decellularized tissue matrices can be used as supporting scaffolds.30 In particular,

decellularized matrices are biodegradable and bioresorbable (supporting the reconstruction of a

completely normal tissue without significant inflammation) and provide efficient scaffolds for cell

seeding, as well as inherent bioactivity and mechanical similarity to native ECM.12,30

After seeding the expanded cell population onto a tissue-specific scaffold (tissue generation

phase), comes the maturation phase, when cells can adapt to the 3-D environment, produce ECM,

establish cell-cell interactions and spread along and through the construct. During this phase, tissue

development is guided by various cues, such as cell-matrix interactions, soluble factors and physical

stimuli (mechanical, electrical and electromagnetic cues).38 The finalized tissue construct can be

further used as an implant or as a test system (e.g. for drug screening or toxicology testing).

One of the most relevant hurdles inherent to tissue engineering is tissue neovascularization,

which is essential to supply oxygen and nutrients to the cells in the 3-D constructs. It is virtually

impossible to expect the neovascularization throughout a cell–scaffold construct in the case of in vitro

tissue engineering.40 As a consequence, the construction of entire vascularized organs in vitro is

precluded by the complexity of generating new vessels. As of today, only cartilage-composed organs

(e.g. ear)41 or relatively simple organs (e.g. trachea)39 have been constructed through tissue

engineering approaches. Still, some tissue engineering-derived products are currently on the market,

including autologous cultured chondrocytes and allogeneic fibroblast products (i.e. Carticel and

Dermagraft, for cartilage and skin replacement, respectively).35

8

I.1.3 Muscle Cells

As mentioned previously, the urethral sphincter is populated by (i) SMCs on the inner layer

and (ii) skeletal myofibers on the outer layer. Given the importance of the urethra musculature, in order

to construct an engineered urethral tissue, to use as a tool for urethral cell therapy, functional layers of

both smooth and skeletal muscle cells must be generated via tissue engineering-related approaches.

I.1.3.1 Smooth Muscle Cells

Bladder smooth muscle fibers have a fusiform shape, are 300-400 m in length and are of the

single unit (visceral) type (i.e. become excited and contract as a single unit).42 SMCs in general can be

derived from eight different origins: neural crest, secondary heart field, somites, various types of stem

cells, mesangioblasts, proepicardium, splanchnic mesoderm and mesothelium.43,44

Bladder smooth muscle differentiation is regulated by epithelial-mesenchymal interactions and

a variety of factors and signalling pathways.42 An epithelial signal is necessary for the induction of

smooth muscle differentiation from the bladder mesenchyme, which seems to govern the pattern of

this muscle.42 Among the factors that regulate SMC differentiation, sonic hedgehog (Shh), expressed

by the bladder urothelium, is one of the most important. Transforming growth factor-β (TGF-),

expressed in the developing bladder, and serum response factor (SRF), a key controller of

muscle-specific gene expression, as well as some DNA-binding proteins, are also involved in bladder

smooth muscle differentiation.42

In contrast to terminally differentiated skeletal myocytes (mentioned in Section I.1.3.2), SMCs

retain remarkable plasticity and can undergo profound and reversible changes in their phenotype in

response to changes in local environmental cues.45 This plasticity is based on the ability of SMCs to

exhibit either a “synthetic” or a “contractile” phenotype. By switching to a “synthetic” phenotype - a

process that can be dependent on arterial injury, growth factors and arteriogenesis - SMCs change

their phenotypic markers and start secreting ECM proteins.45,46

Generally, differentiated SMCs (displaying a “contractile” phenotype) exhibit low proliferation

rates, low synthetic activity, and express a unique repertoire of contractile proteins, ion channels, and

signalling molecules required for the contractile function of these cells.43 Some of these molecules,

especially contractile proteins, can therefore be used as markers for SMCs. Smooth muscle alpha

actin (-SMA) is considered a marker of early SMC differentiation and the most general marker of the

SMC lineage, although it is also expressed by myofibroblasts and endothelial cells under certain

conditions.42,43,46 Smoothelin, a cytoskeletal protein found in mature SMCs, is absent in myofibroblast

cells, providing a potential marker to distinguish between SMCs and fibroblasts.46 Calponin and

SM22 (an actin-associated protein) are considered intermediate markers while smooth muscle

myosin heavy chain (SM-MHC) is a marker of fully differentiated SMCs, thus being considered the late

and most specific maker of differentiated SMCs.42

9

I.1.3.2 Skeletal Muscle Cells

The process of generating skeletal muscle – myogenesis – is used to describe de novo

embryonic muscle formation (embryonic myogenesis) and the regeneration of injured muscle (adult

myogenesis), which are very similar processes in terms of the regulatory molecules involved.47,48

Embryonic myogenesis is a highly controlled mechanism induced in the mesoderm of the embryo,

implicating signalling pathways (mainly Notch and Wnt) and transcription factors, including Six1/4,

Pax3/7 and the so-called myogenic regulatory factors (MRFs) MyoD, Myf5, myogenin (MyoG) and

MRF4 (also known as Myf6), which are collectively expressed in the skeletal muscle lineage.47,48,49

During embryonic myogenesis, some embryonic progenitors specialize into satellite stem

cells, while others skip this quiescent (i.e. not mitotically active) stage and directly differentiate into

myoblasts.47 At this early stage in the lineage progression, cell fate is mainly regulated by transcription

factors Six1/4, Pax3/7, Myf5 and MyoD (Figure I.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.

Both quiescent and activated satellite cells express Pax7 (which is considered an evolutionary

conserved specific marker), while Pax3 is only expressed in some muscle types.47,48,50 Non-committed

satellite cells exhibit a Pax7+Myf5- phenotype whereas committed cells display Pax7+Myf5+.48,50,51

Upon activation, those committed but still quiescent satellite cells (Pax7+MyoD-) simultaneously

upregulate MyoD, enter the cell cycle, and proliferate as Pax7+MyoD+ myoblasts. By downregulating

Pax7 and upregulating MyoG and myosin heavy chain (MHC), myoblasts enter the differentiation

10

program (Pax7-MyoD+MyoG+).50,51 This last stage involves myoblasts aligning, giving rise to myocytes,

and fusing homotypically to form multinucleated myofibers or myotubes.47,48

Once embryonic myogenesis is completed, the remaining satellite stem cells will reside in a

specialized environment within skeletal muscle termed “satellite cell niche”, which serves as a pool of

myogenic progenitor stem-like cells. The formation of this niche ensures muscle regeneration

throughout the adult lifetime, by supporting the self-renewal of satellite cells (in response to stimuli

such as injury) and preventing their differentiation.51,52 Interestingly, satellite cells have also been

described as multipotent stem cells potentially capable of differentiating into other cell types belonging

to the mesenchymal lineage, including adipocytes and osteocytes.53

Within the niche, satellite cells surround myofibers, lying between the plasmalemma and the

basal membrane.47 Moreover, 95% of satellite cells are subjacent to a vessel and separated from it by

the basement membrane, which allows their function to be modulated by other cell types not in direct

contact with them.48,51,54 The differentiation of satellite cells in adult myogenesis, which occurs upon

muscle injury, is highly dependent on extrinsic regulatory factors. Signalling molecules such as

fibroblast growth factor (FGF), members of the TGF-β family, insulin-like growth factor-I and II

(IGF-I/II), hepatocyte growth factor (HGF) and interleukins take part in muscle regeneration.47,48

Quiescent satellite cells express CD34, a surface marker implicated in cell adhesion and signalling;

however, it is downregulated when cells are activated.48,51,55 CD56 (also known as neural cell

adhesion molecule-1) is considered one of the most reliable markers of satellite and myoblast cells in

human muscle (although it is not lineage specific), since it is expressed in quiescent cells and is

further maintained during proliferation and differentiation.52,56

Although satellite cells exhibit tremendous regeneration potential in vivo, they highly depend

on their niche environment to keep their proliferation ability.47,54 This niche “addiction” is a major hurdle

when large cell numbers are required, for instances, for cell therapy purposes (generally 1-5 millions of

cells/kg of body weight)57. After isolation, satellite cells can only divide a small number of times and

rapidly differentiate into myoblasts irreversibly expressing Myf5, MyoD and MyoG.44,48,49 Other

disadvantages of using these cells in cell therapy or tissue engineering settings include poor

accessibility and low numbers of satellite cells obtained from muscle biopsies.47

Although myoblasts may seem a good alternative due to their proliferative capacity, which

makes their expansion in vitro possible, this capacity is largely reduced as the number of rounds of

cultivation (i.e. passages) increases, and so does their ability to fuse and produce myofibers.59

Allogeneic myoblasts transplanted into Duchenne muscular dystrophy patients have been, in early

trials, largely inefficient owing to immune rejection, rapid death (presumed to be caused by anoikis

induction, i.e. apoptotic death caused by reduced contact with ECM components)60, and limited

intramuscular migration.52,58 Finally, the use of myofibers as therapeutic agents is precluded by their

barely detectable postmitotic turnover in healthy young muscle and, consequently, lack of turnover in

vitro.51 Therefore, an alternative cell type with both proliferative capacity and stability in vitro, as well

11

as myogenic differentiation potential, must be selected for cell therapy and tissue engineering

applications involving skeletal muscle cells.

I.1.4 Mesenchymal Stem/Stromal Cells

In the early 1990s, Arnold Caplan at Case Western University used the term “Mesenchymal

Stem Cell” to describe BM-derived cells with the ability to both self renew and differentiate along

classical mesenchymal lineages, including adipocytes, chondrocytes, myocytes, osteoblasts, and

tenocytes.61,62 The term “Mesenchymal Stromal Cells” is also valid, since MSCs belong to the marrow

stroma, providing support and nourishment to HSCs. These cells were first reported by Friedenstein

and colleagues in 1970, who identified the presence of fibroblast-like cells that could be obtained from

the adult BM, form colonies on plastic and differentiate into mature cells of mesenchymal lineages,

referring to them as Colony-Forming Unit-Fibroblasts (CFU-Fs).63,64

I.1.4.1 Definition and In Vitro Characteristics

According to the International Society for Cellular Therapy (ISCT), MSCs are defined by three

criteria: (1) adherence to plastic; (2) specific surface antigen expression (positive for CD73, CD90 and

CD105 in greater than 95% of cells in culture, but negative for hematopoietic markers CD14, CD34

and CD45, as well as CD11b, CD79a and human leukocyte antigen-D related (HLA-DR) surface

markers); and (3) multipotent differentiation potential in vitro (capable of osteogenesis, adipogenesis

and chondrogenesis) demonstrated by staining.65 Currently, these criteria are unanimously considered

as extremely minimalist, as they presuppose that practically every non-clonal culture of cells from any

connective tissue could potentially be classified as a MSC culture under the right culture conditions.66

Although not officially recognized by the ISCT, besides being able to differentiate into stromal

cells, osteocytes, chondrocytes and adipocytes, MSCs are also capable of differentiating into

myotubes in vitro (Figure I.6). 67–69

MSCs belong to the mesodermal lineage, the germ layer that includes blood, bone, BM,

muscle, adipose tissue (AT) and connective tissues.24 Although MSCs are currently defined by their

surface antigen expression, there is no cell surface marker that specifically and uniquely identifies

MSCs. Other expressed markers include Stro-1 (which is associated with the capacity to form

colonies, and, therefore, is gradually lost during culture expansion), CD200 (OX-2), adhesion

molecules such as CD54, CD102 and CD50 (or ICAM-1,2 and 3, respectively), several growth

factors/cytokine receptors and integrins.70–72

Remarkably, MSCs display a unique immune phenotype, described as MHC I+, MHC II-,

CD40-, CD80-, CD86- and regarded as non-immunogenic.73 As mentioned above, MSCs are negative

for CD45 (leukocyte common antigen) and HLA-DR antigens, suggesting that these cells can be

immunoprivileged. In fact, MSCs appear to use an array of mechanisms to minimize rejection by the

12

host including modulation of dendritic and T cell function, as well as the creation of a suppressive

microenvironment.74 Clinically, this immune phenotype represents a great advantage, as it

theoretically allows the transplantation of allogeneic MSCs into a host without the need of

immunosuppression.

As stem cells, MSCs have the ability to proliferate without spontaneously differentiating, while

maintaining the ability to differentiate into multiple cell types in vitro.75,76 These characteristics allow

multiple in vitro passages of expansion to be performed, reaching high cell numbers of undifferentiated

MSCs (e.g. 50 to 375 million cells generated by passage 2 from a 10 ml marrow aspirate)75.

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.

I.1.4.2 MSCs Sources

One other important feature that allows the use of MSCs in Regenerative Medicine relies on

their accessibility and ease of harvest from various tissues. MSCs have been isolated from the BM,

peripheral blood, cord blood, cord Wharton’s jelly, AT, amniotic fluid, compact bone, periosteum,

synovial membrane and synovial fluid, articular cartilage and fetal tissues.64 MSCs have been further

identified in skeletal muscle, hair-follicle and urine.77 However, differences in culture morphology,

13

growth rates, proliferation potential, and differentiation capacity can occur depending on the tissue

from which MSCs are isolated.64

The most promising MSC sources in terms of significance of the number of cells obtained and

the amount of descriptive data in the literature correspond to BM (BM-derived MSCs – BM-MSCs), AT

(adipose tissue derived-stromal/stem cells – ADSCs) and umbilical cord (umbilical cord matrix stem

cells or umbilical cord derived MSCs – UCM-MSCs).

Although BM is considered the traditional source of MSCs, these cells constitute only a small

percentage of about 0.01%-0.001% of the total population of nucleated cells in the marrow, and the

harvesting process can involve an invasive and painful procedure.32,75 BM-MSCs are isolated from the

mononuclear layer of BM aspirates after separation by density gradient centrifugation. Once cultured

in vitro, MSCs adhere to the plastic surface of culture plates, while hematopoietic progenitors and

other non-adherent cells are removed by medium changes.75

Human AT can be routinely obtained as excised surgical specimens or as lipoaspirates (often

resulting in more than 1 L of tissue), and subsequently digested with collagenase and centrifuged.62 It

has been described that a single millilitre of human lipoaspirate can yield between 0.25 to 0.375x106

ADSCs, capable of differentiating into adipocyte, chondrocyte and osteoblast lineages in vitro.62

UCM-MSCs are collected from the umbilical cord by explant culture or by enzymatic digestion

and filtration, achieving yields of 8.6x105 of adherent cells/cm.78 UCM-MSCs present a greater

proliferation capacity and faster growth rate in comparison to BM-MSCs, as well as a lower expression

of HLA-class I.32,78 The umbilical cord presents other advantages such as accessibility, painless

procedures to donors, the possibility of autologous and allogeneic cell therapy and lower risk of viral

contamination.78 However, the number of cells obtained from one single cord unit is scarce for clinical

applications, thereby requiring ex-vivo expansion strategies.

I.1.4.3 In vivo Roles of MSCs

It has been suggested that cells with mesenchymal characteristics reside in virtually all

postnatal organs and tissues.73 The fact that MSCs seem to reside in almost every tissue of the

human body incites questions about the in vivo role of these cells, apart from their differentiation to

mesenchymal lineages (allowing the growth and natural turnover of mesenchymal tissues).

In fact, MSCs are known to secrete a great variety of bioactive factors with paracrine and

autocrine effects (referred to as trophic factors/effects) such as enhanced angiogenesis and tissue

regeneration, inhibition of fibrosis (scar formation) and apoptosis, and stimulation of mitosis and

differentiation of tissue-intrinsic progenitor cells.79,80 It should be noted that these trophic

factors/effects do not lead to the differentiation of the secreting MSCs, but rather primarily affect their

neighbouring cells. Moreover, MSCs act by displaying immunomodulatory activities (including the

suppression of the local immune system), recruiting endogenous MSCs to the injury site and possibly

transferring mitochondria or vesicular components containing mRNA, microRNA and proteins.81

14

MSCs have the ability to migrate chemotactically into tissues showing inflammation and injury,

possibly in response to signals that are upregulated under injury conditions73. After being mobilized to

the injury site, MSCs are presumed to act as curative agents by becoming activated and establishing a

regenerative microenvironment, through the secretion of bioactive molecules and the regulation of the

local immune response.80

In conclusion, the currently proposed mechanisms of action for MSCs in vivo include not only

differentiation but also paracrine/trophic effects, immunomodulatory effects, and transfer of vesicular

components.81 While the differentiation capacity of MSCs benefits tissue engineering-related

applications, the capacity to secrete bioactive factors with regenerative and immunomodulatory effects

could be beneficial for cell therapy applications, such as tissue repair.

I.1.4.4 MSCs in Clinical Trials

The success of MSCs in modulating immune responses and promoting tissue repair in

preclinical studies has prompted the exploration of MSCs in clinical settings.29 MSCs have been

explored to treat myocardial infarction, multiple system atrophy, liver failure and cirrhosis, diabetic

critical limb ischemia and foot ulcer and GvHD, among other diseases, with no severe side affects.29 In

particular, GvHD, a severe reaction followed by allogeneic HSCs/bone marrow transplants, in which

the transplanted cells “reject” the host’s own cells, can be attenuated by MSC co-transplantation.82

Little is known about the mechanisms of suppression of GvHD, however, the release of soluble

factors, induction of regulatory T cells, and repair of damaged target organs are possible

mechanisms.82 As mentioned in Section I.1.2.2., a MSC-based product (i.e. ex-vivo expanded cells)

named Prochymal was already approved for paediatric use in GvHD, in Canada and New Zealand.

MSCs do not show tumorigenic activity, as opposed to pluripotent stem cells (mentioned in

Section I.1.2.2), which makes them an extremely encouraging stem cell type for Regenerative

Medicine proposes. Other clinical MSC applications include tissue engineering constructs or direct

MSC injection for bone and cartilage repair. For autologous cartilage repair, natural matrices

(e.g. alginate and collagen), as well as synthetic polymers are available, with the design of the ECM

being of extreme importance, since it must lead to proper differentiation of MSCs into

chondrocytes.23,83 Autologous MSCs were successfully implanted in patients to enhance

fracture/osteotomy healing, fill bone defects, treat pseudarthrosis, bone cysts, osteonecrosis, or

enhance spinal fusion.23 Besides their astonishing osteogenic differentiation capacity, MSCs are

capable of secreting paracrine factors that enhance angiogenesis during fraction healing.23 These

angiogenic properties give great hopes for MSCs as therapeutic agents for diseases caused by limited

angiogenesis, such as peripheral artery disease (ischemia), myocardial infarction and cerebral

ischemia/stroke.81

However, the exploitation of the trophic properties of MSCs for cell therapy may come with

one disadvantage: the shifting from local delivery of MSC to systemic administration, although less

invasive and more convenient, causes only a small percentage of the infused MSC (often <1%) to

15

reach the target tissue with cell entrapment commonly observed in capillaries within the liver, spleen

and lung.84 Furthermore, the absence of a specific marker of MSCs as well as the

heterogeneity/inconsistency of MSCs isolation and expansion techniques are still major hurdles, which

make it difficult to compare pre-clinical and clinical results from different research groups.81

I.1.4.5 Emerging Issues: Muse cells and the Perivascular Niche Hypothesis

Although there is a general agreement on the multipotency capability of MSCs (capability of

differentiating into most mesodermal cell lineages, excluding the hematopoietic cell lineage), some

reports have emerged in the past decade suggesting that MSCs might be pluripotent (capable of

differentiating into mesodermal, ectodermal and endodermal lineages) or, at least, able to

transdifferentiate.27 However, most of these results have been widely debated due to their

non-reproducibility, which discredits their reliability and authenticity.

Nonetheless, a supposedly pluripotent stem cell type, residing among MSCs, was recently

found by Kuroda and colleagues: multilineage-differentiating stress-enduring (Muse) cells.85 As their

denomination suggests, Muse cells are resistant to stress conditions, namely 16-hour incubation with

collagenase or trypsin, low temperatures, low serum and hypoxia.85,86 This subpopulation of cells

expresses CD105, CD90 and CD29, markers expressed by MSCs, but also pluripotency markers

(SSEA-3, Nanog, Oct3/4, and Sox2).85,86 These cells are able to differentiate into endodermal,

ectodermal, and mesodermal cells both in vitro and in vivo when stimulated by certain combinations of

cytokines and trophic factors.85 However, they only comprise 1% of BM-MSCs grown in adherent

culture, and are thought to exist in a dormant, or quiescent state under normal physiological

circumstances within the cellular niche.85 Muse cells have been efficiently isolated from BM85, AT86

and dermis85, as well as from commercially available cultured fibroblasts and ADSCs87,88, and were

further characterized by different research groups (namely, a research team at the University of

California, Los Angeles)86, which suggests that the existence of these pluripotent MSC-like cells is

possible. Still, the existence of pluripotent cells in adult mesenchymal tissues is a very controversial

subject, and more studies confirming the characterization and differentiation potential of these cells

are required in order to replicate and credit the abovementioned data.

Notably, Muse cells do not seem to show tumorigenicity when transplanted in vivo and do not

require genetic modification strategies to differentiate, which enables their use for clinical cell therapy

applications.85,89 In fact, the therapeutic potential of these cells was very recently explored in animal

models of cerebral ischemia85 and of diabetic skin ulcers86. Through the selection of SSEA-3 positive

cells from the BM and AT by fluorescence-activated cell sorting (FACS) and magnetic-activated cell

sorting (MACS) (mentioned in Section I.2.1.2.), the sorted Muse-enriched and Muse-depleted

populations were transplanted into the mentioned animal models.90,91 In the first study, only the

Muse-enriched population showed engraftment capacity into ischemic tissue, while in the second

study this population displayed greater therapeutic effects to accelerate impaired wound healing

associated with type 1 diabetes.90,91

16

The reported presence of quiescent Muse cells in vivo raises questions about the existence of

a MSC niche, where more primitive MSC-like cells might be maintained under severe stress

conditions, such as hypoxia. In fact, some data suggest that hypoxia enhances not only the

proliferative capacity but also the plasticity of MSCs.70,92,93 However, the presence of a true MSC niche

has not been confirmed nor identified. Nevertheless, a theory suggesting that the MSC niche is of

perivascular nature is currently under development.

At the same time, a major unresolved question relates to the identity of MSCs within their

native tissues. There is growing evidence that pericytes (also referred as Rouget cells or mural cells),

may be the native cells of the ex vivo MSCs.64 MSCs express CD106, or VCAM-1 (vascular cell

adhesion molecule-1), also expressed on blood vessel endothelial and adjacent cells, and have been

identified in perivascular locations.70,94,95 Furthermore, MSCs and pericytes share remarkable

similarities such as their differentiation potential (into osteoblasts, chondrocytes, and adipocytes), the

ability to maintain multipotency at high passage numbers in culture and common marker

expression.71,94 These similarities, in particular the fact that pericytes express cell surface markers that

are identical to those expressed by isolated MSCs94, led Caplan to speculate if all MSC are, in fact,

pericytes.96 However, there is controversy around the connotation of pericytes as “stem cells”, since

they are not know to proliferate and differentiate into mature cell phenotypes in vivo following injury.64

Ultimately, the possibility that MSCs reside in a perivascular niche throughout the body would

give them easy access to all tissues in response to systemic influences related to injury, lending

credence to the notion that MSCs are integral to the healing of many different tissues.70,71 In

conclusion, the existence of a perivascular MSC niche is plausible, where MSCs would interact with

other cell types, ECM and signalling molecules (Figure I.7) and periodically migrate to the bloodstream,

in situations of injury or disease. Additionally, as the O2 tension within the niche is presumed to be

variable70,97, it can be hypothesized that oxygen-dependent cell-specific compartments are likely to

exist, and that hypoxia conditions can be eventually associated with the presence of

primitive/progenitor MSCs (presumably Muse cells).

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.

17

I.1.5 Tissue Engineering-derived Urethral Constructs in Clinical Trials

Few cases of tissue engineering-derived urethral constructs have been developed and tested

on a pre-clinical and clinical basis for the treatment of SUI, contrasting with cell therapy approaches,

which have been more assessed in the literature. Still, a pre-clinical study tested the feasibility of

seeding autologous urothelial cells and SMCs, harvested from bladder biopsies, on tubularized

acellular collagen scaffolds for urethral replacement in canine models, when compared to tubularized

matrices without cells.98 The results showed that the urothelial cells formed a multilayered epithelial

lining on the luminal side of the scaffold, while the muscle cells formed a layer on the outer surface.98

By contrast, the animals that received the tubularized matrix without the cells developed urethral

strictures, urine extravasation, and hematoma and fistula formation.98 These results show the

importance of including cellularized scaffolds in the treatment of incontinence, as the inclusion of

scaffolds alone is not enough for complete tissue healing, and may lead to complications.

The same type of approach was implemented in a clinical study where human patients with

urethral defects underwent urethral reconstruction with tissue-engineered tubularized urethras.99

Bladder biopsies were performed in order to collect SMCs and urothelial cells from the patients, which

were seeded onto tubular scaffolds made from polyglycolic acid.99 Although most patients became

continent after urethral reconstruction, one of the patients, who suffered from SUI caused by a pelvic

disruption that involved the sphincter, needed a pubovesical sling after surgery.99

Thus, in cases of SUI where an intrinsic sphincter deficiency is what mainly causes

incontinence, alternative clinical procedures to reconstruct the sphincter are required. Furthermore, the

need to perform biopsies to collect autologous cells carries additional risks to the patient, as this

surgical procedure may cause further complications in the genitourinary tract. Therefore, novel

autologous cell sources should be explored.

To the extent of the reviewed literature presented herein, no studies using MSCs as a cell

source for tissue engineered-urethra constructs, to be applied at a clinical level, were found. Still,

MSCs have been used in clinical studies where stem cell therapy approaches were implemented to

treat SUI. One particular clinical study aimed to treat male SUI (caused by urethral sphincter

deficiency) by injecting autologous ADSCs into the patients’ rhabdosphincter.100 At 6 months after the

injection, UI improved in terms of leakage volume and the function of the urethral sphincter was

improved in all cases.100

MSCs have been considered one of the most promising types of cells to be used in SUI

therapies.2,101 Nevertheless, a major challenge in using MSCs for muscle tissue engineering is to

robustly direct their differentiation towards the desired cell type and having the cells retaining that

specific phenotype in vitro and, importantly, upon implantation in vivo.

18

I.2 Myogenic Differentiation of MSCs: Current Status I.2.1 Differentiation of MSCs into Skeletal Muscle Cells

The myogenic capability of MSCs has been demonstrated both in vivo, through the

transplantation of MSCs in animal models, and in vitro, through the use of myogenic differentiation

induction strategies. An in vivo animal study demonstrated the survival of BM-MSCs transplanted in

the urethral wall of urethral sphincter injured rats, as well as their differentiation into striated muscles

and nerves.19 The cells were identified within rhabdosphincters on day 14 after transplantation, and

were positive for both desmin and skeletal MHC. Additionally, some injected cells were positive for

PGP9.5, a neural specific protein, suggesting that a small fraction of MSCs differentiated into nerves.19

Several in vitro approaches have been developed in order to differentiate MSCs into

myoblasts. The most common strategies found in the literature rely on the addition of specific soluble

factors and/or DNA demethylation agents, MSC co-culture with myoblasts and the use of patterned

surfaces. Dulbecco's modified Eagle's medium (DMEM) supplemented with Horse Serum (HS), among

other factors, is a commonly used myogenic induction medium used in the literature.69 The

supplementation of DMEM with 5% HS and corticosteroids (dexamethasone and hydrocortisone) to

UCM-MSCs was shown to lead to the sequential expression of MyoD, MyoG and MHC, in accordance

to the myogenic differentiation pattern.102

Regarding the co-culture procedures found in the literature, Gunetti and colleagues concluded

that BM-MSCs do not fuse with striated muscle cells in both in vitro (co-culture with C2C12 myoblasts)

and in vivo (transplantation into rat pelvic muscles) conditions.103 However, when seeded onto a

laminin matrix and plated in the presence of Insulin-Transferrin-Selenium (ITS) supplement and

Epidermal Growth Factor (EGF), MSCs expressed myogenic markers, although myotube formation

was not observed.103 Beier and co-workers observed that MSCs showed enhanced ability to fuse and

contribute to myotube formation when cultured with primary myoblasts under the stimulus of bFGF and

dexamethasone.104 Interestingly, in both studies the authors demonstrated that MSCs constitutively

express certain muscle-specific markers. Additionally, Garza-Rodea and colleagues reported that

human MSCs derived from AT, BM and synovial membrane in co-culture with murine C2C12

myoblasts and plated in the presence of ITS formed hybrid myotubes at low frequencies.105 Notably,

when transplanted into an animal model of muscle damage, AT-derived MSCs contributed the most for

myoregeneration in vivo.105 These results suggest that co-culture with myoblasts may not be sufficient

to fully induce MSCs to differentiate into skeletal muscle and that the combination of specific growth

factors/hormones, some known to promote myogenic differentiation106, may be essential.

Finally, the use of matrices with the purpose of inducing myogenic differentiation of MSCs has

also been investigated, with some successful case studies. Recently, efficient commitment of

BM-MSCs to the myogenic lineage was accomplished through plating on a biomimetic polyurethane

acrylate resin replicating myoblast topography.107 The resin, fabricated through a nano-imprint

lithography technique, induced morphology changes and higher expression of muscle-specific genes

in cultured MSCs (namely MyoD, MyoG and skeletal MHC), when compared to flat substrates.107

19

Analogously, ADSCs were shown to undergo stiffness-induced lineage commitment and

fusion (albeit at low percentages) when plated in a muscle-mimicking matrix with a stiffness value of

10 kPa, without any other form of external stimuli.68 In these conditions, ADSCs surpassed BM-MSCs,

which never underwent stiffness-mediated fusion, thus suggesting that the AT may be a source of

cells with enhanced myogenic potential when compared to cells from BM.

I.2.1.1 Stromal Vascular Fraction: The Myogenesis-committed Subpopulation

The stromal vascular fraction (SVF) of the white AT consists of a heterogeneous population of

cells comprising adipose stromal cells (15-30%), hematopoietic-lineage cells (25-45%), endothelial

cells (10-20%), pericytes (3-5%) and fibroblasts, among others.108,109 According to the ISCT, stromal

SVF cells are identified phenotypically by the following phenotype: CD45-CD235a-CD31-CD34+.

Through plating, the plastic-adherent population (e.g. ADSCs) is isolated, and these cells are known to

retain MSC markers, including CD90, CD73 and CD105 and remain negative for CD45 and CD31.108

Cells derived from the SVF cultured in DMEM supplemented with 5% HS alone have been

reported to spontaneously differentiate into skeletal muscle cells without any additional external

stimuli.110 Di Rocco and co-workers reported that this rare subpopulation of myogenesis-committed

cells only corresponded to 0.001% of total plated cells.110 Concordantly, Shan and colleagues reported

that the myogenic differentiation potential of the white AT SVF-derived cells resides in the

Myf5-lineage progenitors.111 The Myf5-lineage progenitor pool contains subpopulations of adipogenic

and myogenic progenitors that could be isolated in transgenic mice based on stem cell antigen-1

(Sca1) expression (positive and negative expression, respectively).111 When transplanted into an in

vivo model, the Myf5+Sca1- SVF cells expressed MyoD and Pax7 and differentiated into myofibers

expressing low MHC.111 Still, hydrocortisone and/or dexamethasone supplementation together with HS

seem to drive SVF-derived cells to undergo myogenesis (assessed by multinucleation, skeletal MHC

and MyoD detection), at average frequencies around 12% and 15%, without resorting to any kind of

cell separation methodology.69,112

A subpopulation of myogenesis-committed cells was also detected in semi-solid medium

experiments, where clusters of myogenic cells spontaneously emerged from cultures of crude SVF.113

Still, further research regarding the characterization and ideal isolation and expansion of such

myogenic cells from the SVF (namely, the detection of its unique markers) is required in order to

conclude if the achievement of sufficient numbers of skeletal muscle cells from the SVF is possible.

I.2.1.2 Stem Cell Separation Technologies: FACS vs. MACS

The isolation of pure stem cell populations from a heterogeneous suspension (e.g. potential

myogenesis-committed progenitor cells from crude SVF) is a fundamental aspect of stem cell clinical

application and basic research.114 Besides purity, several criteria such as resolution capabilities and

scalability of the process, final cell viability and the achievement of clinically relevant cell numbers are

20

important to consider when choosing the ideal stem cell separation technique, which ultimately

depends on the final application of the cellular product. Stem cell separation methods can be classified

into two categories: techniques based on physical properties (size/density), and techniques based on

affinity.114 For the purpose of the work developed herein, only affinity-based techniques will be

considered, particularly FACS and MACS.

FACS is a powerful technique, with high precision, resolution and sensitivity that allows highly

pure cell populations (over 95%) to be obtained.114 This technique is composed by three major

components: the fluidics, optics and electronics.115 The separation process relies on antibodies tagged

with fluorescent dyes that bind to surface antigens either in the desired or the undesired cell fraction

(in case of positive or negative selection, respectively) in mixed cell suspensions. Through the use of

mechanical vibrations, the input liquid stream is separated into small droplets, allowing the cells to be

analysed individually.115 Each cell is then charged positively or negatively and deflected into sampling

tubes, depending on whether they fluoresce or not (Figure I.8). Still, FACS has limited throughput

(107 cells/hour) and long processing times (3h-6h) and as an alternative, separation using

antibody-coated magnetic beads can be used (Figure I.8).114,115 After mixing the cell suspension and

the beads, the mixture passes through a column under the influence of a strong magnetic field.

Bead-carrying cells are retained while the unbound contaminants are washed from the column. The

final step of the process consists on turning the magnetic field off and eluting the desired cells.

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

1. Magnetic labelling with

MACS Microbeads

2. Positive selection using MACS column

3. Elution of positively

selected cells

Positive fraction

A B

21

One of the advantages of MACS when compared to FACS is the existence of less expensive

commercial systems, while FACS requires specialised technicians to operate the machine. In

particular, for clinical applications requiring automated cell separation on a large scale, a closed and

sterile system (CliniMACS® system by Miltenyi Biotec) is already available.115,118 FACS, on the other

hand, may not be as adequate for clinical-grade applications due to possible contamination risks and

the high costs associated with the equipment and reagents.114,115 Regarding cell viability, MACS is

believed to induce lower shear stress to the cells,119 although cell lysis is likely to occur when eluting

the desired cells from the MACS column. Still, the possibility of cell interference caused by the

magnetic particles themselves is described as the main drawback of using MACS.114,115

I.2.1.3 5-Azacytidine to Induce Myogenesis

5-Azacytidine (5-AZA), a cytidine analogue, was first synthetized in 1964.120 5-AZA was

originally developed and tested as a nucleoside antimetabolite with clinical specificity for acute

leukemias.121 Antimetabolites are chemical compounds that are structurally similar to purine and

pyrimidine nucleotides, becoming readily incorporated into either DNA or RNA, and consequently

interfere with normal cell functions, particularly gene expression.122 Although the genotoxic effect of

these compounds can theoretically occur upon all cells undergoing synthesis of new DNA, they mostly

affect fast-diving cells such as tumour cells, which is the reason why antimetabolites are used as

chemotherapeutic agents.122 Currently, 5-AZA is marketed under the trade name Vidaza, as well as its

deoxy derivative, 5-aza-2’-deoxycytidine (5-AZAd) or Decitabine, for the treatment of myelodysplastic

syndrome (Figure I.9).123 While 5-AZA is incorporated into both DNA and RNA, 5-AZAd affects only

DNA, which makes this compound approximately 10-fold more cytotoxic.123

Figure I.9. Structures of cytidine and its analogs, 5-AZA and 5-AZAd. R=ribose. dR=deoxyribose. Adapted from Christman,

2002.

5-AZA acts by covalently inhibiting DNA cytosine methyltransferases, which become

irreversibly bound to 5-AZA residues present in DNA.123 As methyltransferases are responsible for

DNA methylation (i.e. suppression of specific genes), their inhibition results in passive loss of

methylation, thereby allowing the expression of formerly silenced genes.123 Since the methylation

target for the mammalian maintenance methyltransferase (Dnmt1) is C-phosphate-G (CpG)

dinucleotide pairs in hemi-methylated sites, 5-AZA should indirectly reactivate the expression of genes

with promoter CpG islands.123

22

In 1995, Wakitani and colleagues demonstrated that immortalized rat BM-MSCs exposed to

5-AZAd differentiated into myogenic phenotypes in vitro.67 The cells exhibited a multinucleated

morphology, contracted spontaneously and when exposed to acetylcholine, and expressed skeletal

MHC.67 Since then, several research groups have claimed that 5-AZA-treated MSCs were able to

differentiate to skeletal muscle-like cells124–127, cardiomyocyte-like cells128,129 and osteocytes130.

In one pre-clinical study, 5-AZA-exposed rat ADSCs formed myoblast-like cells that were

injected into the urethral sphincter of animal models of SUI.131 According to the authors, the

myoblast-like cells induced from ADSCs played a positive role in improving the urine-controlling ability

of SUI animal models.131 Still, in vitro, Drost and colleagues concluded that independently of 5-AZA

exposure, MSCs express smooth and striated muscle antigens and that treatment with 5-AZA only

moderately and transiently increases the myogenic character of the cells.126 Burlacu and co-workers

applied several 5-AZA pulses to MSCs and determined that MSCs retained their multipotent capacity

after one pulse with 5-AZA, whereas additional pulses resulted in a restricted differentiation potential

with simultaneous increased ability to accomplish chondrogenic commitment.132 In this study, the

authors stated that the increased expression of muscle-specific genes observed after 5-AZA treatment

was not sufficient for the induction of differentiation into muscle cells such as cardiomyocytes.

Concordantly, Martin-Rendon and collaborators reported that only a small proportion of BM-MSCs

could generate cardiomyocyte-like cells in vitro after treatment with 5-AZA, while Liu and colleagues

demonstrated that BM-MSCs could not be induced to differentiate into cardiomyocytes.133,134

Still, 5-AZA-treated murine-immortalized BM-MSCs were shown to differentiate into

cardiomyocyte-like cells and display contractile activity,128 while human BM-MSCs induced with 5-AZA

were reported to form myotube-like structures positive for myocardium-specific α-actin, sarcomeric

β-MHC and troponin-T, although no spontaneous beat was detected.129

Contrasting with all these studies, Zhou and colleagues reported that 5-AZA treatment

facilitated osteogenic differentiation, which was accompanied by hypomethylation of genomic DNA

and increased osteogenic gene expression.130 It is possible to conclude that the results present in the

literature are, in a general matter, not consistent and even contradictory regarding the effect of 5-AZA

on MSC differentiation.

I.2.2 Differentiation of MSCs into SMCs

Great attention has been put into researching the capacity of MSCs to differentiate into SMCs,

particularly vascular SMCs, with the purpose of overcoming the primary limiting factors in vascular

tissue engineering, namely the construction of blood vessels with adequate SMC layers.135 Moreover,

SMCs are major components of hollow visceral organs, such as the urethra. However, the transition

from a stem cell to a SMC is a complex biologic process that is not yet fully understood. Still, cultured

SMCs and BM-MSCs show similarities such as the expression of -SMA, SM22, calponin and

smoothelin, and transplanted BM-MSCs have been shown to be capable of differentiating into a

23

vascular SMCs phenotype in vivo.136,137 These encouraging findings suggest that it may be possible to

establish a protocol capable of driving BM-MSC into the SMC lineage in vitro.

I.2.2.1 Soluble Factors

TGF-β plays an important role in cell differentiation and vascular remodelling.42 There is

considerable evidence that TGF-β may play a dual role in SMC differentiation, being important for both

initial commitment and further differentiation to SMCs.138 Therefore, several studies have tested the

effect of this soluble factor in MSC differentiation, frequently in association with platelet derived growth

factor-BB (PDGF-BB).139–142 TGF-β was shown to enhance the expression of -SMA in BM-MSCs, as

opposed to PDGF-BB which lowered its expression.139 However, when cultured in the presence of

both these soluble factors, or TGF-β alone, BM-MSCs could be induced to differentiate in a relatively

homogeneous way into SMCs expressing SM22, -SMA, calponin and SM-MHC.46

Wang and collaborators showed that TGF-β induced cell morphology changes as well as an

increase in actin fibers in BM-MSCs.143 According to them, TGF-β coordinates the expression of

gelsolin (an actin-binding protein that is a key regulator of actin filament assembly and disassembly)

and -SMA, to promote the assembly of -actin and actin filaments in MSCs.143 Human iPSCs were

successfully differentiated into functional SMCs through an intermediate stage of multipotent MSCs

(capable of adipogenesis, osteogenesis, chondrogenesis and myogenesis).144 At this stage, the cells

were exposed to differentiation medium containing TGF-β and heparin, and a significant increase in

-SMA, calponin, and SM-MHC (early, middle and late SMC marker genes, respectively) was

observed, as well as the exhibition of a robust fibrillar organization, which suggested the development

of a contractile phenotype. Moreover, tissue constructs prepared from these cells exhibited high levels

of contractility in response to receptor- and non-receptor-mediated agonists.144

BM-MSCs have been successfully used as a source of SMCs for vessel engineering. In a

particular study, BM-MSCs were differentiated into SMCs through the addition of TGF-β and seeded

onto a biomimetic culture system in vitro.141 The engineered vessel walls were further optimized by the

alteration of conditions such as matrix proteins, soluble factors, and cyclic strain, and the constructs

were found to be substantially similar to native vessels.141

I.2.2.2 ERK/MAPK Signalling Pathway

Few studies have shown that the extracellular-signal-regulated kinase (ERK)/

mitogen-activated protein kinase (MAPK) signalling pathway exerts an anti-myogenic effect in SMC

differentiation. This pathway plays an important role in the regulation of cell proliferation, differentiation

and migration, and is under the control of extracellular ligands related to extracellular stimuli such as

growth factors and environmental stresses.145

24

Once the ERK/MAPK pathway is activated, the terminal component of this pathway (MAPK)

translocates from the cytoplasm to the nucleus, in order to induce specific changes in gene expression

in response to extracellular signals.145 These signals come from receptor tyrosine kinases (RTKs) and

are transduced by the activation of the ERK/MAPK cascade (Figure I.10)146. Initially, a protein complex

consisting of the RAS exchange factor son-of-sevenless (SOS) and the growth-factor-receptor bound

protein-2 (GRB2) is recruited to a docking site on the RTKs.146 This results in the activation of the

small GTPase RAS, which triggers the sequential phosphorylation of three kinases: MAPK kinase

kinase (MAPKKK), MAPK kinase (MAPKK) and MAPK.146 Subsequently, the phosphorylated isoform

of the classical MAPK (known as ERK) translocates to the nucleus and phosphorylates and activates

transcription factors such as Elk-1 and c-Myc, which control the expression of genes required for cell

growth, differentiation and survival.146

The ERK/MARK pathway is associated with the repression of SRF, a critical regulator of the

differentiation of vascular SMC.140,147 The activity of SRF is regulated by single interactions with

myocardin or Elk-1.147 Myocardin functions as a very potent myogenic factor for vascular SMC and

induces the expression of SMC gene markers, including SM-MHC.148 Elk-1, on the other hand,

competes for the SRF binding site with myocardin and antagonizes myocardin, thus supposedly

disabling the induction of SMC gene markers and acting as a myogenic repressor.147

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.

Taking these findings into consideration, it has been suggested that by blocking ERK/MAPK

signalling through the use of a MEK inhibitor, the activation of SRF by myocardin would be allowed,

25

hence possibly driving the differentiation of BM-MSCs into SMCs in vitro.140 Using the small molecule

modulators PD98059 and U0126, which inhibit ERK/MEK kinases, Tamama and colleagues were able

to block the ERK/MAPK signalling pathway and upregulate SMC marker gene expression in

BM-MSCs, likely through induction of myocardin expression.140 Furthermore, the authors tested the

ability of the MEK-inhibitor treated BM-MSCs to contract or relax in response to stimuli, and observed

that the cells contracted a collagen gel in response to endothelin, the most potent vasoconstrictive

mediator, as oppose to the non-treated BM-MSCs.140 These promising results suggest that the use of

MEK inhibitors might be an effective approach to differentiate MSCs into SMCs.

I.2.2.3 Other approaches

A recent study demonstrated that high cell density promotes MSC differentiation into

contractile SMCs (measured by the expression of SMC gene and protein markers), when cultured in

growth/myogenic differentiation medium containing bFGF, due to the formation of adherens

junctions.149 This process was found to be mediated through cadherin-11, which acted as a regulator

of myogenesis, increasing the expression of TGF-β and ultimately regulating the expression of SRF.149

Direct cell-to-cell contact was shown to induce MSCs to differentiate into SMCs, through direct

co-culture of BM-MSCs with SMCs.150 By contrast, indirect co-culture and conditioned media did not

induce the expression of SMC gene markers.150

Additionally, mechanical straining and patterns of substrates were also tested to differentiate

MSCs without using biochemical reagents.151 Higher level straining (10%) and grooved surfaces were

shown to positively influence the differentiation of MSCs into SMC-like cells.151 Conversely, coating

with bladder-derived ECM and 3-D dynamic culture conditions were shown be insufficient to induce

SMC marker expression in BM-MSCs in the absence of biochemical stimuli.142 In this study, PDGF-BB

and TGF-β alone induced the expression of smooth muscle genes and proteins, including SM-MHC.142

Recently, sodium butyrate (NaB), a histone deacetylase (HDAC) inhibitor was used in a

strategy to promote rat MSCs to differentiate into SMCs.152 Concentrations of 1.0 and 1.5 mmol/L of

NaB enhanced the mRNA expression of calponin (more than 1-fold), SM-MHC (approximately 2-fold)

and -SMA (less than 1-fold). Moreover, NaB significantly enhanced MSC differentiation into SMCs in

a co-culture system with bladder SMCs and the NaB-induced cells were proven to be functional as

they were sensitive to depolarization.152 However, much like 5-AZA, this compound may act in an

unspecific way or even lead to toxicity, which is a disadvantage when compared to the use of growth

factors and/or matrices to induce differentiation.

Lastly, the use of highly porous microcarriers, produced using thermally induced phase

separation, have been developed as a method aiming the expansion and differentiation of ADSCs into

smooth muscle-like cells.153 The authors induced the cell’s differentiation by using a differentiation

medium containing TGF-β, which resulted in a significant increase in the mRNA expression of cell

contractile apparatus components caldesmon (approximately 5-fold), calponin (4.5-fold), and SM-MHC

26

(approximately 3.5-fold) at 14 days culture.153 This approach has the advantages of using

microcarriers, which allow cell expansion and differentiation to occur as a one step process and can

potentially act as a delivery vehicle for the attached cells for therapeutic applications.

I.3 Aim of Studies

The aim of this MSc project was to develop strategies to direct MSCs to differentiate into

skeletal muscle and SMCs for urethral sphincter engineering. To achieve this goal, the effects of

compounds that are described in the literature as myogenic inducers in MSCs were tested, and the

myogenic potential of cells derived from the SVF was explored. Ultimately, the discovery of the

appropriate biochemical stimuli required to establish a laboratory-based myogenic differentiation

protocol (i.e. the ideal formulation of a potential skeletal/smooth muscle induction media) was targeted.

The following specific objectives were pursued:

(i) Studying the effects of 5-AZAd in the myogenic differentiation of ADSCs;

(ii) Exploring the potential of SVF-derived cells to undergo myogenesis without the use of

demethylation compounds, co-culture strategies or in vivo engraftment;

(iii) Developing strategies for the isolation of a cell subpopulation from SVF with

potentially increased myogenic potential;

(iv) Studying the effects of compounds that act upon the activation or repression of

specific signalling pathways important in smooth muscle differentiation (e.g. small molecules such as

PD98059) in MSCs.

Finally, a decellularization protocol based on mechanochemical action was tested in porcine

urethras, with the aim of potentially combining this method with the knowledge gathered in this thesis

to further develop urethral sphincter engineering strategies.

27

II. Materials and Methods

II.1 Human-derived samples

AT samples (lipoaspirates) were obtained following donor informed consent from the abdomen

and thigh areas of both female and male healthy donors with ages ranging from 30 to 52 years old, at

Clínica de Todos-os-Santos, Lisbon, Portugal. A BM sample was obtained following donor informed

consent from a 36-year-old male healthy donor at Instituto Português de Oncologia, Lisboa, Portugal.

II.2 Culture Media

II.2.1 Ex-vivo expansion of ADSCs, BM-MSCs and SVF-derived cells

All cells were maintained in culture using low glucose (LG) (1g/L) Dulbecco’s modified Eagle’s

Medium (DMEM), supplemented with 10% fetal bovine serum (FBS, Gibco®, Carlsbad, CA) and

antibiotics (0.025μg/mL penicillin and 0.025U/mL streptomycin). The FBS used for ADSCs and

BM-MSCs culture was MSC-qualified, which means it was certified for MSC research. On the other

hand, the connotation “AT-SVF cells” is referred to cells derived from AT that were isolated and

cultured in FBS that was not MSC-qualified (standard FBS). These cells were not labelled here as

MSCs, as the SVF is composed by several distinct cell populations.108 The medium was replaced

every 3 to 4 days to diminish toxic metabolites and provide fresh nutrients, thus promoting cell growth.

Cell expansion was performed in static culture systems (T-flasks, BD FalconTM Biosciences©, Franklin

Lakes, NJ) in an incubator at 37ºC with a humidified atmosphere of 5% CO2. Images of the cell’s

morphology (in bright field microscopy) were captured with Leica Camera CD350F Software in a CK40

Microscope (Olympus®).

II.2.2 Thawing and cryopreservation of SVF cells, ADSCs and BM-MSCs

Cryopreserved SVF cells (previously isolated by an enzymatically-based method154) were

thawed and recovered by dilution in a proportion of 1:5 in DMEM+20%FBS supplemented with DNase

(1:100 dilution of 1mg/mL aliquots, Roche Applied Science, Hague Road, IN) while ADSCs and

BM-MSCs were recovered in DMEM+20%FBS. The cells were then centrifuged for 7 minutes at 1250

rpm, resuspended and plated in the appropriate culture media at cell densities between 3000 and

5000 cells/cm2 for ADSCs and BM-MSCs or at 10 000 cells/cm2 for SVF cells. When 80-90% of

confluence was reached, the cells were harvested with accutase (Sigma®, St. Louis, MO) or trypsin

0.05% (Gibco®) supplemented with 1 mM EDTA (Gibco®) for 7 minutes. Enzymatic digestion was

stopped with DMEM+10%FBS in a proportion of 3:1 and the cells were centrifuged for 7 minutes at

1250 rpm. Viable cells were counted under an optical microscope using the trypan blue dye exclusion

method (Gibco®), and part of the cells were cryopreserved. In order to protect the cells from damage

during freezing or thawing, dimethylsulfoxide (DMSO), a cryopreservative compound, was used.155

The cells were resuspended in RecoveryTM cell culture freezing medium (Gibco®) (which contains

DMSO) or in a solution of 10%DMSO (Merck Millipore©, Billerica, Massachusetts, USA)+90%FBS and

28

distributed into freezing vials (1mL/vial). The cells were then stored overnight in a -80ºC freezer and

transferred to a liquid/vapour phase nitrogen container for long-term storage.

II.3 Multilineage differentiation assays

SVF-derived cells, ADSCs and BM-MSCs were plated on 24-well plates with an initial density of

3 000 cells/cm2 for adipogenesis and osteogenesis. When 100% confluence was reached, the

medium was replaced by the respective differentiation medium (osteogenic (Gibco®) or adipogenic

(Gibco®)). For chondrogenesis, a pellet of 0.3-0.5x106 cells was resuspended and droplets of this

suspension were plated in a 24-well low attachment plate in chondrogenic medium (Gibco®). The

medium was changed every 3 to 4 days.

II.3.1.1 Adipogenesis Staining

The cells were washed with PBS (Gibco®) and fixed with 2% paraformaldehyde (PFA)

solution (Sigma®) for 30 minutes at room temperature (RT). After being washed with distilled water,

the cells were stained with 0.3% Oil Red-O solution (Sigma®) for 1 hour at RT. Lastly, the cells were

washed twice in distilled water and observed under a microscope.

II.3.1.2 Osteogenesis Staining

The cells were washed in PBS and fixed with 2% PFA for 5 minutes at RT. Afterwards, the

cells were washed and kept in distilled water for 15 minutes. Then cells were stained and incubated

with reagent X, containing 0.1M Tris-HCl solution (Sigma®), the substrate Naphthol AS MX-PO4

(0.1mg/mL) (Sigma®) dissolved in dimethylformamide (Fischer Scientific®) for alkaline phosphatase

staining, and 0.6mg/mL Red Violet LB salt (Sigma®) for 45 minutes at RT. After incubation, the cells

were washed three times, kept in distilled water and observed under the microscope. The Von Kossa

staining protocol was then performed. The cells were washed with PBS and stained with 2.5% (w/w)

silver nitrate (Sigma®) for 30 minutes at RT. The cells were washed three times with distilled water

and observed under the microscope.

II.3.1.3 Chondrogenesis Staining

The cells were washed in PBS and fixed in 4%PFA for 30 minutes at RT. Then the cells were

washed twice and incubated with Alcian Blue 1% (Gibco®) for 1 hour at RT. After being washed with

PBS and kept in distilled water, the cells were observed under the microscope.

II.4 Immunophenotype characterization

Once isolated, ADSCs, BM-MSCs and SVF-derived cells were characterized

immunophenotypically according to a panel of extracellular markers.65 BM-MSCs were additionally

tested for CD14 to evaluate monocyte contamination, as well as HLA-DR, and SVF-derived cells were

tested for satellite cell markers (CD34 and CD56). The cells were treated with 0.05% trypsin, accutase

29

or TrypLETM 1X (Gibco®) when stated, centrifuged for 7 minutes at 1250 rpm and resuspended in

PBS. The cells were distributed into FACS tubes (BD FalconTM) and incubated for 15 minutes at RT in

the dark with the following mouse anti-human monoclonal antibodies (1:10 dilution):

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.

Surface Markers Brand Conjugated Fluorophore Isotype

CD14 BioLegend® PE IgG1


CD34 BioLegend® FITC IgG1


CD56 BD Biosciences© PE IgG1


CD90 BioLegend® FITC IgG1


HLA-DR BioLegend® PE IgG2a

IgG2a BD Biosciences© PE -

IgG1/IgG1 BD Biosciences© FITC/PE -

Control groups were incubated with FITC-/ PE- conjugated mouse IgG1 isotype antibodies

(1:10 dilution). After incubation, 2 mL of PBS was added to each FACS tube, and the cells were then

centrifuged for 5 minutes at 1500 rpm and resuspended in 1% PFA. Quantitative analysis was

performed using a FACSCalibur flow cytometer (Becton Dickinson®, San Jose, CA) and with FlowJo

software (Version 8.8.6, Tree Star Inc., Ashland, Oregon, USA). MFI values were calculated by

dividing the geometrical mean fluorescence values of the sample by the ones of the isotype alone.104

II.5 5-aza-2’-deoxycytidine treatment II.5.1 Effect of 5-aza-2’-deoxycytidine on cell viability

ADSCs were seeded into 24-well plates (BD FalconTM) at cell densities of 3000 and

5000 cells/cm2 in DMEM+10%FBS (MSC-qualified). The cells were allowed to adhere to tissue culture

flasks for 24 hours, after which the medium was replaced by myogenic differentiation medium

composed by DMEM+2% HS (Sigma®) supplemented with 0, 2, 5, 7.5 or 10 μM of 5-AZAd

(Sigma®).126 After 24 hours of 5-AZAd exposure, the medium was replaced by DMEM+2%HS alone.

The cells were cultured for 2 weeks with medium changes every 3 to 4 days. At days 3, 5, 7, 12 and

14 after the 24-hour 5-AZA pulse, cell viability was assessed using the trypan blue exclusion test. The

cells were washed with PBS and harvested with accutase for 7 minutes, or with 0.05% or 0.25%

trypsin when stated. The cells were centrifuged for 7 minutes at 1250 rpm and resuspended in PBS.

II.5.2 Effect of 5-aza-2’-deoxycytidine on cell apoptosis and morphology

ADSCs were plated at a cell density of 5000 cells/cm3 on T75 flasks and 12-well plates

(BD FalconTM) in DMEM+10%FBS (MSC-qualified). Myogenic induction was performed as mentioned

30

above, with cell exposure to 0, 2 or 10 μM of 5-AZAd in DMEM+2%HS. To quantify the percentage of

apoptotic and necrotic cells after 2, 7 and 14 days of 5-AZAd exposure, the cells were stained with

FITC-conjugated anti-Annexin V and PE-conjugated anti-PI antibodies (Kit from Life Technologies®,

BD Biosciences©). The cells were analysed by flow cytometry within 1 hour. A cytoskeleton

morphology assay was performed in order to assess possible morphology changes induced by

5-AZAd. Cells plated in 12-well plates were stained for F-actin with Phalloidin-Tetramethylrhodamine

(TRITC) (1 μg/mL, Sigma®).

II.6 CD34+ SVF-derived cell sorting and myogenesis induction

Human SVF-derived CD34+ cells were isolated using CD34 MicroBeads (kit from Miltenyi

Biotec®), according to the manufacturer’s instructions. Crude SVF cells and magnetically sorted cells

from the enriched and depleted fractions were analysed by flow cytometry and plated in T25 flasks at

10 000 cells/cm2 in myogenic expansion medium156,157, composed by DMEM + 20%FBS +

10-6M dexamethasone (Sigma®) + 2.5 ng/mL bFGF (Sigma®).

The effects of gelatin (0.2%, Sigma®) and fibronectin (50 μg/mL, Sigma®) coatings, as well as

the use of high glucose (HG) (4.5 g/L) DMEM medium (Gibco®) (versus DMEM-LG, 1 g/l Glucose),

were tested for the CD34-enriched and CD34-depleted fractions. Once the cells reached confluence,

they were harvested with 0.05% trypsin for 7 minutes, tested for CD34 and CD56 expression by flow

cytometry and plated at 4500 cells/cm2 in T25 flasks and at 3000 cells/cm2 in 12-well plates, in the

same conditions. Myogenic induction was performed when the cells reached confluence, by the

addition of DMEM-HG + 2%HS for 8 days with medium changes every 3 to 4 days. After 2, 4 and 8

days of medium replacement, the cells were tested for the expression of Pax7, MyoD, myogenin and

skeletal MHC (when stated) by flow cytometry and immunofluorescence. Cells plated on 12-well plates

were also stained with Phalloidin-TRITC (1 μg/mL, Sigma®) at day 8 of induction to assess

cytoskeleton morphology and nuclei alignment and fusion.

II.6.1 CD34/CD56 decay assessment in normal and ultralow attachment plates

Fresh SVF cells were thawed and subjected to MACS sorting for CD34+ cell enrichment.

Unsorted cells and cells from the positively selected fraction were plated at 10 000 cells/cm2 in 6-well

plates and in 24-well ultralow attachment plates, in myogenic expansion medium and in skeletal

muscle cell growth medium (Zen-Bio, Inc., Research Triangle Park, NC). After 3 and 4 days of culture,

the cells plated in 6-well plates were harvested (0.05% trypsin for 7 minutes) and the expression of

CD34 and CD56 was assessed by flow cytometry. For cells plated in ultralow attachment plates, a

medium change was performed once by medium collection, centrifugation at 12500 rpm for 7 minutes

and cell resuspension in fresh medium. After 6 days in culture, the medium was collected and cells in

suspension were plated on gelatin-coated T12.5 plates. Aggregate adhesion was observed and after 4

days in culture the cells were harvested (0.25% trypsin for 10 minutes) and the expression of CD34

and CD56 was assessed by flow cytometry.

31

II.7 Smooth muscle differentiation induction

After isolating and expanding AT-SVF cells and BM-MSCs in DMEM+10%FBS (standard and

MSC-qualified FBS, respectively), the cells were plated at 5000 cells/cm2 in T25 flasks and 12-well

plates in the same conditions. When confluence was reached, the cells were exposed to smooth

muscle induction medium, composed by DMEM-LG supplemented with 3%FBS (standard FBS) and 1,

10 and 30 μM of PD98059 (Calbiochem®, Merck Millipore©, 25 mM stock solution in DMSO) for 7

days. DMEM-HG supplemented with 3%FBS plus 10 μM of PD98059 was also tested in AT-SVF cells.

Cells plated in DMEM+3%FBS, DMEM+10%FBS and DMEM-HG+10%FBS alone were tested as

control groups. The expression of smooth muscle markers -SMA, calponin, SM-MHC and desmin

(when stated) was evaluated after 2, 4 and 7 days of medium replacement by intracellular flow

cytometry and immunofluorescence. Bladder SMCs plated in bladder SMC growth medium were used

as positive control (both from Zen-Bio, Inc.). The expression of the previously mentioned markers was

evaluated by flow cytometry after 14 days of culture in P5 SMCs and by immunofluorescence after 21

days of culture in P6 SMCs.

II.8 Immunofluorescence staining

Cells cultured on 12-well plates were washed with PBS and fixed with 4% PFA for 20-30

minutes at RT, followed by rehydration with PBS. Cell permeabilization was performed with Blocking

Solution (10% NGS+ 0.1%Triton X100 (Gibco®)+PBS) for 15 minutes at RT. The cells were washed

with PBS and incubated with the primary antibody in Staining Solution

(5%NGS+0.1%Triton X100+PBS) at a concentration of 1μL/mL for 2 hours at RT. After being washed

with PBS, the cells were incubated with the respective secondary antibody (1:500 dilution) for 1 hour

at RT (see Table II.2). The cells were washed and incubated in a 1.5 μL/mL solution of

4',6-diamidino-2-phenylindole (DAPI, Sigma®) in PBS for 2 minutes for nuclei staining. The stained

cells were washed with PBS and viewed with a CK40 Microscope equipped with DAPI, TRITC and

FITC filters. Images were edited with Image-J (Version 1.48, NIH, Maryland, USA).

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.

Primary Antibody

Dilution Brand Secondary Antibody

Isotype Dilution Brand

Pax7 1:100 SCBT® Goat anti-mouse FITC IgG2a 1:500 Abcam®

MyoD 1:100 Abcam® Goat anti-rabbit Alexa 488/PE IgG 1:500 Invitrogen™

Myogenin 1:200 BD Biosciences© Goat anti-mouse Alexa 488/PE IgG 1:500 Invitrogen™

-SMA 1:40 SCBT® Goat anti-mouse Alexa 488/PE IgG 1:500 Invitrogen™

Calponin 1:100 Abcam® Goat anti-mouse Alexa 488/PE IgG 1:500 Invitrogen™

Desmin 1:200 Abcam® Goat anti-rabbit Alexa 488/546 IgG 1:500 Invitrogen™

SM-MHC 1:40 SCBT® Goat anti-mouse Alexa 488/PE IgG 1:500 Invitrogen™

32

II.9 Intracellular staining for flow cytometry

Intracellular staining was performed in order to assess the expression of skeletal

muscle-specific markers (Pax7, Myod, MyoG and skeletal MHC), smooth muscle-specific markers

(α-SMA, calponin and SM-MHC) and desmin. Cells cultured in T25 flasks were harvested with 0.05%

trypsin for 7 minutes, centrifuged for 7 minutes at 1250 rpm and resuspended in PBS. The cells were

permeabilized and fixed in FACS tubes using the BD Cytofix/Cytoperm™ Fixation/Permeabilization

Solution Kit (BD Biosciences©) according to the manufacturer's instructions. After the

fixation/permeabilization steps, the cells were incubated with a primary monoclonal antibody, washed

with PBS and permeabilization buffer, and then labelled with the respective secondary antibody (Table

II.2). Control tubes were incubated only with the secondary antibody. The cells were resuspended in

PBS and analysed by flow cytometry within 1 hour using a FACSCalibur flow cytometer and FlowJo

software. At least 1000 gated cells were considered for the flow cytometry analysis. MFI values were

calculated as previously mentioned.

II.10 Decellularization of Porcine Urethras

Four urethras from euthanized F1 Large White X Landrace female pigs, weighting between 28

and 30 kg, were obtained from Faculdade de Medicina Veterinária da Universidade de Lisboa, in

accordance with the Direcção Geral de Alimentação e Veterinária guidelines. The lower urinary tract

(bladder and urethra) was harvested immediately after slaughter, frozen and transported to the

laboratory in appropriate containers, and left overnight at -20ºC. Excess AT was removed with

tweezers and the bladders were cut transversely 2 to 4 cm above the bladder neck. The urethras were

cannulated from the bladder side (Jelco® I.V. Catheter, 14G, Smiths Medical, USA) and ligated with

sutures (Ethicon™ A185 sutures, Johnson & Johnson, USA). The decellularization process was

performed in a dynamic system. A catheter was connected to a peristaltic pump (Ecoline VC-380,

Ismatec®, Wertheim, Germany) set at 40 mL/minute, through L/S 14 G tubing (Masterflex,

Gelsenkirchen, Germany) and an adaptor. The urethras were placed inside a vessel with constant

agitation (set at 340 rpm). Prior to the decellularization process, the urethras were submerged and

perfused with distilled water for 24 hours. Decellularization was initiated once the water solution was

replaced by 0.5% sodium dodecyl sulfate (SDS) solution (Sigma®) and constant perfusion of the

urethra was allowed for at least two days, after which the direction of the perfusion was switched. The

detergent solution was renewed everyday (24-hour cycles). By the end of at least 5 days, all 0.5%

SDS solution was removed and replaced by distilled water for 60 minutes. Fresh distilled water was

then pumped for 24 hours to remove detergent residues before the urethras were recovered and

thawed at -20ºC.

33

III. Results and Discussion

In this section, the results regarding the isolation and characterization (Subsection III.1) of the

cell populations that were subsequently used in skeletal and smooth muscle differentiation studies

(Subsections III.2 and III.3, respectively) will be presented and discussed. Additionally, a brief

description of a decellularization protocol adopted for the decellularization of porcine urethras and its

results and efficacy are included (Subsection III.4).

III.1 Mesenchymal Stem/Stromal Cell isolation

Lipoaspirates obtained from healthy donors were previously processed and SVF cells were

isolated according to the enzymatic isolation protocol from Shah and colleagues.154 The SVF cells

were cryopreserved in vials and were subsequently used for the isolation of MSCs (when the culture

medium included MSC-qualified FBS) or AT-SVF cells (when standard FBS, e.g. not MSC-qualified,

was used) (Table III.1). The culture medium was changed every 3 to 4 days, allowing the

non-adherent cells to be removed, until 80-90% confluence was reached.

Table III.1. Age and gender of the AT donors and the assays performed with each sample.

Donor Information Assays Performed

SVF Donor Samples Reference

Age of Donor (years)

Gender of Donor

Skeletal Muscle Differentiation

Smooth Muscle Differentiation

SVF-1 - Female Yes Yes

SVF-2 38 Female Yes -

SVF-3 30 Male Yes -

SVF-4 52 Female Yes Yes

SVF-5 36 Female Yes -

A BM sample collected from a 36-year-old male healthy donor was isolated in

DMEM+10%FBS and the adherent cells displayed typical fibroblastic shapes. These cells were further

used for a smooth muscle differentiation assay (Section III.3).

SVF cells plated in different medium formulations showed distinct morphology and

proliferation characteristics. In particular, the use of MSC-qualified FBS or standard FBS affected the

morphology of the isolated cells. When MSC-qualified FBS was used, the cells adhered after

24 hours, allowing the identification of typical MSC-shaped cells (displaying a “fibroblast-like” shape)

after 4-5 days in culture (Figure III.1A). By contrast, when standard FBS was used, few adhered cells

could be observed in the first 2-3 days of isolation, and their shape was more heterogeneous. Early in

culture, some adherent cells looked roundish while others were more elongated. Elongated cells

usually displayed long cytoplasmic edges. As the days in culture increased, the cells started

elongating and a heterogeneous cell population was obtained. While most cells had fibroblastic

shapes, some cells displayed pericyte-like morphologies with several cytoplasmic extensions or

protrusions (Figure III.1B). Pericyte-like cells were obtained when three distinct medium formulations

34

supplemented with standard FBS were used (DMEM+10%FBS, DMEM+3%FBS and DMEM+20%FBS

supplemented with dexamethasone and bFGF, or myogenic expansion media) but were not observed

in SVF cells isolated with MSC-qualified FBS. These data may indicate that different cell populations

are obtained from the SVF depending on the type of serum that is used in the isolation step, or that

the MSC-like culture obtained with standard FBS is more heterogeneous.

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.

Among the cells isolated with standard FBS, the three medium formulations used in the

studies performed herein also affected cell proliferation rate, shape and size. These changes were

more noticeable with increased culture time. Cells plated with DMEM+3%FBS and DMEM+10%FBS

tended to suffer from contact inhibition, usually reaching 90% confluence levels at most, and adopted

a wider and flattened shape with prolonged culture time, which may be a sign of senescence.

Conversely, cells plated in myogenic expansion medium sustained a highly proliferative state,

frequently reaching 100% confluence, and maintained an elongated and spindle-like morphology. This

differences in cell proliferation and morphology can be explained by the presence of potent mitogens

(namely bFGF) in this medium formulation, which promote cell proliferation.

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.

A B

A B

35

III.1.1 Mesenchymal Stem/Stromal Cell Characterization

III.1.1.1 Immunophenotypical Characterization

After reaching confluence, the cells isolated from the SVF and BM were harvested and

characterized immunophenotypically using the surface antigens indicated by Bourin and co-workers108

and Dominici and colleagues,65 including: CD73, CD90 and CD105 (all expressed by MSCs), CD14

(expressed in monocytes), CD34 (hematopoietic progenitor/endothelial antigen also expressed in

satellite cells), CD31 (platelet endothelial adhesion molecule), CD45 (lymphocyte common antigen,

expressed in all hematopoietic cells) and HLA-DR (MHC class II cell surface receptor, not expressed

in MSCs).

According to the obtained flow cytometry results, the isolated populations expressed MSC

markers at proportions indicated by the ISCT that allow MSC classification (>95% expression for

CD73, CD90 and CD105 and <2% expression for CD45, CD31 and HLA-DR). The expression of

CD14 was higher than the indicated by Dominici and colleagues (less than 2%) for the cells tested for

this marker. However, this probably occurred because of the antibody clone used (HCD14), which was

reported to stain cross-reactive MSC populations.158 CD34 expression was higher for AT-SVF cells.

This marker is expressed in crude SVF and at variable levels in plated SVF-derived cells, depending

on the culture conditions.108 The presence of low serum conditions seemed to enhance CD34

expression in AT-SVF cells when compared to cells plated in myogenic expansion medium (Table III.2).

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

Expression (%)

ADSCs DMEM+10%FBS*

AT-SVF cells Expansion

Media

AT-SVF cells DMEM+3%FBS 15 day culture

AT-SVF cells DMEM+3%FBS 25 day culture

BM-MSCs DMEM+10%FBS*

CD14 5.5 11 - - 24

CD31 - 0.10±0.2 0.7 0 -

CD34 3.8 3.6±2.1 32 20 1.0

CD45 0.9 0±0 0.50 0 2.0

CD73 99 99±0.4 - 98 99

CD90 99 97±2.3 97 99 98

CD105 99 98±0.5 - 93 99

HLA-DR 0.30 0 - 0 0

III.1.1.2 Multilineage Differentiation Assays

The isolated ADSCs, AT-SVF cells and BM-MSCs were characterized in terms of multilineage

differentiation ability, i.e. the ability of MSCs to differentiate into adipocytes, chondrocytes and

osteocytes under specific in vitro tissue culture-differentiating conditions. This is also one criteria for

defining MSCs, according to the Dominici’s criteria.65 As shown in Figure III.3, the induction of

chondrogenesis resulted in the formation of spheroids, which marked positively for Alcian Blue,

36

indicating the presence of cartilage mucopolysaccharides namely glycosaminoglycans. On the other

hand, lipid droplets could be detected after the induction of adipogenic differentiation by Oil Red-O

staining. Finally, osteoblasts were detected by alkaline phosphatase and Von Kossa stainings.

Alkaline phosphatase stained pre-osteoblasts, while the Von Kossa staining allowed the visualization

of calcium deposits, indicative of matrix mineralization.

The results obtained below indicate that cells isolated from the BM and from the adipose SVF

(whether standard FBS or MSC-qualified FBS was used) are multipotent.

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.

There is some controversy in the literature regarding the currently used criteria for defining

MSC populations in vitro, mainly because of the fact that no exclusive marker for MSC identification is

presently known. Several authors have questioned the specificity and reliability of the relatively

minimal criteria in the ISCT definition of MSC, as these criteria refer to widely shared properties of

cells from connective tissues in general, and thus do not to allow to say, without ambiguity, that cells

meeting the criteria are only MSC and not some other cell type.66,159 Since then, the ISCT has

published a joint statement together with the International Federation for Adipose Therapeutics and

Science (IFATS) with several criteria that can be used to identify SVF cells and the respective stromal

BM-MSCs AT-SVF cells

Oste

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Dif

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Ad

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ic

Dif

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on

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ADSCs

37

adherent cell fraction (e.g. ADSCs).108 The SVF harbours a highly heterogeneous cell population,

composed by pericytes (3-5%), endothelial cells (10-20%), stromal cells (15-30%) and

hematopoietic-lineage cells (25-45%).108 The degree of heterogeneity depends, in part, on the AT

depot site and the digestion protocol.108 Although not mentioned in this official statement, other

adherent cell subpopulations such as SMC-like cells and myotube-forming cells have been

successfully isolated from the SVF.95,110,159 Thus, it is possible that the SVF includes small

percentages of myogenic-committed cells that can be useful in therapeutic approaches targeting

muscle cells.

The true in vivo identity of AT progenitors has also been a cause of debate in the literature.

ADSC location within human AT was reported to be essentially perivascular, which led some authors

to propose that ADSC are vascular precursor (stem) cells - rather than “fat stem cells” - at various

stages of differentiation capable of differentiating in situ into host tissue-specific cell types.95

Analogously, the discovery of a subpopulation of perivascular cells in multiple human organs that

expressed both MSC and pericyte markers led Arnold Caplan to speculate if all MSCs are, in fact,

pericytes.94,96

Given the inherent heterogeneity of the SVF, the isolation of morphologically and

phenotypically distinct populations is somewhat expected, especially when growth conditions not

optimized for MSCs are employed. The immunophenotypic characterization assay performed herein

suggests that stromal cells are the main components of the isolated SVF-derived cells in all the

medium formulations tested (see Table III.2). Moreover, CD31 expression levels were lower than 1%,

indicating that the adherent cell fraction obtained was not contaminated with endothelial cells (which

are present at 10-20% in the SVF). Still, given the phenotypic similarities between MSCs and other

cell types, particularly pericytes and fibroblasts94,160, non-stromal cells are likely to have been isolated

when not-MSC qualified FBS was used, resulting in the appearance of pericyte-like cells. Despite the

morphology differences observed, lineage differentiation studies indicate that all the isolated

populations (denominated here as BM-MSCs, AT-SVF cells and ADSCs) are capable of mesenchymal

trilineage differentiation (Figure III.3).

Cell morphology is an important parameter not only to evaluate cell isolation and differentiation

but also senescence. Here, changes in cell morphology with increasing culture time were observed in

medium not supplemented with mitogenic factors. A study in which Wharton’s Jelly-derived MSCs and

ADSCs were compared morphologically and in terms of proliferation capacity over long-term cultures

identified three main cell morphologies: (a) spindle-shaped, elongated; (b) polygonal, with few, small

membrane protrusions and (c) flattened, with irregular shape and large nuclei.161 Increasing passaging

of ADSCs resulted in gradual acquisition of the last mentioned type of morphology together with

increases in cell size, which the authors concluded to be signs of senescence, as cells with those

characteristics were 𝛽-galactosidase positive.161 One other study reported that independently of the

plating density and cell passage, the proportion of flat cells increases over culture time in

38

BM-MSCs.162 However, flat clones were able to give rise to spindle-like cells, indicating that flat MSCs

are not terminally differentiated and still hold some clonogenic potential.162

Although most researchers believe that spindle-like cells represent the true multipotent

subpopulation, no correlation between cell shape and “stemness” was yet established.162 This type of

information would be of extreme importance for the development of protocols aiming the myogenic

differentiation of MSC populations, especially since flat cells are thought to be more mature cells and

thus possess diminished differentiation potential.

III.2 Differentiation of Mesenchymal Stem/Stromal Cells into Skeletal Muscle Cells

III.2.1 Myogenic induction using 5-aza-2’deoxycytidine

5-azacytidine (5-AZA) and its deoxy analogue 5-aza-2’deoxycytidine (5-AZAd) are described

in the literature as myogenic inducers in BM-MSCs, ADSCs and UCM-MSCs in both in vitro and in

vivo studies at concentrations ranging from 0.1 to 10 μM. However, since these compounds indirectly

induce alterations in the chromatin conformation of DNA, potentially leading to mutations or cytotoxic

effects, cell viability assays (which are currently lacking in the literature in myogenic differentiation

studies involving these two compounds) should be performed. For myogenic induction, the protocol

used by Drost and colleagues was adopted.126 Briefly, ADSCs were allowed to adhere to the plastic

surface of tissue culture plates for 24 hours, after which 5-AZAd diluted in DMEM+2%HS was added

to cells for 24 hours (day 0), followed by medium replacement to DMEM+2%HS alone (day 1). Thus,

day 2 after 5-AZAd exposure (or treatment) refers to 24 hours in the presence of the demethylating

agent plus 24 hours without the agent. The same rationale was considered for other time points (i.e. 7

and 14 days of treatment). No 5-AZAd was added to control cells.

III.2.1.1 Cell Viability Assay

A preliminary study was conducted with P2 ADSCs in order to determine if a 24-hour pulse of

5-AZAd at concentrations of 2, 5, 7.5 and 10 μM led to cytotoxic effects, through a cell counting assay.

Cell morphology was also evaluated by optical microscopy. Additionally, two cell densities were tested

(3000 and 5000 cells/cm2).

ADSCs treated with 7.5 and 10 μM of 5-AZAd developed extremely thin and elongated

morphologies, as opposed to control ADSCs, which maintained their fibroblastic shape

(Section III.2.1.3). Additionally, when harvested with accutase and viewed on a haemocytometer

under an optical microscope, two important observations were made: firstly, the cells exposed to 7.5

and 10 μM were still plate-adherent after accutase treatment; secondly, the unattached cells exposed

to 7.5 and 10 μM of 5-AZAd displayed a very thin and elongated morphology, instead of the typical

round shape adopted by MSCs upon harvesting and myoblasts after enzymatic digestion treatment,

and stained positively for trypan blue (Figure III.4). Both observations represent abnormal cell

behaviours, suggesting that 5-AZAd at 7.5 and 10 μM induced some cell cytotoxicity.

39

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.

At day 7 the cells were harvested with 0.05% trypsin for 5 minutes to evaluate if a harsher

enzymatic digestion protocol would efficiently detach the cells treated with 7.5 and 10 μM of 5-AZAd

from the plates. No changes in cell detachment efficiency were observed, as these cells remained

adherent to the plates. At day 14, 0.25% trypsin was tested but with the same outcome. It was

concluded that for future studies, a harsher enzymatic formulation should be used to detach 5-AZAd

treated cells, such as TrypLETM (1X). Interestingly, accutase and trypsin were used for cell detachment

in several reviewed studies and there was no reference in the literature to unusual effects of 5-AZAd in

cell adherence to plastic or trypsin resistance. Thus, it is possible that increased cell attachment could

be a consequence of unspecific effects induced by the demethylation agent used herein.

A pronounced decrease in cell viability was confirmed by cell counting for 5-AZAd at 7.5 and

10 μM, for which no viable cells were observed after day 1 of 5-AZAd treatment. In fact, in these

conditions, all cells stained positively for trypan blue at days 3, 5, 7, 12 and 14 after 5-AZAd

exposure (Figure III.5).

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.

A B

40

No plated cells could be observed for the 5 μM of 5-AZAd at days 3, 5, 7, 12 and 14, probably

due to an error in the initial plating of the cells, therefore, the cell viability for this concentration could

not be evaluated. Control cells and cells exposed to 2 μM of 5-AZA were similar in terms of cell

morphology and viability. Regarding the two cell densities tested, the results showed that with

5000 cells/cm2 higher cell viabilities were achieved in comparison with 3000 cells/cm2 (Figure III.5).

From the results obtained in this assay, two hypotheses were formulated: (i) 5-AZAd at

concentrations above 7.5 μM induced early cell apoptosis, between 24 and 72 hours after 5-AZAd

treatment; (ii) 5-AZAd at concentrations above 7.5 μM induced abnormal cytoskeleton alterations that

affected cell morphology. These two hypotheses were tested through a cell apoptosis assay and a

phalloidin staining assay, respectively.

III.2.1.2 Cell Apoptosis Assay

P1 ADSCs were plated on 12-well plates, exposed to 2 and 10 μM of 5-AZAd for 24 hours and

subjected to a propidium iodide (PI)/Annexin V flow cytometry analysis to prove 5-AZAd at 10 μM

induced early cell apoptosis. This assay gives more information than the trypan blue exclusion test

(which only detects viable and necrotic cells), since the PI/Annexin V flow cytometry assay allows

viable, apoptotic and necrotic cells to be distinguished through the use of FITC-labelled annexin and

PI molecules. When cells start becoming apoptotic, phosphatidyl serine phospholips flip to the

extracellular surface of the cell, and annexin can bind to them, thus detecting early apoptotic cells.163

PI, a fluorescent dye that binds to DNA, enters the cell and binds to DNA when the cell is late

apoptotic and the membrane is no longer intact, thus allowing the detection of necrotic cells.163

After 2, 7 and 14 days of 5-AZAd treatment the cells were analysed and it was possible to

discriminate between viable (Annexin V-/PI-), apoptotic (PI-/Annexin+) and necrotic (PI+/Annexin-,

PI+/Annexin+) cells (Table III.3 and Figure III.6).163

After day 2 of 5-AZAd treatment, the cells exposed to 10 μM were necrotic (92%), while most

of the cells exposed to 2 μM were viable (89%), as well as the control cells (94%). These results prove

that 10 μM of 5-AZAd induces early cell apoptosis even before 48 hours after 5-AZAd treatment, as

the cells were already necrotic at this timepoint. Moreover, since at this concentration only 8% of the

cells were viable, cell apoptosis was not assessed for this 5-AZAd concentration at days 7 and 14,

since at least 1 000 cells (gated) are required for an accurate flow cytometry analysis to be performed.

The number of cells rose as the number of days in culture increased, indicating that the cells

proliferated throughout the assay. Cells treated with 10 μM 5-AZAd did not achieve the minimal

number of gated events for flow cytometry analysis (Table III.3). Furthermore, the dot plot chart

obtained for this population of cells indicated that it was mostly composed by cell debris, since this

population was located on very low values of forward scatter (FSH, i.e. measure of cell size) and side

scatter (SSH, i.e. cell complexity) (Figure III.7). This was also an indicator that the cells were no longer

viable.

41

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.

Day 2 Day 7 Day 14

Control 2 μM 10 μM Control 2 μM Control 2 μM

Necrotic (%) 5.7 5.2 92 4.0 4.9 2.0 1.8

Apoptotic (%) 0.67 6.1 0 0.72 0.84 0.29 0.46

Viable (%) 94 89 8.0 96 94 98 98

Gated Cells 9564 9417 88 9387 9389 8656 8496

Figure III.6 Effect of 5-AZAd on cell viability.

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.

These results indicate that it is not viable to treat MSCs with concentrations of 5-AZAd equal

or above 10 μM, since it leads to early cell necrosis. Few studies regarding the effects of 5-AZA on cell

viability were found in the literature. The 50% inhibitory concentration of 5-AZA after 24 hours of

induction was reported to be around 40 μM, with little decrease in viability at concentrations below

10 μM.130 The fact that cells treated with 10 μM of 5-AZA have been successfully used in the literature

in flow cytometry analysis also suggests that 5-AZA might be significantly less toxic than the analogue

used herein. Still, 5-AZA can be cytotoxic, as Tomita and colleagues reported that BM-MSCs

A B

42

incubated with concentrations of 5-AZA above 20 μM not only appeared abnormal but also more than

50% necrosed.127 Additionally, the authors observed damage in BM-MSCs cultured with 5 and 10 μM

5-AZA for 24 hours.127

III.2.1.3 Microscopic Analysis of Cell Morphology

Control ADSCs (Figure III.8A) and ADSCs exposed to 2 μM of 5-AZAd maintained a

fibroblast-like shape, while ADSCs exposed to 10 μM appeared unhealthy and displayed very

heterogeneous morphologies. Essentially two types of cells were observed: very thin and elongated

cells and broad, irregular cells, sometimes with multiple cytoplasmic projections (Figure III.8B). Some

multinucleated cells could be observed in all conditions.

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).

It has been described that 5-AZA-treated BM-MSCs (3, 5 and 10 μM of 5-AZA) show more

heterogeneous morphologies and alterations in size and growth properties.133 In particular, the

presence of two types of cells, very thin and elongated vs. broad and irregular, is in accordance with

this previous report.133 Although the authors observed that the thin elongated cells were able to

replicate faster and form colonies, no relation between these two types of morphologies and cell

function, growth or differentiation has been established.

III.2.1.4 Phalloidin Immunofluorescence Staining

In order to visualize possible alterations in the cytoskeleton morphology of cells exposed to

5-AZAd, FITC-phalloidin and DAPI were used to stain their cytoskeleton (F-actin) and nuclei,

respectively. Control cells and cells treated with 2 μM 5-Azad (Figure III.9) were similar in terms of

morphology and confluence throughout the culture time. Very thin cells could be observed 2 days after

5-AZAd treatment with 10 μM, which is in accordance with the observed cell morphology under the

A B

43

optical microscope. No fluorescence was detected at days 7 and 14 after 5-AZAd treatment (10 μM),

probably due to cell necrosis, as shown in the cell apoptosis assay. These results indicate that after 48

hours of 10 μM 5-AZAd treatment severe cell morphology alterations are induced.

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.

III.2.1.5 Surface Marker Expression

The expression of surface markers was assessed by flow cytometry at days 2, 7 and 14 after

5-AZAd treatment. Cells treated with 10 μM of 5-AZAd did not achieve sufficient cell numbers for a

reliable flow cytometry analysis at day 2 after treatment (Table III.4). As the cell apoptosis assay

indicated that the cells exposed to this concentration of 5-AZAd were no longer viable, their surface

marker expression was not assessed at days 7 and 14.

The expression of the following markers was evaluated (Table III.4 and Figure III.10): CD73,

CD90, CD105, CD34, CD45 and CD56 (neural adhesion molecule, expressed in skeletal muscle).

CD56 was evaluated as a marker of myogenesis, in order to assess if 5-AZAd induced myogenesis.164

44

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.

Control 2 μM 10 μM

Day 2 (%) Day 7 (%) Day 14 (%) Day 2 (%) Day 7 (%) Day 14 (%) Day 2 (%)

CD73 100 100 100 100 100 100 83

CD90 98 90 92 99 87 72 51

CD105 100 93 96 100 72 92 79

CD34 11 8.4 4.5 10 6.2 3.8 0.89

CD56 0.37 0.19 0.10 0.13 0.11 0 0

CD31 0.23 0.16 0 0.82 0.29 0.12 0

CD45 0.32 0 0 0 0 0.09 17

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.

5-AZAd-treated cells decreased the expression of MSC markers CD73, CD90 and CD105.

This decrease was observed 2 days after treatment for ADSCs treated with 10 μM of 5-AZAd, but only

after day 7 for cells treated with 2 μM of 5-AZAd. Moreover, a significant increase in CD45 expression

was detected in ADSCs treated with 10 μM of 5-AZAd (17%). One study showed that MSCs

expressed CD45 after 5-AZAd induction (1 μM) through immunohistochemical staining and real time

PCR techniques.165 However, this increase in CD45 is probably not associated with haematopoietic

lineage differentiation but rather with apoptosis, since CD45 is thought to have a role in apoptosis

regulation.166 Control ADSCs and ADSCs treated with 2 μM of 5-AZAd expressed CD34 at day 2 after

5-AZAd treatment, and the expression of this marker decreased as the culture days increased.

Importantly, no CD56 expression was detected, suggesting that 5-AZAd did not induce

myogenesis. Still, many reports claim that 5-AZA induces MSCs to differentiate into myogenic

lineages, although contradictory conclusions were found amongst the studies present in the literature

(see Section I.2.1.3 – 5-Azacytidine to Induce Myogenesis).67,124–132,134 The fact that 5-AZA is being

studied for both cardiomyocyte and myogenic differentiation proves this lack of consistency.

Furthermore, the results are difficult to compare, as different research groups use different culture

media compositions and distinct markers to induce and prove myogenic differentiation, respectively.

45

Importantly, although described in the literature as being effective in inducing myogenesis, the

treatment of MSCs with 5-AZA presents one great disadvantage: the unspecific mechanism of action

of this antimetabolite. In fact, the mechanism through which 5-AZA induces myogenesis is not yet

understood, although some studies have suggested potential mechanisms for its action, such as

inducing the demethylation and activation of a myogenic determination locus that is usually

transcriptionally inactive.167

Wakitani and co-workers were the first to report that an immortalized line of rat BM-MSCs

could be differentiated into contractile muscle cells by using 5-AZA (concentrations ranging from 3 to

50 μM) or 5-AZAd (from 0.1 to 10 μM). Interestingly, of the concentrations tested, 0.3 μM of 5-AZAd

gave the highest incidence of myogenic conversion, according to the authors.67 Analogously, in 1999,

Makino and colleagues successfully differentiated immortalized rat BM-MSCs into contractile

cardiomyogenic cells expressing desmin, α-MHC and α-actin using 3 μM 5-AZA.128 Since then, 5-AZA

and its deoxy analogue have been widely tested in immortalized and primary cell lines.

Liu and collaborators concluded that rat BM-MSCs could not be induced to differentiate into

cardiomyocytes using 5-AZA unless the cells were immortalized.133 This is a plausible hypothesis,

since immortalized cell lines are more prone of overcoming the toxic effects of molecules such as

5-AZA than primary cell lines, and could eventually be reprogrammed into myogenic differentiation in

response to 5-AZA. It is also possible that the cells differentiate due to a random reprogramming of

gene expression induced by the drug, rather than to the cell’s ability towards myogenic

differentiation.133 As a cytidine analogue, 5-AZA is capable of being incorporated into the DNA,

subsequently acting by covalently inhibiting DNA cytosine methyltransferases. From a regulatory

perspective, it would be very challenging to approve a treatment modality involving the use of cells

exposed to a molecule that can potentially give rise to unpredictable effects, such as toxicity or

tumorigenicity. This demethylation agent may cause an unregulated expression of various genes,

disabling its use in therapeutic contexts, in which the cells cannot present any kind of unexpected

behaviours.168 Consequently, research in the field of muscle engineering using MSCs as a source of

muscle-like cells should focus on the establishment of novel protocols adequate for clinical

applications.

In conclusion, in the experiments presented herein, 5-AZA did not induce myogenesis and at

10 μM led to early cell necrosis and immunophenotypical and cell morphology alterations. The study

presented herein has limitations such as lack of statistical relevance, as only one population of ADSCs

was tested immunophenotypically, and the fact that only one marker was used to detect myogenesis

(CD56 expression). Still, the results presented in this thesis give a new insight about the effects of

5-AZAd on cell apoptosis and morphology. Moreover, it is possible that 5-AZAd affects the expression

of cell adhesion molecules, as standard enzymatic digestion procedures were not sufficient to detach

5-AZAd-treated cells from culture plates, a phenomenon that was not described in the reviewed

literature.

46

III.2.2 Myogenic induction of CD34+ cells from the Stromal Vascular Fraction

One particular study confirmed the presence of cultured Pax7-expressing AT-SVF cells that

spontaneously differentiate into myotube-like cells in the presence of myogenic differentiation medium

alone.110 However, their percentages within the SVF were reported to be very low (0.001% of total

plated cells) and their characterization concerning marker expression is not known, making it difficult

to isolated them. Moreover, an extensive literature search indicated that myogenic-committed cells

have not yet been identified in uncultured SVF. Thus, with the aim of assessing if myogenic-committed

cells exist in the human white AT, intracellular flow cytometry staining for Pax7 was performed in a

fresh SVF sample in order to identify Pax7-expressing cells (Figure III.11).

100

101

102

103

104

Pax7 (FITC)

0

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Pax7 (FITC)

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Pax7 (FITC)

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6.23

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.

Three subpopulations were identified in the dot plot chart (Figure III.11A), which were not

evenly enriched with Pax7+ cells. Subpopulation 2, placed at low FSH and intermediate SSH values,

corresponding to cells with low size but with some complexity, was the most enriched population

(7.7%), followed by subpopulation 3 (5.3%) and subpopulation 1 (2%). The identification of a

considerable percentage of Pax7-expressing cells in the SVF, which translated into numbers

corresponded to 1.05x105 cells in a total of 49.6x105, distributed into 3 gated subpopulations,

remarkably indicates that myogenic-committed cells do exist in vivo in the adult AT and that these

cells can potentially be isolated. Therefore, a method for further isolating Pax7+ cells from the SVF

was aimed. However, as it is not possible to isolate/sort cells based on the expression of intracellular

SVF Subpopulation

1 2 3

Gate Frequency (%) 41.9 8.57 11.7

Gated events 5912 1210 1653

Pax7 expression (%) 2.0 7.7 5.3

Pax7-expressing Cells (total, 105)

1.05

C

A

B

47

markers, these cells could not be sorted according to Pax7 expression. An extracellular maker was

thus needed for this purpose.

Several protocols for satellite cell/skeletal muscle stem cell isolation have been successfully

established. Among them, satellite cell sorting through CD34/CD56 positive selection is a common

method that has allowed the isolation of satellite cells with different commitment levels and

differentiation potentials.156,157 In particular, it has been shown that CD34-CD56+ satellite cells are

purely myogenic, while CD34+CD56+ satellite cells are also capable of adipogenic differentiation.157

Based on this information, enzymatically processed SVF cells were submitted to CD34 cell sorting

through MACS with the aim of potentially isolating Pax7-expressing cells. This methodology was

repeated and the obtained cells were used for several assays (Table III.5). The cells obtained after

plating will be designated here as AT-SVF cells, as standard FBS was used for their isolation.

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.

Experiments Performed

SVF Sample Reference

CD34/CD56 Expression

Pax7 expression MACS Low attachment plate cultivation

Myogenic Differentiation Assay

SVF-1 - Yes - - -

SVF-2 Yes - Yes - -

SVF-3 Yes - Yes - Yes

SVF-4 - Yes Yes Yes -

Cell pool* Yes - Yes - Yes

III.2.2.1 Expression of myogenic markers in SVF cells before and after MACS

Fresh SVF cells were subjected to MACS in order to sort cells according to CD34 expression

(Table III.6). CD34 and CD56 expression was then determined for the CD34 positive and negative

fractions through flow cytometry analysis. The term “CD34-enriched” and “CD34-depleted” refers to

the initial status of fresh cells (and not to cells after passages in culture). The term “CD34+ cells” refers

to cells that effectively expressed CD34 (confirmed by flow cytometry). Of notice, CD34 expression

decreased substantially in the first passages in culture.

The yield of the sorting process could be obtained for two of the sorted SVF samples. This

yield represents the number of CD34+ cells present in the CD34-enriched fraction divided by the

number of CD34+ cells present in the initial SVF sample. Both values were calculated by multiplying

the total number of cells by the corresponding percentage of CD34+ cells (obtained by flow cytometry

analysis). The yield values obtained (33% and 58%) seem reasonable, when compared to recovery

values obtained for the same target molecule and MACS system in the literature (78% after one cycle

and 52% after two cycles, for umbilical cord blood cells).169

48

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.

SVF Sample Reference

Total Cells (106) Before MACS

CD34+ cells (106) Before MACS

Cells (106) CD34+ fraction

CD34+ cells (106) After MACS

Purity (%)

Cells (106) CD34-

fraction

MACS Yield (%)

SVF-2 8.6 - 1.6 1.5 94 3.4 -

SVF-3 5.2 3.0 1.3 1.0 77 2.0 33

SVF-4 4.8 - 0.57 - - 0.33 -

Cell pool 4.1 2.0 1.5 1.2 80 0.60 58

After the sorting process, CD34 and CD56 expression was assessed in the CD34-enriched

and depleted fractions obtained for samples SVF-2, SVF-3 and the cell pool (Figures III.12-III.14).

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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.

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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.

CD34-enriched CD34-depleted

% MFI % MFI

CD34 98 90 23 3.4

CD56 8.6 2.0 0.20 1.0

Gated events

16 802 21 112

Unsorted

SVF CD34-enriched CD34-depleted

1 2 1 2

% MFI % MFI % MFI % %

CD34 77 26 97 75 89 32 1.6 14

CD56 32 3.4 94 11 33 5.3 11 0.70

Gated events

931 6449 1713 1208 918

CD34-depleted

1 (%) 2 (%)

CD90 3.1 14.1

CD73 7.2 3.8

CD31 32.3 20.4

CD45 96.2 36.9

CD105 1.4 1.5

A B C

D

1

2

A B C

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E

Unsorted SVF

CD34-depleted CD34-enriched

CD34-depleted CD34-enriched

D

49

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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).

Over 70% of cells were CD34+ in the two samples of unsorted SVF analysed by flow

cytometry, while CD56 expression was more divergent (32% and 0.50%). CD34 expression in the

enriched cell fractions reached values around 90%. Some CD34 expression was detected in the

negatively sorted fractions, which may have lowered the yield of the separation process. Still, it is

possible that these cells are CD34dim (i.e. expression slightly increased when compared to the

negative control). Since the magnetic sorting process depends on the number of magnetically-labelled

antibody molecules bound to the desired target protein (which translates into stronger or weaker

retentions in the column), cells with fewer CD34 molecules on their surface are more prone to be

washed from the column.

The results shown in Figure III.13E indicate that hematopoietic cells are the main constituents

of the CD34 negative fraction. In one of the positively sorted samples, a subpopulation of cells positive

for both CD34 and CD56 was obtained (Subpopulation 1, Figure III.13B). To the extent of the literature

reviewed herein, besides satellite cells, only myoendothelial cells are known to simultaneously

express CD34 and CD56.170,171 These results indicate that a population with myogenic potential may

have been sorted by performing CD34 MACS. However, the results show clear donor-to-donor

variability in regard to the obtained populations and their levels of marker expression.

Based on these results, and with the aim of determining if sorting CD34 positive cells is, in

fact, a method of enriching the percentage of Pax7-expressing cells from the SVF, MACS was

performed and the levels of Pax7 were assessed before and after the process, for both

CD34-enriched and depleted fractions (Figure III.15). The results indicate that sorting CD34-positive

cells allowed the enrichment of Pax7-expressing cells: the expression of this phenotype increased

over 7-fold in the CD34 positive fraction (4.3%) in comparison to the initial SVF sample (0.56%). On

the other hand, no significant Pax7 expression was detected in the CD34 negative fraction, which

suggests that most Pax7-expressing cells present in the tested SVF sample express CD34, indicating

that they may hold an immature satellite cell-like phenotype (CD34+CD56+).

Unsorted SVF

CD34- depleted

CD34-enriched

1 2

% MFI % % MFI % MFI

CD34 75 28 6.6 93 80 85 30

CD56 0.50 1.8 0.40 6.8 2.6 3.1 1.7

Gated events 5217 5390 6327 3047

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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.

III.2.2.2 Plating and isolation of CD34-enriched and CD34-depleted fractions

The cell fractions obtained previously were plated onto tissue culture flasks with the aim of

assessing if distinct cell populations could be obtained in the two fractions and if the levels of CD34

and CD56 were maintained after plating. Additionally, cells from both fractions were plated in the

presence of gelatin and fibronectin substrates (i.e. coating of the plastic culture surface), to promote

the adhesion of myogenic-committed cells, and in the presence of high glucose DMEM (DMEM-HG)

medium, to test if increased glucose concentrations promote cell differentiation.172 The cells were

plated into the following conditions: a) non-coated dishes, b) gelatin-coated dishes,

c) fibronectin-coated dishes and d) fibronectin-coated dishes in the presence of DMEM-HG medium.

Unsorted SVF cells were plated onto non-coated dishes.

An expansion step was performed in the presence of myogenic expansion medium (DMEM

supplemented with standard FBS, not MSC-qualified, dexamethasone and bFGF)157 and the levels of

CD34 and CD56 were determined after the first passage.

Surprisingly, no cells derived from the CD34-depleted fraction adhered to tissue culture plates,

even in the presence of a gelatin and fibronectin coating. 48 hours after plating, only floating cells

were observed and after 7 days in culture no cells were adherent, and so the cells were discarded.

The assay was repeated to account for biological variability and the same outcome was obtained.

Moreover, plated cells from the CD34-enriched fraction displayed similar sizes and morphologies

when compared to unsorted SVF cells. Based on both these observations, it was hypothesized that

Unsorted SVF

CD34- Enriched

CD34-depleted

1 2

Gate Frequency (%)

39.1 53.6 35.9 19.5

Gated events 6232 10381 2224 4098

Pax7 Expression (%)

0.56 4.2 0 0.26

Pax7-expressing cells (104) 2.72 2.39 - 0.02

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Unsorted SVF CD34-enriched

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51

only the CD34+ cells within the unsorted SVF were able to adhere and proliferate in culture. This

hypothesis was further tested in Section III.2.2.4.

The results presented below (including in Section III.2.2.3) correspond to cells initially

belonging to a cell pool of SVF cells (Figure III.14). After 10 days in culture, unsorted SVF cells were

completely confluent, while cells from the CD34-enriched fraction were not evenly spread across the

plates and grew as dense colonies (Figure III.16). Since unsorted SVF cells were not submitted to the

MACS pre-separation filter, which has 30 μm diameter pores, large cell aggregates could have been

initially plated in this condition, which may have contributed to the achievement of high confluence

levels earlier.

In low confluence areas, pericyte-like cells could be observed (Figure III.16). The expression

of CD34 and CD56 was determined after the first passage (Figure III.17). For all conditions, a main

population and a subpopulation of cells (placed at lower SSH and FSH values) could be identified. The

results suggest that the presence of coating had great influence on CD56 expression, since cells

cultured on gelatin and fibronectin-coated plates presented CD56 expression levels around 80%, while

SVF (unsorted cells) and non-coated cells presented negligible levels of CD56 expression.

Conversely, CD34 expression was more variable between the tested conditions. Cells cultured on

fibronectin-coated plates exposed to DMEM-HG medium presented the lowest levels of CD34

expression (0.8%), while non-coated cells presented the highest (21%). The high levels of CD56

expression (and contrastingly lower CD34 expression) might indicate that the cells adopted a

phenotype similar to activated satellite cells, which tend to lose CD34 expression and are

characterized phenotypically as CD34-CD56+. This experiment was repeated with cells from a different

donor (SVF-2), but significantly lower levels of CD34 and CD56 expression were obtained. One of the

factors that might have influenced these results is cell confluence and/or plating density, since

different confluence levels were achieved in the two assays. Thus, the assay needs to be repeated to

account for these factors.

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.

A

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CD34-enriched fraction-derived cells

SVF No coating Gelatin Fibronectin LG Fibronectin HG

% MFI % MFI % MFI % MFI % MFI

CD34 (MP) 7.9 2.0 21 2.9 9.3 1.8 14 2.3 0.80 1.2

CD56 (MP) 0 - 0.20 1.7 90 7.1 84 20 80 5.1

Gated events 3317 7103 6085 2285 2835

CD34 (SP) 21 2.4 17 4.1 5.2 1.0 0 - 0 -

CD56 (SP) 0.60 1.2 0 - 54 4.4 20 4.8 4.8 4.0

Gated events 414 1112 656 367 657

Total cells (106) 2.48 0.57 1.83 0.44 1.76

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).

III.2.2.3 Myogenic Differentiation Assay

Myogenic induction was performed by replacing the plating medium by DMEM-HG

supplemented with 2% HS, a medium commonly used to induce myogenic induction in MSCs. To

determine at which stage of myogenesis the cells were, four stage-specific markers expressed in the

myogenic differentiation process were used: Pax7 (expressed in satellite cells), MyoD (expressed in

proliferating and differentiating myoblasts), myogenin (expressed only in differentiating myoblasts) and

skeletal muscle MHC (expressed in differentiated myotubes). After 2, 4 and 8 days of medium

replacement, the expression of these markers was evaluated through intracellular flow cytometry and

immunostaining methodologies.

The expression of myogenic markers before medium replacement was evaluated on cells

cultivated on gelatin- and fibronectin-coated (HG) plates, the only two conditions for which enough cell

numbers were obtained to perform intracellular staining for flow cytometry analysis (Figure III.18).

Surprisingly, the results indicate that cells cultured on gelatin- and fibronectin-coated (HG)

plates expressed MyoD and myogenin (over 97% and 84% of the gated main population, respectively)

before the medium replacement. Moreover, the MFI values show that MyoD was being significantly

expressed, as well as myogenin, although at lower levels.

SVF No coating Gelatin Fibronectin

(LG) Fibronectin (HG)

53

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Main Population Subpopulation

Gelatin Fibronectin (HG) Gelatin Fibronectin (HG)

% MFI % MFI % MFI % MFI

Pax7 7.3 1.7 32 2.2 0.10 1.1 1.4 1.3

MyoD 98 89 99 92 76 14 92 20

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)


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



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 (%)


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

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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

Subpopulation 1 Subpopulation 2 Subpopulation 1 Subpopulation 2 Subpopulation 1 Subpopulation 2

% MFI % MFI % MFI % MFI % MFI % MFI

CD34 91 36 45 4.8 94 42 82 11 98 42 82 11

CD56 15 2.8 0 - 8.9 3.3 0 - 19 3.0 0 -

Gated events 1911 1255 1205 1361 3093 4008

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.

III.2.2.5 Culture of SVF-derived cells on low attachment plates

Satellite cells usually require a substrate to adhere to, given the fact that they adhere slower

to culture dishes when compared to other cell types such as MSCs.110 Once satellite cells adhere,

they quickly begin differentiating into myoblasts, instead of proliferating in an undifferentiated state.

MSCs, on the other hand, start proliferating greatly once they adhere, making it difficult to isolate more

quiescent cells that may exist in the SVF (e.g. satellite cells). Therefore, in an attempt to isolate

myogenesis-committed cells from the SVF, cells derived from fresh SVF and cells from the

CD34-enriched fraction sorted by MACS were plated onto low-attachment plates. The use of

low-attachment plates for satellite cell expansion has been implemented for mice Pax7+ cells (after cell

isolation through FACS) with success, as Pax7+ cells were able to proliferate and form aggregates in

suspension.176 Two types of medium were tested: the expansion medium mentioned above and a

commercially available skeletal muscle cell growth medium formulation. After 6 days of culture on low

attachment plates, aggregates in suspension could be observed. The medium was then collected and

cells in suspension were plated on gelatin-coated plates. Adhesion of the aggregates to the culture

SVF – Day 2

Skeletal Muscle Medium

Expansion Medium

% MFI % MFI

CD34 89 45 86 47

CD56 35 8.6 9.3 8.3

Gated events 3196 4191

SVF D2

Expansion Medium

SVF D2 Skeletal Muscle

Medium

SVF D3

Skeletal Muscle Medium

SVF D3

Expansion Medium

CD34-enriched D3

Expansion Medium

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59

plate was observed after 24 hours in culture. Aggregates were formed by typical AT-SVF cells that

were able to proliferate in suspension, and no satellite/myoblast-like cells were observed

(Figure III.23). The cells were harvested after 4 days and the expression of CD34 and CD56 was

assessed by flow cytometry. No CD56 expression was obtained in any of the conditions, suggesting

that myogenic-committed cells were not isolated.

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.

III.2.2.6 Discussion

Most of the data in the literature indicate that MSC myogenic differentiation requires either

DNA demethylation treatment, co-culture with myoblasts or in vivo muscle engraftment. As

demonstrated in Section III.2.1, DNA demethylation treatment, under the conditions tested in this

thesis, did not reveal to be a plausible option. Conversely, few reports of spontaneous myogenic

differentiation of cells derived from the SVF, without any of the previously mentioned stimuli, have

emerged.69,110–113 Some of these studies demonstrated that the SVF contains subpopulations of cells

that exhibit myogenic phenotypes in vitro and that are capable of differentiating into myotubes.110,111

When reading the existing literature on this matter, one important question can be raised: does the

myogenic potential of the AT relies, in fact, on the presence of a low percentage of

myogenic-committed cells, as concluded by these authors110,111, or is the multipotent adherent fraction

of the SVF (e.g. ADSCs) capable of myogenic potential per se in the presence of adequate stimuli?

The results obtained herein give evidence that both hypotheses might be correct.

To the extent of the literature reviewed herein, this is the first time that Pax7-expressing cells

are identified in uncultured fresh human SVF samples. Thus, it is reasonable to claim that

myogenic-committed cells do exist in vivo in the adult AT, as Pax7 is the most reliable marker for

identifying skeletal muscle progeny. Still, this assay should be repeated in order to account for

biological variability and to assess the average percentage of Pax7+ cells in the SVF. Additionally, it

would be interesting to determine if MyoD- and myogenin-expressing cells exist in fresh uncultured

SVF, in order to conclude if myogenic cells at different stages of commitment can be obtained from the

AT. Although the isolation of Pax7+ cells was not successful, even after the implementation of two

different strategies, these results open a new window of clinical applications for the AT. For instances,

if sufficient numbers of myogenic cells are accessible from the AT and an efficient isolation method is

60

developed, the SVF could become a reliable alternative source of skeletal muscle cells, thus

eliminating the need of invasive muscle biopsies. Moreover, it has been hypothesized that the

immunosuppressive properties of undifferentiated ADSCs increase the feasibility of transplantation

procedures in damaged muscle, which represents an important advantage in a clinical therapy

perspective.177

Although CD34-cell sorting through MACS enhanced the content of Pax7+ cells by 7-fold,

other markers should be used for isolating these cells, as the results obtained herein indicate that over

80% of SVF cells that adhere to tissue culture plates in the first 48 hours of culture are CD34+. Hence,

for future studies, CD56 should be a promising marker for sorting myogenic cells from the SVF, as

AT-derived MSCs do not express this marker.178 FACS could have also been used as sorting

methodology, instead of MACS. Indeed, FACS allows the achievement of highly pure fractions, and

holds greater precision, resolution and sensitivity, which are important qualities when the isolation of

rare subpopulations is aimed. Importantly, an anti-CD56 antibody attached to a bright fluorophore

(e.g. PerCP) should be chosen, in order to allow the isolation of cells expressing different amounts of

the antigen (e.g. CD56bright and CD56dim cells). The use of antibodies with inferior relative brightness,

such as FITC-conjugated antibodies, may result in less efficient cell separations, especially if the

desired cell population does not significantly overexpress the chosen antigen.179

According to the ISCT, uncultured SVF cells express CD34 at moderate levels (>20%), and lose

CD34 expression over few cell passages108. Three uncultured SVF samples were assessed herein by

flow cytometry and considerably high levels of initial CD34 expression were obtained (75, 77 and

98%). Moreover, in the first 48 hours of culture, CD34+ cells represented over 80% of total cells

(depending on the culture medium used), and even higher values of expression were reached at 72

hours of culture. These results are in accordance with the observation that the CD34-depleted cell

fraction obtained by MACS lacks proliferative potential. One similar study was performed by Maumus

and colleagues, who sorted CD34+ cells from human AT by MACS and concluded that only CD34+

cells possessed colony-forming activity.180 These results indicate that the proliferative potential of the

SVF relies on CD34+ cells, which deeply contrasts with UC-derived MSCs and BM-MSCs that do not

express CD34. Additionally, the maintenance of CD34 and CD56 expression seemed to greatly

depend on the culture medium and the presence of a substrate (i.e. coating). In Table III.2, it was

observed that low serum medium stimulated CD34 expression in P0 AT-SVF cells. From the results

obtained in Figure III.17, no clear correlation between cell sorting, presence/absence of substrate and

CD34 expression could be made. However, CD56 expression was clearly augmented in the presence

of substrates (gelatin or fibronectin).

Cells plated on a coated plastic surface adopted a phenotype similar to activated satellite cells

(CD34-CD56+) which is consistent with the high MyoD and myogenin levels observed for cells cultured

on gelatin- and fibronectin- (LG) coated plates before cell re-plating. These results suggest that the

expansion medium in which the cells were isolated might have positively influenced the cells’

commitment to the myogenic lineage. Moreover, it can be proposed that AT-SVF cells possess some

61

myogenic potential per se and begin expressing myogenesis-related proteins in the presence of

adequate biochemical stimuli (e.g. dexamethasone and bFGF). In fact, dexamethasone

supplementation is known to promote myogenic differentiation in BM-MSC, leading to the formation of

desmin-positive myotubes in a dose-dependent manner, and enhance the proliferation and

differentiation of myoblasts.106,181,182 Concordantly, a study showed that significantly higher numbers of

MSC nuclei were involved in myotube formation when bFGF and dexamethasone were added to

co-cultures with myoblasts when compared to DMEM medium supplemented with HS alone, thus

suggesting a possible role of dexamethasone and bFGF as pro-myogenic factors.104 However, it

should be noted that dexamethasone is not a specific pro-myogenic factor, as it also promotes

adipogenic, osteogenic and chondrogenic differentiation in MSCs.170 In future studies, the cells should

be expanded in medium free of these factors, in order to allow the proliferation/maintenance of the

cells in an undifferentiated state and avoid early myogenesis commitment.

The presence of gelatin coating was reported to promote myoblast differentiation presumably

by collagen interactions mediated by a myoblast cell surface glycoprotein named gp46, which is

reduced in differentiation-defective mutants of rat skeletal myoblasts.183 The expression of gp46

remains constant during myoblast replication and fusion but decreases markedly postfusion, which is

in accordance with the pattern of collagen synthesis in L6 myoblasts (peaking just as cells align and

dropping after fusion).183,184 Through the use of a compound that specifically inhibits collagen

hydroxylation, it was concluded that collagen-deficient L6 myoblasts were unable to align and fuse if

the compound was added before myoblast alignment.185 These evidence suggest that collagen is

essential at early stages of myogenic commitment but dispensable for the subsequent steps of

myogenic differentiation. Still, in a more recent study in which the role of fibronectin and gelatin

coating on myogenesis was explored, it was concluded that fibronectin promoted myoblast migration

and more organized collective alignment, and gave rise to a larger number of myotubes, with more

nuclei per myotube, when compared to gelatin.186 Conversely, other authors observed no significant

differences in myotube development in myoblasts cultured on gelatin and fibronectin coated plates in

low serum conditions.187 Moreover, these authors concluded that exogenous matrix substrates are not

essential for serum free cultures if matrix secretion is triggered and the culture surface supports the

retention of secreted matrix proteins.187

In the conditions of the experiments presented herein, the presence of coating had impact on

myogenic marker expression, as gelatin coating increased skeletal MHC expression by 10-fold after 8

days of induction (34% versus 3.3% for cells in the absence of coating). Ultimately, it is possible that

the presence of different ECM molecules influences the progression of myogenesis by the exertion of

distinct effects at different stages (initial alignment and commitment versus myoblast fusion and

myotube formation). Thus, it would be interesting to attempt to mimic the skeletal muscle ECM

composition pattern and its dynamic changes when muscle regeneration occurs, in order to optimize

myogenesis in vitro.

62

Myogenin and MyoD expression were not maintained for long-term culture periods, and the

expression of both markers was drastically reduced after day 2 (gelatin and fibronectin HG conditions)

and 8 of media replacement, respectively. One particular study involving UC-derived MSCs showed

that, in the presence of dexamethasone and hydrocortisone, MSCs expressed MyoD at day 3 of

culture and myogenin at day 7, but both markers were not expressed after 2 weeks.102 It should be

noted that a switch in the secondary antibody used for MyoD at day 8 might have influenced the

results obtained, since MyoD expression was still observed by immunofluorescence at day 8, albeit at

lower intensities when compared to day 4. In order to avoid possible interferences caused by the

secondary antibodies, the use of labelled primary antibodies should be preferred in future studies.

Still, the drastic decrease in MyoD expression, which was not followed by myogenin

upregulation, might have had two possible causes. Firstly, MyoD decreases may be a consequence of

the increasing proportion of broad flattened cells with culture time, as lower fluorescence intensity was

observed in these cells when compared to high confluent areas filled with spindle-like cells. Secondly,

the myogenic induction medium used herein might not have sustained and promoted the last steps of

myogenic differentiation (cell fusion and myotube formation). Consequently, it is possible that

important factors might have been lacking in the medium formulation used. Common myogenic

differentiation medium formulations include DMEM supplemented with insulin and transferrin, or

hydrocortisone, dexamethasone and FBS, apart from HS, which is usually used in ranges from 2 to

5%.31,105,112,157 Importantly, DMEM-HG medium has been reported to not only promote adipogenic

differentiation in skeletal muscle and ADSCs but also inhibit myogenesis in C2C12 myoblasts, while

DMEM-HG and high insulin activate myoblast differentiation.173,188

Unsorted SVF cells demonstrated the same MyoD expression levels as sorted and coated

cells, and the same type of population was obtained for all conditions. Thus, CD34 sorting is probably

a dispensable step, which is a positive aspect since cell sorting is an expensive operation that often

leads to cell losses. In future studies, the myogenic marker expression of unsorted SVF cells cultured

on coated plates should be also tested.

One surprising result was the presence of two roundish multinucleated cells in a low

confluence area of a gelatin-coated well. Since skeletal muscle are the only kind of multinucleated

cells existing in the human organism, it can be postulated that myotube formation occurred in P0

(initial cell passage, upon initial cell seeding after isolation), and that these myotubes were unable to

elongate once re-plated. This atypical morphology could have been caused by cytoskeletal arrestment

caused by cell trypsinization and inadequate plating conditions (e.g. plating density and/or medium

composition). In fact, myotubes are known to undergo anoikis, a response induced by either a loss of

cell anchorage, anchorage to inappropriate ECM proteins or disrupted cell adhesion to the ECM,

consequently exhibiting spherical morphologies and eventually detaching from culture plates.189 In

particular, laminin–211 was reported to be important in preventing this phenomenon.189 The possibility

of myotube formation accentuates the notion that myogenesis progressed dynamically and rapidly at

early stages in culture, which contrasts with the current established myogenic differentiation protocols,

that can last up to 6 weeks in order to significantly detect skeletal MHC expression.69,102,112

63

Under forced MyoD expression, MyoD-expressing ADSCs were shown to have the capacity to

fuse with dystrophic myoblasts, restore dystrophin expression and to form fibers.190 In a very recent

study, MyoD expression was induced in rat ADSCs through the transfection of small activating RNA

targeting MyoD gene promoters, in a process known as RNA activation.191 Desmin expression could

be detected after 14 days of transfection, and was even more evident after 21 and 28 days.191 Herein,

natural MyoD expression was accomplished, which represents a major advantage when compared to

genetically modified or transfected cells for future clinical applications.

MyoD expression was detected not only in the nucleus but also in cytoplasmic areas.

Transcription factors are synthetized in the cytosol, and many are known to interact with cytosolic

proteins and translocate to the nucleus, where they exert biological effects. Therefore, it is plausible to

propose that MyoD is either (i) being arrested in the cytosol by unknown interactions with other

proteins, and that its translocation to the nucleus is only triggered once the cells receive enough

stimuli to proceed to myogenic differentiation or (ii) cytoplasmic arrestment is being caused due to

MyoD turn over (degradation) by the ubiquitin proteasome system.192

Lastly, to further enhance myotube formation, specific biomaterials or substrates with different

stiffnesses could be tested. ADSCs were shown to undergo stiffness-induced lineage commitment and

fuse (albeit at low percentages – 2%) when plated in a muscle-mimicking extracellular matrix (10 kPa)

surpassing BM-MSCs, which never underwent stiffness-mediated fusion.68 The same group of

researchers developed a mechanically-patterned hydrogel that led to increased fusion rates and

fusing cell numbers in comparison with unpatterned matrices.193 Thus, the importance of physical cues

in the form of ECM composition, stiffness and patterning, should not be underestimated.

III.3 Differentiation of Mesenchymal Stem/Stromal Cells into Smooth Muscle Cells

As already mentioned, human adult SMCs have limited proliferation capacity, unless the cells

are immortalized, which makes the ex vivo expansion of these cells for tissue engineering applications

very unviable.135 With the aim of achieving smooth muscle differentiation of MSCs from two different

sources (SVF and BM), PD98059 (a compound already successfully used in one other study for this

purpose)140 was tested. This compound belongs to the small molecules category, which are becoming

increasingly popular in stem cell self-renewal and differentiation studies, mainly because of their ability

to specifically act on proteins part of signalling pathways.194 PD98059 is a potent and selective

inhibitor of MAPKK, a key protein in the ERK/MAPK pathway. This pathway is believed to play an

important part in inhibiting the activity of SRF, a critical regulator of vascular SMC differentiation.140

A commercially available line of human bladder SMCs was used as positive control. These cells

were initially spindle-like but became larger and more rectangular in shape as time in culture

increased (Figure III.24). The expression of smooth muscle markers was tested in control bladder

SMCs by flow cytometry and immunofluorescence (Figure III.25).

64

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.

0 200 400 600 800 1000

Forward Scatter

0

200

400

600

800

1000

Sid

e S

catt

er

47.4

17.6

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.

The flow cytometry results obtained for marker expression in control cells surprisingly show

poor expression levels for SM-MHC (Figure III.25). Calponin and α-SMA expression was higher but did

Population

1 (%) 2 (%)

α-SMA 32 13

Calponin 51 28

SM-MHC 4.5 2.7

1

2

α-SMA

SM-MHC

Calponin

Calponin

Desmin

Desmin

Merge

Merge

A B

A B

C

65

not come close to 90-100% (51% and 32% for Population 1, respectively), which was the expected

expression range. Thus, desmin was included in the marker panel in immunofluorescence testing.

Although this intermediate filament is not smooth muscle specific, as it is expressed also in skeletal

muscle and cardiomyocytes, it can be used as an indicator of myogenic lineage differentiation. Low

fluorescence intensity could be detected for α-SMA and SM-MHC, while most cells clearly expressed

calponin and desmin.

P1 AT-SVF cells and P3 BM-MSCs isolated and plated in DMEM+10%FBS (standard FBS

and MSC-qualified FBS, respectively) were induced with DMEM+3%FBS supplemented with different

concentrations of PD98059, after reaching confluence. The cells displayed spindle-like morphologies

(Figure III.26) and were in culture for 6 days (BM-MSCs) and 9 days (AT-SVF cells) before induction.

Figure III.26. BM-MSCs (A) and SVF-derived cells (B) plated in DMEM+10%FBS, cultured for 3 and 9 days, respectively.

During BM-MSC expansion, P2 cells were tested for monocyte contamination by flow

cytometry with monocyte-specific marker CD14. The low percentage of expression obtained for this

marker (0.18 %) indicated that a monocyte-free population was achieved.

Cell morphology and viability did not suffer any significant changes from PD98059

supplementation. The effects of PD98059 in the expression of smooth muscle markers α-SMA,

calponin, SM-MHC and desmin were tested in the abovementioned cells after 2, 4 and 7 days of

induction. Different conditions were tested in the two assays. For BM-MSCs, two controls

(DMEM+10%FBS and DMEM+3%FBS) were implemented, and 1 μM, 10 μM and 30 μM of PD98059

were tested, based on the work of Tamama and co-workers.140 For AT-SVF cells, DMEM-HG medium

supplemented with 10 μM of PD98059 was added as a test condition, and DMEM-HG supplemented

with 3%FBS was the only control used. A medium change was performed at day 4.

The flow cytometry results obtained for BM-MSCs indicate that the maximum expression

values for α-SMA and calponin were reached at day 2 of induction, and the expression of these

markers gradually dropped. However, the expression levels obtained for 30 μM at day 2 were close to

the ones of the positive control (29 for α-SMA and 34 for calponin versus 32 and 51, respectively, in

A B

66

the positive control). For SM-MHC, little expression was obtained for 10 μM, 30 μM and

DMEM+10%FBS alone at day 2 (1.9, 1.3 and 1.0, respectively), which decreased to values close to 0

at days 4 and 7. Surprisingly, for AT-SVF cells no expression was obtained for α-SMA, calponin and

SM-MHC. It should be noted that due to a stock rupture the secondary antibody used for the AT-SVF

assay was different (in terms of the conjugated fluorophore) from the one used in the BM-MSCs

assay. Desmin expression, on the other hand, achieved its maximum at day 4 for most conditions.

Still, for the condition using 30 μM of PD98059, 22% desmin expression was reached at day 7. In

general, PD98059 concentration was not directly correlated with increased marker expression, apart

from BM-MSCs at day 2. Additionally, cells from the control conditions frequently presented

expression levels higher than cells exposed to PD98059.

Regarding the immunofluorescence results obtained, calponin- and desmin-expressing cells

could be observed in all conditions (Figure III.27). Importantly, fluorescent cells were only observed

near the edges of the wells, which suggests that 3-D-like spatial stimuli might promote myogenic

differentiation. Few cells were positive for both desmin and calponin. Calponin-expressing cells

presented broad polygonal morphologies and increased sizes, as opposed to desmin-expressing cells,

which tended to be smaller and more spindle-like. In BM-MSCs, very low intensity fluorescence could

be observed for α-SMA and SM-MHC at day 2 in cells near the edges of the wells for all conditions.

Conversely, no fluorescence was detected for α-SMA and SM-MHC in any condition for AT-SVF cells.

In general, higher fluorescence intensities were observed for calponin in cells exposed to PD98059,

however, the same correlation was not valid for desmin.

As already mentioned, smooth muscle cells are of great importance in tissue engineering

applications in general, as these cells line the vessels of the vascular system and are major

components of hollow visceral organs. Neovascularization is essential to supply oxygen and nutrients

to the cells in 3-D constructs, making this one of the biggest challenges that the tissue engineering

field needs to overcome.40 This challenge is hindered by the fact that smooth muscle cells do not

possess enough proliferative capacity to allow for significant cell numbers to be obtained for

Regenerative Medicine purposes.135 Moreover, SMC extraction from patients involves invasive tissue

biopsies. Consequently, other cell types with increased availability and expansion capabilities in vitro,

such as MSCs, have been strategically chosen as cell sources for differentiation assays directed

towards the smooth muscle lineage for tissue engineering applications.141,195 MSCs are the ideal cell

source given the phenotypic similarities between these two cell types.139 In fact, two decades ago,

cultured BM-MSCs were shown to express SMC markers such as the ones used herein (α-SMA,

calponin and SM-MHC) in long-term (7 week) confluent cell cultures.136 Since then, several studies

have successfully implemented different strategies to induce MSCs to differentiate into SMCs.

67

AT-SVF cells Day 7

BM-MSCs Day 7

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 Alexa4

88 for calponin and goat-anti rabbit Alexa 546 for desmin. Scale bar = 50 μm.

BM-MSCs AT-SVF cells

Day 2 Day 2

1 μM 10 μM 30 μM

Control 3%FBS

Control 10%FBS

1 μM 10 μM 10 μM (HG) 30 μM Control HG

3%FBS

α-SMA 8.2 5.8 29 7.5 25 0 0 0.20 0.10 0.10

Calponin 18 29 34 21 28 0 0 0 0 0

SM-MHC 0 1.9 1.3 0.5 1.0 0 0.20 0 0 0

Desmin - - - - - 4.5 3.8 3.0 3.5 4.5

Day 4 Day 4

1 μM 10 μM 30 μM

Control 3%FBS

Control 10%FBS


3%FBS

α-SMA 4.2 1.9 1.2 7.2 11 0.70 0.80 0 0.10 0.20

Calponin 20 14 14 13 15 0.30 0.50 0 0 0.30

SM-MHC 0 0 0 0.20 0 0 0 0 0 0

Desmin - - - - - 6.3 11 5.6 3.5 8.5

Day 7 Day 7

1 μM 10 μM 30 μM

Control 3%FBS

Control 10%FBS


3%FBS

α-SMA 0.10 0.50 0.50 1.0 0.20 0.10 0.50 0.50 1.0 0.20

Calponin 0 0.30 1.0 0.20 0.30 0 0.30 1.0 0.20 0.30

SM-MHC 0 0 0 0.10 0 0 0 0 0.10 0

Desmin - - - - - 3.6 4.2 4.2 22 8.5

1 μM 10 μM 10 μM (HG) 30 μM Control (HG)

1 μM 10 μM 30 μM Control (3%FBS) Control (10%FBS)

A

B

C

68

The SMC line used herein as positive control showed surprisingly low levels of SM-MHC

expression and less than 50% expression for α-SMA (assessed by flow cytometry). Still, in the

company’s website it is claimed that each lot is tested by flow cytometry for the smooth muscle

markers calponin and α-SMA.196 Hence, it is possible that the lower expression levels obtained here

are due to the primary antibodies used. Importantly, since SMCs show phenotypic plasticity, it is also

possible that the cells were not cultured for sufficient time periods to allow for a complete phenotype

switch into the contractile-phenotype (characterized by higher expression of contractile proteins such

as α-SMA, calponin and SM-MHC).197 Therefore, it can be hypothesized that most cultured SMCs still

retained a synthetic-phenotype (e.g. proliferative phenotype) at the time of marker expression

assessment, which ultimately resulted in the exhibition of low expression levels for contractile proteins.

Serum alone (3% and 10% FBS) seemed to induce the expression of SMC markers in

BM-MSCs and AT-SVF cells, possibly because of proteins and factors existing in the serum that might

stimulate smooth muscle-lineage commitment. α-SMA is described as one of the earliest SMC

markers expressed in undifferentiated BM-MSC culture, under normal and reduced serum conditions,

as well as in AT-SVF cells.136,139,198 Besides α-SMA, all isoforms of calponin and desmin were shown

to be expressed in undifferentiated BM-MSCs.199 Herein, this phenomenon was more evident for

calponin and desmin, for which positive cells were detected by immunofluorescence in all tested

conditions, including the control cells. Therefore, the only marker that can truly be considered as

“smooth muscle-specific” is SM-MHC, which is not spontaneously expressed by MSCs. Interestingly,

calponin-expressing cells tended to have broad polygonal shapes, resembling SMCs in confluent

culture.

In the conditions of the present study, PD98059 did not provide enough stimuli to significantly

and consistently enhance the expression of smooth muscle markers. No expression was detected by

flow cytometry for AT-SVF cells, for all markers except desmin. Since calponin-expressing cells were

observed at all timepoints in the immunofluorescence assay, it is possible that the secondary antibody

used in the flow cytometry analysis influenced the results. Again, fluorophore-conjugated primary

antibodies should be used in order to avoid possible fluorophore-derived variability in the results.

Several PD98059 pulses might be necessary to fully induce smooth muscle differentiation

(e.g. medium replacement every day for 5 days). Tamama and co-workers achieved the upregulation

of smooth muscle markers SM-MHC, caldesmon and α-SMA in BM-MSCs with PD98059. However,

the authors of this study did not mention how many pulses/medium changes were performed over the

induction time, which lasted 5 to 7 days according to the authors.140 No other studies regarding the

effect of PD98059 in MSC differentiation were found in the literature, except for one report in which it

was concluded that this molecule blocks osteogenic differentiation and promotes adipogenic

differentiation in a dose-dependent manner in MSCs.200 Conversely, several studies have

demonstrated that TGF-β is successful in inducing MSCs to the SMC lineage, and a correlation

between TGF-β production and embryonic SMC differentiation has been

established.46,138,139,142,143,195,201 Therefore, TGF-β might be a promising candidate to test in future

69

studies since several data are already available in the literature, which potentially facilitates the

establishment of a reproducible and efficient differentiation protocol. Other strategies may include the

use of commercially available smooth muscle differentiation medium, which was reported to induce

the expression of calponin, caldesmon, SM-MHC, SM22 and smoothelin.198

The results obtained herein for AT-SVF cells indicate that DMEM-HG did not enhance smooth

muscle differentiation. Interestingly, in a study where several medium formulations were tested for

AT-SVF expansion, it was concluded that cells isolated and expanded in DMEM-HG supplemented

with standard FBS more closely resembled SMCs than MSCs.159 Conversely, cells isolated in smooth

muscle cell medium (basal medium) presented a significantly higher frequency of adipogenic

progenitors.159 Taken together, these data reinforce the importance of medium formulation selection

when isolating and expanding AT-SVF, especially if the enrichment of non-stromal cell populations is

aimed. Thus, in future studies, AT-SVF isolation with DMEM-HG medium can be tested in order to

promote SMC enrichment.

Importantly, marker expression by immunofluorescence was only observed in cells near the

edges of the wells, which indicates that physical cues namely 3-D orientation should be vital for

directing the differentiation of MSCs into myogenic lineages. In a study where the effects of

biomechanical straining and surface patterns of substrates were tested on MSCs (without using any

biochemical reagents for cell proliferation and differentiation), it was concluded that higher level

straining and a grooved surface (rather than a flat surface) positively influenced the differentiation of

MSCs into smooth muscle-like cells.151 These results are in accordance with the abovementioned

observation. In one other study, the combination of mechanical (uniaxial strain) and biochemical

(TGF-β addition) stimuli was tested, and the results indicated that higher expression of a smooth

muscle phenotype at a transcriptional level was obtained in the presence of both types of stimuli.202

Lastly, in one study where three mechanical forces relevant to the cardiovascular system (cyclic

stretch, cyclic pressure, and laminar shear stress) were applied independently to BM-MSCs, cells

exposed to cyclic stretch expressed SMC proteins with increasing levels of differentiation with

increasing magnitudes of stretch.203 These studies show that surface patterning and mechanical

stimulation also play an important part in inducing myogenic differentiation and that biochemical cues

namely medium composition should not be the only factor taken into consideration when developing a

differentiation protocol. Additionally, the effect of ECM molecules such as gelatin, fibronectin or

collagen coatings should be also tested, as the presence of these ECM molecules was described to

further contribute to the stimulation of SMC differentiation in BM-MSCs.141 Finally, as the primary

function of SMCs is to contract and relax, cell contraction assays (e.g. measure of carbachol response

in collagen gels) should be performed after assessing smooth muscle marker expression (in case of

positive results).

70

III.4 Decellularization of Porcine Urethras

The development of strategies for urethral sphincter engineering requires the selection of

appropriate scaffolds for cell seeding, to be used once proper in vitro myogenic differentiation

protocols are established. In Section I.1.2.3 several potential scaffold materials were mentioned;

however, decellularized tissue matrices present advantages such as biological inherent bioactivity and

mechanical similarity to native ECM. Thus, protocols targeting efficient urethral tissue decellularization

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

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.

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