Differentiation plasticity of stromal vascular fraction cells ...

$: Differentiation plasticity of stromal vascular fraction cells ...$
Differentiation plasticity of stromal vascular fraction cells towards adipose tissue

and endothelial structures

Von der Medizinischen Fakultät

der Rheinisch-Westfälischen Technischen Hochschule Aachen

zur Erlangung des akademischen Grades

einer Doktorin der Medizin

genehmigte Dissertation

vorgelegt von

Melanie Esther Vicky Wosnitza

aus

Düsseldorf

Berichter: Herr Universitätsprofessor

Dr.med. Dr.univ.med. Prof. h.c.mult. Norbert Pallua

Herr Universitätsprofessor

Dr.med. Hans Merk

Tag der mündlichen Prüfung: 21. April 2009

Diese Dissertation ist auf den Internetseiten der Hochschulbibliothek online verfüg-

bar.

2

Differentiation plasticity of stromal vascular fraction cells towards

adipose tissue and endothelial structures

Der Medizinischen Fakultät der Rheinisch-Westfälischen Technischen Hochschule

Aachen vorgelegte Dissertation zur Erlangung des akademischen Grades einer Dok-

torin der Medizin

von

Melanie Esther Vicky Wosnitza

aus Düsseldorf

3

CONTENTS

ABBREVIATIONS ............................................................................................. 6

GERMAN SUMMARY ....................................................................................... 8

SUMMARY ...................................................................................................... 11

INTRODUCTION ............................................................................................. 12

Adipose Tissue ....................................................................................................................... 12 Source and location ............................................................................................................ 12

Cellular components ........................................................................................................... 12

Harvesting of adipose tissue derived cells ......................................................................... 13

Miscellaneous populations due to isolation and culture methods ....................................... 14

Adipose derived adult stem cells ............................................................................................ 16 Adipose tissue as source of mesenchymal stem cells ....................................................... 16

Adipose derived adult stem cells with a phenotype of bone marrow stem cells ................. 18

Relevant CD markers and their expression pattern ............................................................ 23

Differentiation capacity towards multiple lineages .............................................................. 23

Mesodermal differentiation ................................................................................................. 27 Adipogenesis ................................................................................................................................ 27 Chondrogenesis ............................................................................................................................ 29 Osteogenesis ................................................................................................................................ 30 Skeletal myogenesis ..................................................................................................................... 31 Cardiomyogenesis ........................................................................................................................ 31 Immune cells ................................................................................................................................. 32

Ectodermal differentiation ................................................................................................... 33 Neurogenesis ................................................................................................................................ 33 Epithelial differentiation ................................................................................................................. 33

Endodermal differentiation ................................................................................................. 34 Hepatogenic and intestinal differentiation ..................................................................................... 34

Vasculogenesis and angiogenesis ......................................................................................... 34 Angiogenesis in adipose tissue .......................................................................................... 35

4

Adipogenesis and angiogenesis ......................................................................................... 35

STUDIES ......................................................................................................... 41

MATERIAL AND METHODS .......................................................................... 44

Technical equipment .............................................................................................................. 44

Reagents ................................................................................................................................ 45

Culture media ......................................................................................................................... 47

Buffers and solutions .............................................................................................................. 49

Isolation of SVF cells .............................................................................................................. 50 CD31 Dynabead separation of SVF cells ........................................................................... 52

CD31 MicroBead separation of SVF cells .......................................................................... 54

Culturing of SVF cells ......................................................................................................... 56

Cell splitting ........................................................................................................................ 57

Fibronectin and MatrigelTM coating ..................................................................................... 58

Human dermal microvascular endothelial cell (HDMEC) isolation ......................................... 58

Human dermal fibroblast isolation .......................................................................................... 59

Human umbilical vein endothelial cell isolation ...................................................................... 59

Flow cytometry analysis ......................................................................................................... 60

Immunostaining with von-Willebrand-factor ........................................................................... 61

Determination of glycerol-3-phosphate dehydrogenase (GPDH) activity ............................... 61

Oil Red O staining .................................................................................................................. 63

Cell counting ........................................................................................................................... 63

Statistical evaluation ............................................................................................................... 63

RESULTS ........................................................................................................ 64

Comparison of excised fat tissue versus liposuction material ................................................ 64

Characterisation of surface markers and morphological changes in SVF cells in culture ...... 67

In vitro differentiation of unpurified SVF cells under endothelial medium conditions ............. 68

Splitting and purity analysis of CD31- and CD31+ SVF cells .................................................. 70

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Differentiation potential of CD31- and CD31+ SVF cells towards EC lineage ......................... 72

In vitro differentiation of SVF cells under adipogenic medium conditions .............................. 76

DISCUSSION .................................................................................................. 79

Tissue engineering with adipose derived adult stem cells ..................................................... 80 Present problems and hurdles............................................................................................ 80

How to manage insufficient vascularisation ........................................................................ 81

Characterization of the SVF ................................................................................................... 81

ADAS cell differentiation towards endothelial cells ................................................................ 84 Therapeutic angiogenesis .................................................................................................. 85

Endothelial cell differentiation to adipocytes ....................................................................... 88

Arteriosclerosis and adipose tissue ........................................................................................ 90 The impact of progenitor cells in atherosclerosis ............................................................... 94

CONCLUSION ................................................................................................. 95

REFERENCES ................................................................................................ 96

PUBLICATIONS ............................................................................................ 110

Full papers ............................................................................................................................ 110

Published abstracts .............................................................................................................. 110

Scientific presentations ........................................................................................................ 110

ACKNOWLEDGEMENT................................................................................ 112

ERKLÄRUNG § 5 ABS. 1 ZUR DATENAUFBEWARUNG .......................... 113

CURRICULUM VITAE ................................................................................... 114

6

ABBREVIATIONS

1,25 (OH)2D3 1alpha,25-dihydroxyvitamin D3 IgG Immunoglobulin

acLDL Acetylated low-density lipoprotein IL-3 Interleukin-3

ACLP Aortic carboxypeptidase-like protein IL-6 Interleukin-6

ADAS cell Adipose-derived adult stem cell IMDM Isocove’s Modified Dulbecco’s Medium

Ang Angiopoietin KCl Kalium chloride

aP2 Adipocyte fatty acid binding protein KDR Kinase insert domain receptor

APC Allophycocyanin KHCO3 Potassiumhydrogencarbonate Kit

AT Adipose tissue LDL Low-density lipoprotein

ATP Adenosintriphosphate LPL Lipoprotein lipase

ATRA All-trans retinoic acid Mac-1 Macrophage antigen-1/CD11b

BAT Brown adipose tissue MAP2ab Microtubule associated protein 2 a/ b

bFGF Basic fibroblast growth factor MCP-1 Monocyte chemotactic protein-1

BMP-2 Bone morphogenetic protein-2 M-CSF Macrophage colony-stimulating factor

BMSC Bone marrow stem cell MMP Matrix metalloproteinases

BSA Bovine serum albumin MOMA-2 Anti-macrophage monoclonal antibody-2

CaCl2 Calcium chloride MS Metabolic syndrome

CD Cluster of differentiation MSC Mesenchymal stem cell

C/EBP CCAAT/enhancer binding protein Myf Myogenic factor

C/EBP-α CCAAT/enhancer binding protein alfa MyoD1 Myogenic differentiation 1

CNS Central nervous system NaCl Natrium chloride

CO2 Carbone dioxide NAD Atrial natriuretic peptide

CYP Cytochrome P450 NaHCO3 Sodium bicarbonate

DAPI 4’,6’-Diamidin-2’-phenylindol-dihydrochlorid Neu N Neuronal nuclear protein

DiI 1,1'-dioctadecyl-3,3,3',3’-tetramethylindocarbocyanine

perchlorate

NH4Cl Ammonium chloride

DMEM Dulbecco’s Modified Eagle Medium NMDA N-methyl-D-aspartate

DMSO Dimethylsulfoxid NF Neurofilament

EC Endothelial cell NO Nitric oxide

ECGM Endothelial Cell Growth Medium PA Plasminogen activator

ECM Extracellular matrix PAI-1 Plasminogen activator inhibitor 1

ED2 Anti perivascular monocytes/macrophages antibody PBS Phosphate buffered saline

EDTA Ethylene diamine tetraacetic acid PD Population doublings

EGF Epithelial growth factor PDGF Platelet-derived growth factor

EGM-2 (MV) Endothelial Growth Medium 2 (Microvascular) PE Phycoerythrin

eNOS Endothelial nitric oxide synthetase PECAM-1 Platelet endothelial cell adhesion molecule 1

EPC Endothelial progenitor cell PFA Paraformaldehyde

F12 Ham’s F12 PFF Pulsating fluid flow

FA Fatty acid PLGA Poly(lactic-co-glycolic acid)

FACS Fluorescence activated cell sorting PPARγ Pperoxisome proliferator-activated receptor gamma

FCS Fetal calf serum PRELP Proline/arginie-rich end leucine-rich repeat protein

FGF Fibroblast growth factor PTH Parathyroid hormone

FITC Fluorescein-Isothiocyanat RBM Reconstituted basement membrane

Flk-1 Fetal liver kinase 1 SCC Sideward scatter

Flt-1 Fms like tyrosine kinase 1 SCD1 Stearoyl-CoA desaturase

FSC Forward scatter SCF Stem cell factor

7

GABA Gamma-amino butyric acid SCGF-β Stem cell growth factor beta

GFAP Glial fibrillary acidic protein SD Standard deviation

Glut4 Glucose transporter 4 SM Smooth muscle

GPDH Glycerol-3-phosphate dehydrogenase SMC Smooth muscle cell

Gy Gray SSC Sideward scatter

HDL High density lipoprotein SVF Stromal vascular fraction

HDMEC Human dermal microvascular endothelial cell TE Tissue engineering

HEPES N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid TGF-β Transforming growth factor beta

hATSC Human adipose tissue stem cell TIMI-3 Thrombolysis in Myocardial Infarction 3

hMADS cell Human multipotent adipose-derived stem cell TIMP Tissue inhibitor matrix metalloproteinases

hTERT Human telomerase TNF-α Tumor necrosis factor alfa

HMG-CoA 3-hydroxy-3-methylglutaryl coenzyme A VCAM-1 Vascular cell adhesion molecule 1

HNF4α Hepatocyte nuclear factor 4 alfa VE-cadherin Vascular endothelial cadherin

hS Human autologous serum VEGF Vascular endothelial growth factor

HSC Hematopoietic stem cell VEGF-R1 Vascular endothelial growth factor receptor 1

HUVEC Human umbilical vein endothelial cell VEGF-R2 Vascular endothelial growth factor receptor 2

IBMX Isobutylmethylxantine vWF Von-Willebrand factor

ICAM-1 Intercellular adhesion molecule 1 WAT White adipose tissue

IGF-1 Insulin-like growth factor 1

8

GERMAN SUMMARY In den letzten Jahrzehnten hat sich gezeigt, dass Fettgewebe weitaus mehr darstellt als ein

simples Energiespeicherorgan. Es besteht neben reifen Fettzellen, sog. Adipozyten, aus

Fettvorläuferzellen, auch Präadipozyten genannt. Präadipozyten gehören zu den mesen-

chymalen Stammzellen des Körpers und konnten das erste Mal vor mehr als dreißig Jahren

aus humanem menschlichem Fettgewebe isoliert werden. Seither sind die Vorläuferzellen im

Fettgewebe mehr und mehr in das Interesse der Forschung gerückt. Es ließ sich zeigen,

dass Präadipozyten keine homogene Zellpopulation darstellen, wie anfangs geglaubt, son-

dern ganz im Gegenteil ein heterogener Mix aus verschiedenen Zelltypen sind. Die Hetero-

genität ist eine Konsequenz der Methode zur Gewinnung von Fettvorläuferzellen, nämlich

Fettgewebsexzision oder Liposuktion. Sowohl das Herausschneiden als auch das Absaugen

von Fett beinhaltet die Gewinnung eines ganzen Gemisches von stromalen Zellen, nicht nur

selektiv von Fettvorläuferzellen. Die isolierten Zellen sind neben den klassischen Präadipo-

zyten auch Endothelzellen, endotheliale glatte Muskelzellen, Fibroblasten, Makrophagen und

Blutzellen; daher spricht man auch allumfassend von der stromalen vaskulären Fraktion

(SVF). Es hat sich gezeigt, dass sich SVF Zellen nicht nur zu reifen Adipozyten differenzie-

ren lassen, sondern auch in viele andere mesenchymale sowie nicht-mesenchymale Zellty-

pen. Die SVF des Fettgewebes gehört zu den Zellpopulationen, die sehr schwer zu charakte-

risieren sind. Daher ist die Terminologie auch nicht einheitlich. Sehr häufig finden sich neben

dem Begriff adipose SVF Bezeichnungen wie human adipose tissue-derived stem cells oder

adipose lineage cells. Ihre außerordentliche Differenzierungskapazität und ihre Oberflä-

chenmerkmale unterstreichen die Ähnlichkeit von SVF Zellen zu Knochenmarkstammzellen.

Aufgrund ihrer breiten Differenzierungskapazität in meso-, ekto- und endodermale Zelltypen

können die fettgeweblichen SVF Zellen als multipotente Stammzellen betrachtet werden.

Neueste Forschungsergebnisse legen nahe, dass Fettgewebszellen und Endothelzellen ei-

nen gemeinsamen Zellvorläufer haben könnten. Dies wird durch die enge Beziehung zwi-

schen Angiogenese und der Entstehung von Fettgewebe unterstrichen. Angiogenese und

Adipogenese beeinflussen sich gegenseitig; so verbessert die Interaktion zwischen Endo-

thelzellen und Adipozyten das Wachstum beider Zelltypen und bewirkt gleichzeitig eine Re-

duktion der Apoptoserate. In Hinblick auf eine gemeinsame Vorläuferzelle bleibt die For-

schung allerdings noch entscheidende Erklärungen schuldig, insbesondere der Verbin-

dungsweg zwischen Fettgewebe und Endothel, sowie das Differenzierungspotential der Zel-

len in die eine oder andere Richtung über eine gemeinsame Vorläuferzelle konnten noch

nicht demonstriert werden. Diese Themen waren daher ein zentraler Fokus dieser Arbeit.

9

SVF Zellen wurden aus dem Fettgewebe isoliert und ihre Oberflächeneigenschaften, das CD

(cluster of differentiation) Profil, untersucht. Für die Isolation von SVF Zellen kommen zwei

Methoden in Frage, zum einen die Fettgewebsexzision mit anschließender Präparation der

gewonnenen Fettgewebsläppchen oder die Liposuktion, bei der die anschließende Gewebs-

präparation entfällt. Um beide Methoden hinsichtlich ihrer Unterschiede in der Zellgewinnung

zu vergleichen, wurden die Oberflächeneigenschaften von SVF Zellen aus exzidiertem Ge-

webe und aus liposuktioniertem Gewebe mittels Durchflusszytometrie bestimmt. Hierbei

wurde ein besonderes Augenmerk auf endotheliale Marker sowie Stammzellmarker gelegt,

um eventuelle Kontaminationen mit anderen Zelltypen aufzudecken. Des Weiteren wurde die

Expression der CD Marker während einer Zellkulturphase untersucht, um Veränderungen

der Zelloberflächeneigenschaften im Kultivierungsverlauf darzustellen. Dies ist insbesondere

von Bedeutung, wenn SVF Zellen zur Zellvermehrung über mehrere Passagen kultiviert wer-

den und erst dann für Differenzierungsexperimente verwendet werden sollen. Das zelluläre

CD Profil wurde neben der Durchflusszytometrie mit Hilfe von Fluoreszenzfärbungen be-

stimmt. Als endotheliale Marker wurden hauptsächlich CD31, vWF und CD105 verwendet,

als Stammzellmarker CD34, KDR und S100. Weiterhin wurde die fettgewebliche SVF mit

Hilfe von magnetischen Beads (Dynabeads und MicroBeads) in eine CD31+ und eine CD31-

Fraktion aufgeteilt und die Differenzierungskapazität der jeweiligen CD31-Gruppe in Rich-

tung Fettgewebe sowie Endothel parallel untersucht. Die Differenzierung zu Endothel wurde

mit Hilfe verschiedener Endothelzellmedien und der Kultur auf Extrazellularmatrix (Matri-

gelTM) durchgeführt. Hier wurde das Augenmerk insbesondere auf die Ausbildung kapillärer

Strukturen auf MatrigelTM gelegt, zudem wurden die zellulären Oberflächeneigenschaften mit

Hilfe der Durchflusszytometrie bestimmt. Die Fettzelldifferenzierung der CD31- und CD31+

SVF Zellen erfolgte mittels Kultivierung in einem Fettzelldifferenzierungsmedium über insge-

samt 21 Tage. Die Differenzierung wurde mit dem Glycerol-3-Phosphat Dehydrogenase

(GPDH) Enzymassay sowie durch Öl-Rot Färbung und Zellzählung validiert.

Frisch isolierte SVF Zellen aus exzidiertem Fettgewebe zeigen signifikante Unterschiede in

ihrer Oberflächenbeschaffenheit im Vergleich zu Zellen aus Liposuktionsgewebe. SVF Zellen

aus exzidiertem Fettgewebe sind in 5-10% positiv für CD31, die Zellen aus Liposuktionsge-

webe zeigen hingegen einen deutlich höheren Anteil an CD31 positiven Zellen von bis zu

40%. Ähnliches gilt für KDR. Beide Zellpopulationen haben einen vergleichbaren, deutlich

positiven Nachweis von CD34. Im Verlauf kam es bei beiden Fraktionen nach 14 Tagen in

der Zellkultur zu einem Abfall der Stammzellmarker (CD34, KDR, S100). Um das endothelia-

le Differenzierungspotential von CD31- SVF Zellen zu evaluieren, wurden die Zellen auf Mat-

rigelTM Extrazellularmatrix in Endothelzelldifferenzierungsmedium kultiviert. Hier zeigten die

CD31- Zellen eine ausgeprägte Tendenz, dreidimensionale, tubuläre Strukturen im Sinne

10

eines kapillären Netzwerkes auszubilden. Die Ergebnisse wurden durch die Durchflusszyto-

metrie bestätigt. Frisch isolierte SVF Zellen ohne weitere Selektierung differenzieren in vitro

zu ca. 90% zu reifen Adipozyten. Hier wurden CD31- und CD31+ SVF Zellen im Fettzelldiffe-

renzierungsmedium kultiviert und die Ausbeute der reifen Fettzellen bestimmt. Die CD31-

Zellfraktion differenziert zu fast 100% zu Adipozyten. Interessanterweise sind auch die

CD31+ SVF Zellen in der Lage eine Fettzelldifferenzierung zu vollziehen, hier zeigten ca.

65% der Zellen Lipidvakuolen. Die Bestimmung der GPDH Aktivität ergab ebenfalls einen

Wert von 64% ± 11%, verglichen mit der GPDH Aktivität der CD31- Zellen. Während der Dif-

ferenzierung zu reifen Adipozyten trugen die CD31+ SVF Zellen weiterhin die CD31-

Dynabeads auf der Zelloberfläche.

Die frisch isolierten SVF Zellen des Fettgewebes zeigen unabhängig von der Isolationsme-

thode eine starke Expression von Stammzellmarkern. Die SVF enthält jedoch auch eine Po-

pulation von CD31+ Zellen, insbesondere in Liposuktionsgewebe. Dies weist auf eine hohe

Kontaminationsrate mit Endothelzellen in liposuktioniertem Fettgewebe im Vergleich zu exzi-

diertem Gewebe hin. Insgesamt fallen die Stammzellmarker während der Kultivierung deut-

lich ab, was wiederum in Übereinstimmung mit der Erfahrung ist, dass Präadipozyten ihre

Differenzierungskapazität nach mehreren Passagen verlieren.

Die Ergebnisse der endothelialen und adipogenen Differenzierung belegen, dass die fettge-

webliche SVF einen CD31- Zelltypen enthält, der zu Adipozyten sowie Endothelzellen diffe-

renziert werden kann. Zudem gibt es CD31+ Zellen, die, obwohl sie einen endothelialen Phä-

notyp besitzen, in reife Fettzellen differenziert werden können. Diese Resultate demonstrie-

ren die Fähigkeit der SVF Zellen sich in beide Richtungen, und zwar Adipozyt sowie Endo-

thelzelle, differenzieren zu können. Allgemein gesprochen zeigen die Ergebnisse die Plastizi-

tät ausgereifter, mesenchymaler Zellen aus dem Fettgewebe, eine Differenzierung zwischen

Endothel und Fettgewebe sowie auch umgekehrt vollziehen zu können.

Des Weiteren könnten diese Resultate von Relevanz für die Pathogenese der Arteriosklero-

se sein. Hier scheinen Transdifferenzierungsprozesse endothelialer Vorläuferzellen eine Rol-

le zu spielen.

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SUMMARY Recent research findings postulate that adipocytes and endothelial cells (ECs) may share a

common progenitor. However, the interlinking pathways between adipose tissue and endo-

thelium, and the differentiation potential of cells to convert from one tissue into the other via

progenitor cells, have not been elucidated so far and are therefore the focus of this thesis.

Stromal vascular fraction (SVF) cells were isolated from liposuction aspirates or excised adi-

pose tissue and separated into CD31+ and CD31- populations by magnet-assisted cell sort-

ing. Differentiation to fat tissue was induced in both CD31 fractions. Differentiation was as-

sayed enzymatically and by cell counting. Maturation to endothelium was performed with

vascular endothelial growth factor, insulin like growth factor-1 plus 2% FCS and confirmed by

flow cytometry and tube formation assays on Matrigel™. The results presented here show

that the SVF contains a CD31-, S100+ cell type which can differentiate into adipocytes and

EC. The SVF also comprises CD31+ cells that, although they have an endothelial phenotype,

can be converted into mature adipocytes. These findings demonstrate the potency of SVF

cells to perform both adipogenic and endothelial differentiation. Further, they reveal the plas-

ticity of mature cells of mesenchymal origin to undergo conversion from endothelium to adi-

pose tissue and vice versa.

INTRODUCTION

ADIPOSE TISSUE

SOURCE AND LOCATION

In the last decades the belief was corroborated that adipose tissue is much more than a sim-

ple fat storage. Adipose tissue plays a central role in energy homeostasis and was found to

be a highly active metabolic organ335. Many secretory factors produced by fat cells influence

the endocrine regulation of the body, affecting growth, metabolism and behaviour. Fat tissue

is a highly specialized connective tissue located in different places of the body and displaying

various functions. In humans, white adipose tissue composes as much as 20% of the body

weight in men and 25% of the body weight in women. A major portion is the subcutaneous

fat tissue located directly underneath the dermis. Furthermore, adipose tissue is found in

intraabdominal locations, as omental fat pads and surrounding several organs of the human

body. Dependent on the location there are two different types of fat: white and brown adipose

tissue. Both cushion and insulate the body, but each of them has further specialized func-

tions as well. White adipose tissue (WAT) is mainly found in the subcutaneous regions, but

also in the Omentum Majus, lying intraabdominally between the abdominal muscles and the

visceral organs. WAT basically functions as energy source for the body, whereas brown adi-

pose tissue (BAT) displays the heat source, primarily at young age. BAT is surrounding

nearly all organs in the abdomen as well as in the chest. During the natural aging process

brown fat tissue is gradually replaced by white adipose tissue.

CELLULAR COMPONENTS

The primary cellular component of adipose tissue is a collection of so called adipocytes, lipid

filled fat cells. Focussing on the two different types of fat tissue, morphological and ultrastruc-

tural properties of brown and white adipocytes reflect the different functions in thermoregula-

tion and energy balance. White adipocytes contain only a single large fat vacuole, whereas

brown fat cells consist of several smaller vacuoles and a much higher number of mitochon-

dria76. Brown and white adipose tissue are both innervated by the sympathetic nervous sys-

tem, which controls metabolic and lipolytic activity as well as the vascularisation in adipo-

cytes18,19.

White mature adipocytes consist of about 90% lipids303 and each of them stands in close

contact with at least one capillary, providing a vascular network that allows continued growth

of the organ304. A further component located in the adipose organ is the stromal vascular

fraction (SVF), stromal cells which are mainly located around the blood vessels. This adipose

13

SVF contains a large population of preadipocytes, first identified by Poznanski et al. in 1971

following the enzymatic digestion of adipose tissue282,317. Preadipocytes were recently identi-

fied to have major properties of adult mesenchymal stem cells. These precursor cells are

present troughout adult life, and differentiation into new mature adipocytes can still be

achieved in old animals211,269.

HARVESTING OF ADIPOSE TISSUE DERIVED CELLS

A number of factors have to be taken into consideration when isolating preadipocytes or SVF

cells from adipose tissue. Overall cell yield is determined by donor age, obesity, the anatomi-

cal site of harvest, the methods of tissue harvest, storage and preadipocyte isolation and the

in vitro culture environment (for review158). Besides the isolation method of preadipocyte that

defines the cell numbers harvested, the donor side exhibits a major factor for the adipogenic

capacity of preadipocytes. In general human abdominal subcutaneous preadipocytes seem

to have a greater potential for adipogenesis as omental or intraabdominal preadipo-

cytes150,265,384, however, other authors state that there is no difference in adipogenic potential

between both locations352,399. In subcutaneous locations abdominal preadipocytes have a

greater potential to convert into mature adipocytes than femoral preadipocytes146. Addition-

ally proliferation capacity in subcutaneous preadipocytes is superior to those of omental cells

since subcutaneous cells proliferate faster under in vitro conditions399. Additionally the donor

age plays an important role in proliferation rates of harvested cells where a negative correla-

tion can be observed between age and doubling capacity33,399.

To release the broad spectrum of different cell types harboured in adipose tissue, minced fat

lobules from excised tissue or plain lipoaspirate are used. The material is washed after har-

vesting to remove possible blood contamination. The extracellular matrix (ECM) holding the

adipocytes in place is digested with collagenase solution, followed by centrifugation to divide

mature fat cells from stromal cells. The cells located in the stromal fraction mainly appear in

a fibroblast-like morphology after adherence to cell culture dishes, macroscopically indistin-

guishable from factual fibroblasts. At the beginning of fat tissue research these cells were

referred to as preadipocytes or adipoblasts5,52,130, as they were found to give rise to fat vacu-

ole-containing cells. Both terms refer to fat cell precursors, but adipoblasts were regarded to

be the juvenile adipose stem cell, contrariwise preadipocytes as the cell type in between adi-

poblast and adipocyte5. However this could not be asserted. Instead, the term preadipocyte

was established over the last three decades and is still frequently used in the literature for

precursor cells derived from fat tissue65,79,92,118,140,158,160,200,350,407.

Preadipocytes are mainly located in the surrounding of small blood vessels and between

mature fat cells in the stromal vascular fraction of fat tissue260,288,290 and can be recruited a

lifelong6. Nevertheless, harvested adipose stromal cells display a heterogeneous cell popula-

14

tion directly after isolation and are not as homogenous as initially believed. Regarding the

isolation method of adipose stromal cells it is likely that not solely preadipocytes are digested

from the ECM but also cells located in the blood vessels and the vessel walls. As a result the

adipose SVF not exclusively comprises preadipocytes but also microvascular endothelial

cells, endothelial smooth muscle cells, fibroblasts, macrophages, and blood cells (Figure

1)427,434.

SVFSVF

preadipocytes

endothelialcells

smooth musclecells

fibroblasts

macrophages

bloodcells

Major progress was made in the last few years concerning the nomenclature of the adipose

stromal cells due to newly discovered cell features and cell characteristics158,314,433. At the

beginning of fat tissue research the differentiation process from preadipocyte to adipocyte

was believed to be an irreversible process, with mature fat cells as an end-stage product. In

vitro studies, however, have demonstrated that the phenomenon of differentiation seems to

be reversible. It has been proven in cell culture experiments that dedifferentiation of adipo-

cytes back into preadipocytes is possible281,334,430. In addition, “preadipocytes” have been

found to be able to differentiate not only into fat cells but also into many other mesenchymal

and non-mesenchymal cell lineages55,382,433. Further research revealed that adipose stromal

cells have surface characteristics comparable to bone-marrow stem cells (compare Table

1)133. Taking their remarkable differentiation potential433, these cells can be regarded as adult

mesenchymal stem cells capable of multipotency372,388,433.

MISCELLANEOUS POPULATIONS DUE TO ISOLATION AND CULTURE METHODS

Although intensive research has been performed on fat tissue progenitor cells, the adipose

SVF remains a cell population very hard to characterize. Consequently, in recent literature

Figure 1. Adipose stromal

vascular fraction (SVF)

Freshly isolated SVF cells

from fat tissue display a het-

erogeneous cell population

comprising contrasting cell

types, such as preadipocytes,

endothelial cells, smooth mus-

cle cells, fibroblasts, blood cells

and macrophages.

15

many different terms refer to this cell mixture isolated from adipose tissue by collagenase

digestion. Most often these cells are called preadipocytes, but adipose tissue-derived stem

cells113,260, adipose-derived adult stem cells23, adipose tissue mesenchymal stem cells244,382,

multi-lineage cells from adipose tissue98, adipose tissue-derived stromal cells109,133, or adi-

pose lineage cells314 have been used to describe this population as well. Since different do-

nor sites such as abdominal, gluteal, mammary or intraabdominal fat tissue are applied for

SVF harvesting, nonuniform populations of preadipocytes are often used for cell surface

characterisation and differentiation assays. Consequently, literature results vary widely re-

garding the surface protein expression and the differentiation plasticity of adipose stromal

cells.

Two miscellaneous procedures are described for primary isolation of SVF cells, liposuction

and plain excision during procedures, such as abdominoplasty or breast reduction. It remains

controversial whether liposuction aspirate or excised adipose tissue displays the superior

source for preadipocyte harvesting. Some studies state that liposuction aspirates represent a

better source compared to excised fat405, others claim the contrary176. After excision of adi-

pose tissue (AT), fat lobules need to be isolated by removing capillaries and connective tis-

sue, a possible contamination source for endothelial cells and fibroblasts. Liposuction is per-

formed by manually applying negative pressure using a 10-cc-syringe with a blunt tip

cannula80. Due to the harvesting procedure the isolation by liposuction damages not only AT

but also the surrounding structures including capillaries and fibrous tissue. Therefore minced

fat tissue from lipoaspirates contains vessels and connective tissue besides the fat compart-

ment.

The adipose SVF harbours not only preadipocytes, but also other mature cell types, e.g. en-

dothelial cells (CD31+/CD34+), macrophages (CD14+/CD31+), and blood cells (see Figure 1).

Certain cell fractions can be isolated with magnet-assisted cell sorting due to the expression

of surface proteins including stem cell markers like CD34260 or kinase insert domain receptor

(KDR)60,252, macrophage markers as CD14350, and fibroblast markers. This cell sorting can

assist in creating a more pure and homogenous population of SVF cells by using a negative

selection method to gain the non-labelled cells, the classical preadipocytes.

Rodriguez et al.331 established a method to isolate two distinguishable cell populations from

the adipose SVF based on adhesion characteristics. Cells either displayed a fast- or a slow-

adherence one. Both populations exhibited similar characteristics concerning doubling time

(36h) and differentiation potential towards adipocytes and osteoblasts for up to 60-80 pas-

sages. After this growth period slow-adherent SVF cells ceased to proliferate and also to

differentiate. In contrary fast-adherent SVF cells exhibited a slow proliferation rate over time

(72h) and expressed significant levels of telomerase activity, whereas slow-adherent SVF

cells did not. Fast-adherent SVF cells responded mainly to basic fibroblast growth factor

16

(bFGF) with expansion, but not to epithelial growth factor (EGF) or platelet-derived growth

factor (PDGF) and could be cultured for more than 200 passages. This fast-adherent cell

type from the adipose SVF was termed human multipotent adipose-derived stem cells

(hMADS cells)332.

ADIPOSE DERIVED ADULT STEM CELLS

ADIPOSE TISSUE AS SOURCE OF MESENCHYMAL STEM CELLS

Stem cells are defined as being qualified to distribute renewal cells for a particular mature

cell type29. Three miscellaneous aspects delineate this special cell fraction and underline

their uniqueness and difference from all other cell populations harboured in the human body.

To begin with, stem cells are capable to replicate themselves leading to self-renewal and an

extensive proliferation capacity. Second, stem cells display an immature and unspecialized

cell type, which indicates that they do not have any tissue specificity and therefore lack spe-

cialized, tissue-specific functions. Thirdly, stem cells are able to differentiate into at least one

specialized cell type. How many different cell phenotypes a stem cell can convert into, known

as stem cell plasticity or potency, can be outlined with different stem cell-defining terms.

Stem cells may be categorized as either totipotent, pluripotent, or multipotent, whereby the

stem cell is able to form all, most, or a small number of miscellaneous cells of the body. Fur-

ther, stem cells capable of producing the blood cells of an organism are defined as hemato-

poietic stem cells.

Several sources of stem cells were established in the past to harvest multipotent stem cells

from lineages of the three different germinal sheets (Figure 2). According to the embryonal

sheet and tissue location, stem cells can be classified as mesodermal, ectodermal or endo-

dermal412. Stem cells exist in humans throughout adult life in special stem cell niches418. Fur-

ther stem cells can be harvested as embryonic stem cells from the inner cell mass of the pre-

implantation blastocyst. Embryonic stem cells were isolated from either human, non-human

primates or mice from 3-5 days old embryos43,403. Still capable of pluripotency, these cells

have the capacity to undergo nearly the full range of differentiation pathways, including all

three germinal lines264,318,343,359,416. Differentiation phenotypes include adipocytes, endothelial

cells, chondrocytes, muscle cells, neuronal cells as well as blood cells and several others

(Figure 2)43,165,236,403. However, major concerns regarding ethical topics in application of em-

bryonic stem cells in research, restrict a wider range of promising clinical usage and trials. By

applying adult stem cells instead, most of these ethical problems can be avoided.

17

Epiblast-like pluripotent SC

Endodermal SCMesodermal SCEctodermal SC

LiverSC

PancreaticSC

IntestinalSC

MesenchymalSC

Hemangioblast

Vascular SCHematopoieticSC

SkinSC

NeuralSC

Epithelial cellMelanocyteHair follicle

NeuronsAstrocyte

Oligo-dendrocyte

LymphoidProgenitor

cell

MyeloidProgenitor

cell

EndothelialProgenitor

cell

EndothelialCell

Pericyte

PreadipocyteFibroblastOsteoblast

ChondroblastMyoblast

Cardiomyoblast

AdipocyteFibrocyte

Smooth muscle cellOsteocyte

ChondrocyteMyocyte

Cardiomyocyte

Hepatocyte

Islet cell

Intestinalcell

MonocyteMakrophageGranulocyteErythrocyte

PlateletDendritic

cell

T-lymphocyteB-lymphocyte

NK cell

Adult stem cells, often termed mesenchymal or somatic stem cells, are mature, undifferenti-

ated cells which retained their unique capacity to differentiate into a wide spectrum of differ-

ent tissue types. They display a reserve of fibroblast-like cells that lie secluded in mature

tissue yet can home to and repair damaged tissue when necessary219. Therefore these cells

display an interesting source for multiple clinical applications135,258,259. Since most mesen-

chymal stem cells (MSCs) can be easily isolated, cultured and expanded, application for a

cell-based therapy is highly attractive, e.g. the treatment of children with osteogenesis imper-

fecta168, patients with Alzheimer’s37,375,376, Parkinson’s191,394, and Crohn´s disease122,

Duchenne muscular dystrophy332, burns321,357, cerebral ischemia193, spinal cord injuries196,

hematopoietic recovery214, heart disease131,170,373, osteoarthritis105,327 and others.

Mesenchymal stem cells preserve their ablility to differentiate into mature specialized cell

phenotypes28,30,38,135,164,294,312,403 and perform self-renewal combined with maintaining their

Figure 2. Postnatal stem and progenitor cells

Differentiation cascade of stem and progenitor cells from mesoderm, ectoderm and endoderm.

18

specific stem cell plasticity and phenotype for several generations218. Adult stem cells com-

prise bone marrow stromal cells (BMSCs), hematopoietic stem cells (HSCs), endothelial pro-

genitor cells, as well as neural, dermal, hepatic and adipose stem cells, among others.

Located in distinct stem cell niches in divergent tissues and organs, adult stem cells can be

found all over the human body. One of the best known niches exists in bone marrow, har-

bouring HSCs as well as MSCs185. Additiononally multiple other sources for MSCs exist in

the human body, including trabecular bone285, skeletal muscle428, dermis428, synovium344,

deciduous teeth395, lung, liver, intestine, pancreas, heart35, central nervous system (CNS)

tissue, human umbilical cord blood106,209, human placenta178, perivascular cells346 as well as

adipose tissue209.

Adipose tissue comprises a stromal vascular fraction comparable to the stromal fraction of

bone marrow. SVF cells from adipose tissue have been identified as mesenchymal cells,

being directly derived from mesenchyme, just like bone marrow433. Stromal cells from fat tis-

sue include adult mesenchymal stem cells, often referred to as preadipocytes, and in addi-

tion various other cell types as mentioned earlier98,113,434. Under stem cell supporting culture

conditions, the SVF turns into a more homogeneous population, termed human adipose-

derived adult stem (ADAS) cells. ADAS cells have nearly equal morphology and characteris-

tics as BMSCs. Most notably they demonstrate extensive expansion potential, plasticity to

undergo multilineage differentiation and surface protein phenotype. ADAS cells are therefore

an alternative source for mesenchymal stem cells96,113,133,186,233,304,326,332. Additionally an ideal

source of stem cells should be abundant, accessible and readily isolated with minimal risk to

the patient. ADAS cells seem to outfit precisely this assortment of specifications.

ADIPOSE DERIVED ADULT STEM CELLS WITH A PHENOTYPE OF BONE MARROW

STEM CELLS

The best known origin for mesenchymal stem cells is the stroma of bone marrow which con-

tains adult stem cells, the BMSCs. ADAS cells have been identified as mesenchymal cells as

they are located in the stroma of adipose tissue, which is derived from mesenchyme much

like bone marrow. Located between mature adipocytes both stem cell populations (ADAS

cells and BMSCs) are harboured in a highly vascularised tissue with close contact to blood

circulation.

Not surprisingly ADAS cells exhibit a differentiation repertoire similar to that described for

bone marrow stem cells. The cells are multipotent adult stem cells and have the ability for

multilineage differentiation429,433,434 comprising the mesenchymal and non-mesenchymal

branches. When cultured in vitro, ADAS cells as well as BMSCs show a fibroblast-like mor-

phology after adhering to the culture flask. Both cell types can be easily cultured and ex-

panded in vitro. Further similarities between ADAS cells and BMSCs have been described in

19

the literature focussing on growth kinetics and expansion capacity, cell senescence, gene

transduction efficiency, gene transcription and protein surface expression.

Noted exceptions comprise the presence of CD49d as well as the low expression of CD133,

the lack of telomerase activity and absence of STRO-1 and CD62L on ADAS cells (Table 1).

CD49d, STRO-1 and CD62L are thought to be molecules involved in cell homing to bone

marrow or adipose tissue compartments. STRO-1 is expressed by stromal elements in hu-

man bone marrow and valuable for identification and functional characterisation of human

bone marrow stromal cell precursors134. Both cell types are stromal populations isolated from

fatty compartments based on their ability to adhere to plastic.

It remains questionable if CD106, CD117 and ABCG2 are expressed on ADAS cells due to

diverse reports in the literature (Table 1).

In contrary to BMSCs lack ADAS cells the natural activity of telomerase but by transduction

with a retrovirus containing the gene for the catalytic subunit of human telomerase (hTERT)

stable clones can be produced and isolated188. The tranduced clones (hATSC-TERTs) have

high telomerase activity, which could be maintained for more than 100 population doublings

(PD). hATSC-TERTs had a normal karyotype and replicated constantly, whereas control

cells underwent senescence-associated proliferation arrest after 36-40 PD. Supplementary

hATSC-TERTs revealed an increased bone-forming capacity188. These results further sup-

port a similarity between BMSCs and hATSCs as studies with BMSCs demonstrated that

telomerase expression not only extends the proliferative life-span but also maintains the os-

teogenic differentiation potential of these stem cells.

The following table summarizes the surface expression patterns of CD markers for ADAS

cells and BMSCs.

Table 1. Characterization of adipose tissue stem cells compared to bone marrow stem cells

Cluster of Differentiation Surface Expression

Adipose Tissue Stem Cells

BoneMarrow

Stem Cells Reference

CD3 - Strem et al. 2005

CD4 (MHC class II co-receptor; T4) - - Katz et al., 2005, Pittenger et al., 1999

CD8 (MHC class I co-receptor; T8) - Katz et al., 2005

CD9 (tetraspan) + +

De Ugarte et al., 2003, Erickson et al., 2002, Gimble

et al., 2003, Gronthos et al., 2001, Guilak et al.,

2006, Strem et al., 2005, Zuk et al., 2002

CD10 (CALLA) + +

Aust et al., 2004, De Ugarte et al., 2003, Erickson et

al., 2002, Festy et al., 2005, Gimble et al., 2003,

Gronthos et al., 2001, Strem et al., 2005, Zuk et al,

2002,

CD11 (alfa-integrin) - +/-

Aust et al., 2004, Barry et al., 2004, Gimble et al.,

2003, Gronthos et al., 2001, Katz et al., 2005, Pit-

tenger et al., 1999, Suva et al., 2004

CD13 (aminopeptidase) + + Aust et al., 2004, De Ugarte et al., 2003, Erickson et

20


Gronthos et al., 2001, Mitchell et al., 2006, Planat-

Benard et al., 2004, Rodriguez et al., 2005, Strem et

al., 2005, Talens-Visconti et al., 2006, Zuk et al.,

2002

CD14 + low/- -

Baksh et al., 2004, Curat et al., 2004, Gimble et al.,

2003, Gronthos et al., 2001, Jorgensen et al., 2004,

Lee et al., 2004, Minguell et al., 2001, Miranville et

al., 2004, Sengenes et al., 2005, Strem et al., 2005,

Suva et al., 2004, Zuk et al., 2002

CD15 - Rodriguez et al., 2005, Strem et al., 2005

CD16 - Strem et al., 2005, Zuk et al., 2002

CD18 (β-2 integrin) - - Gimble et al., 2003, Katz et al., 2005, Pittenger et

al., 1999

CD29 (β-1 integrin, fibronectin

receptor) + +

Aust et al., 2004, Baksh et al., 2004, Cao et al.,

2005, De Ugarte et al., 2003, Erickson et al., 2002,

Gimble et al., 2003, Gronthos et al., 2001, Guilak et

al., 2006, Katz et al., 2005, Lee et al., 2004, Mitchell

et al., 2006, Pittenger et al., 1999, Silva et al., 2003,

Strem et al., 2005, Zuk et al., 2002

CD31 (PECAM-1) + low/- -

Curat et al., 2004, De Ugarte et al., 2003, Fraser et

al., 2006, Gimble et al., 2003, Mitchell et al., 2006,

Planat-Benard et al., 2004, Strem et al., 2005, Wos-

nitza et al, 2006, Zuk et al., 2002

CD33 - Strem et al., 2005

CD34 (sialoumucin) +/- -

Baksh et al., 2004, De Ugarte et al., 2003, Erickson

et al., 2002, Festy et al., 2005, Fraser et al., 2006,

Gimble et al., 2003, Gronthos et al., 2001, Jorgen-

sen et al., 2004, Koc et al., 2000, Lee et al., 2004,

Minguell et al., 2001, Miranville et al., 2004, Mitchell

et al., 2006, Planat-Benard et al., 2004, Reyes et al.,

2001, Strem et al., 2005, Suva et al., 2004, Wosnitza

et al, 2006, Zuk et al., 2002

CD36 + Festy et al., 2005

CD38 - Strem et al., 2005

CD41a (gpIIb) - Katz et al., 2005

CD44 (hyaluronate receptor or

phagocytic glycpprotein-1) + +

Aust et al., 2004, Baksh et al., 2004, Cao et al.,

2005, De Ugarte et al., 2003, Erickson et al., 2002,

Fraser et al., 2006, Gimble et al., 2003, Gronthos et

al., 2001, Lee et al., 2004, Minguell et al., 2001,

Mitchell et al., 2006, Pittenger et al., 1999, Rodri-

guez et al., 2005, Silva et al., 2003, Strem et al.,

2005, Zuk et al., 2002

CD45 (LCA) +/- -

Aust et al., 2004, Baksh et al., 2004, Curat et al.,

2004, De Ugarte et al., 2003, Fraser et al., 2006,

Gimble et al., 2003, Gronthos et al., 2001, Jorgen-

sen et al., 2004, Koc et al., 2000, Lazarus et al.,

1995, Minguell et al., 2001, Miranville et al., 2004,

Reyes et al., 2001, Strem et al., 2005, Suva et al.,

2004, Zuk et al., 2002

CD49b (alfa-2 integrin) + + Katz et al., 2005, Pittenger et al., 1999, Rodriguez et

21

al., 2005

CD49d (alfa-4 integrin) + -

De Ugarte et al., 2003, Erickson et al., 2002, Fraser

et al., 2006, Gimble et al., 2003, Gronthos et al.,

2001, Katz et al., 2005, Pittenger et al., 1999, Silva

et al., 2003, Strem et al., 2005, Zuk et al., 2002

CD49e (alfa-5 integrin) + +


al., 2002, Gronthos et al., 2001, Katz et al., 2005,

Pittenger et al., 1999, Silva et al., 2003, Strem et al.,

2005, Zuk et al., 2002

CD49f (alfa-6 integrin) - Katz et al., 2005

CD50 (ICAM-3) - - Gimble et al., 2003, Minguell et al., 2001

CD51/61 (vitronectin receptor) +/- +/- Katz et al., 2005, Pittenger et al., 1999, Silva et al.,

2003, Wosnitza et al., 2006

CD54 (ICAM-1) + +


et al., 2003, Gronthos et al., 2001, Strem et al.,

2005, Zuk et al., 2002

CD55 (DAF) + +

De Ugarte et al., 2003, Erickson et al., 2002, Festy

et al., 2005, Gimble et al., 2003, Gronthos et al.,

2001, Strem et al., 2005, Zuk et al., 2002

CD56 (NCAM) - Gimble et al., 2003, Gronthos et al., 2001, Strem et

al., 2005, Zuk et al., 2002

CD59 (complement protein) + +



Gronthos et al., 2001, Strem et al., 2005, Zuk et al.,

2002

CD62E (E-selectin) - - Gimble et al., 2003, Gronthos et al., 2001, Pittenger

et al., 1999, Zuk et al., 2002

CD62L (L-selectin) - + Katz et al., 2005, Pittenger et al., 1999

CD62P (P-selectin) - - Katz et al., 2005, Pittenger et al., 1999, Strem et al.,

2005

CD63 + Mitchell et al., 2006

CD65 + Festy et al., 2005

CD71 (transferrin receptor) + + Baksh et al., 2004, Gimble et al., 2003, Zuk et al.,

2002

CD73 (5´nucleotidase) + Gimble et al., 2003, Mitchell et al., 2006

CD90 (Thy-1) + +

Aust et al., 2004, Baksh et al., 2004, De Ugarte et

al., 2003, Gimble et al., 2003, Jorgensen et al.,

2004, Katz et al., 2005, Lee et al., 2004, Mitchell et

al., 2006, Pittenger et al., 1999, Rodriguez et al.,

2005, Silva et al., 2003, Strem et al., 2005, Suva et

al., 2004, Talens-Visconti et al., 2006, Zuk et al.,

2002

CD104 - Gimble et al., 2003, Strem et al., 2005, Zuk et al.,

2002

CD105 (endoglin) + +

Baksh et al., 2004, Cao et al., 2005, De Ugarte et

al., 2003, Erickson et al., 2002, Fraser et al., 2006,

Gimble et al., 2003, Gronthos et al., 2001, Guilak et

al., 2006, Jorgensen et al., 2004, Knippenberg et al.,

2005, Lee et al., 2004, Majumdar et al., 1998, Rodri-

guez et al., 2005, Strem et al., 2005, Talens-Visconti

et al., 2006, Zuk et al., 2002

22

CD106 (VCAM-1) +/- +

Baksh et al., 2004, De Ugarte et al., 2003, Erickson

et al., 2002, Fraser et al., 2006, Gimble et al., 2003,

Gronthos et al., 2001, Gronthos et al., 2003, Jorgen-

sen et al., 2004, Katz et al., 2005, Majumdar et al.,

1998, Pittenger et al., 1999, Strem et al., 2005, Zuk

et al., 2002

CD117 (c-Kit) +/- +

De Ugarte et al., 2003, Katz et al., 2005, Lee et al.,

2004, Minguell et al., 2001, Rodriguez et al., 2005,

Strem et al., 2005, Zuk et al., 2002

CD133/AC133 + low/- +

Gronthos et al., 2001, Gronthos et al., 2003, Katz et

al., 2005, Rodriguez et al., 2005, (Prentice et al.,

2004)

CD138 (syndecan-1) + Katz et al., 2005

CD140a (PDGFR-alfa) + + Katz et al., 2005, Pittenger et al., 1999

CD144 (VE-cadherin) + low/- Mitchell et al., 2006, Strem et al., 2005

CD146 (Muc18) + +


et al., 2003, Gronthos et al., 2001, Strem et al.,

2005, Zuk et al., 2002

CD166 (ALCAM) + +


al., 2002, Gimble et al., 2003, Gronthos et al., 2001,

Guilak et al., 2006, Knippenberg et al., 2005, Mit-

chell et al., 2006, Strem et al., 2005, Zuk et al., 2002

CD243 (MDR-1) - Katz et al., 2005

ABCG2 +/- + Katz et al., 2005, Mitchell et al., 2006

aldehyde dehydrogenase + Mitchell et al., 2006

alfa smooth muscle actin + + Gimble et al., 2003, Minguell et al., 2001

Collagen I + + Gimble et al., 2003, Minguell et al., 2001

Collagen II + + Gimble et al., 2003, Minguell et al., 2001

Glycophorin A - Rodriguez et al., 2005

HLA-ABC + +

Aust et al., 2004, Baksh et al., 2004, Barry et al.,

2004, Cao et al., 2005, Gimble et al., 2003, Katz et

al., 2005, Planat-Benard et al., 2004, Silva et al.,

2003, Suva et al., 2004

HLA-DR - -

Aust et al., 2004, Barry et al., 2004, Gimble et al.,

2003, Gronthos et al., 2001, Katz et al., 2005, Lee et

al., 2004, Rodriguez et al., 2005, Silva et al., 2003

KDR/Flk1 (VEGF-R2) + Cao et al., 2005, Mitchell et al., 2006, Wosnitza et

al., 2006

Osteonectin + + Gimble et al., 2003, Katz et al., 2005

Osteopontin + + Gimble et al., 2003, Katz et al., 2005, Minguell et al.,

2001

S100 + Atanassova et al., 2001, Kato et al., 1988, Wosnitza

et al., 2006

SH3 + + De Ugarte et al., 2003, Koc et al., 2000, Zuk et al.,

2002

STRO-1 +/- +

Baksh et al., 2004, De Ugarte et al., 2003, Fraser et

al., 2006, Gimble et al., 2003, Gronthos et al., 2001,

Gronthos et al., 2003, Majumdar et al., 1998, Rodri-

guez et al., 2005, Strem et al., 2005, Zuk et al., 2002

telomerase - + Katz et al., 2005

23

Vimentin + + Gimble et al., 2003, Minguell et al., 2001

vWF low + Mitchell et al., 2006, Wosnitza et al., 2006

RELEVANT CD MARKERS AND THEIR EXPRESSION PATTERN

In this thesis, the major aim was to differentiate between cells of endothelial versus adipo-

genic origin, and also to determine the level of maturation of the screened cells. Therefore,

the following markers were selected to be applied for flow cytometry analyses. CD31, also

known as platelet endothelial cell adhesion molecule 1 (PECAM-1), is commonly used for

labeling endothelial cells but is also expressed by monocytes, platelets, lymphocytes, and

granulocytes. CD51/61 labels various types of endothelial cells while vWF is expressed by

endothelial cells and platelets. CD105, or endoglin, is expressed by endothelial cells, acti-

vated monocytes and tissue macrophages, hematopoietic progenitor cells, as well as stromal

cells of certain tissues including bone marrow. Expression of CD105 is increased on acti-

vated endothelium in tissues undergoing angiogenesis and is a component of the TGF-beta

receptor system in human umbilical vein endothelial cells (HUVECs). CD34 is found on he-

matopoietic stem cells, vascular endothelial cells, embryonic fibroblasts and adult nervous

tissue. CD34 is absent on adult hematopoietic cells. VEGF signalling is performed through

KDR or VEGF receptor 2, a receptor tyrosine kinase. KDR is expressed during vasculogene-

sis and hematopoiesis and can be found on circulating endothelial progenitor cells, mature

vascular endothelial cells, and HUVECs. Acetylated-LDL (acLDL) can be applied to differen-

tially label endothelial progenitor cells, vascular endothelial cells, and macrophages. S100

labels nervous tissue, skeletal and cardiac muscle, malignant melanoma cells and some

other cell types. Additionally, it is well accepted as an early marker of adipogenesis21,199.

DIFFERENTIATION CAPACITY TOWARDS MULTIPLE LINEAGES

ADAS cells have been proven to be capable of multilineage differentiation429,433,434. As stem

cells from adipose tissue have their origin in the mesoderm, it is no longer surprising that

these cells can undergo different mesenchymal lineage conversions. Interestingly besides

the classic lineage differentiation, ADAS cells are likewise capable of undergoing transdiffer-

entiation towards the non-mesenchymal branches. By differentiating into neural cells, epithe-

lium, cardiomyocytes, skeletal muscle cells and hepatocytes, these cells show a wide plastic-

ity towards the ectodermal and endodermal lineage (Figure 3 and 4).

Clonal expansion of ADAS cells has proven that at least one part of the plasticity is situated

in a fraction of multipotent cells332,433. Despite pluri- and multipontent populations, fat tissue

harbours above all bi- and unipotent stem cells, solely capable of undergoing only two or one

differentiation pathways261,332.

24

To investigate multipotency, Rodriguez et al. created a single clone from fast-adherent ADAS

cells and proved that two out of 12 clones were able to undergo multilineage differentia-

tion331. The remaining ten clones had bipotent capacity. In contrary to these findings Kang et

al. found that a high percentage of single clones derived from primate ADAS cells were able

to retain adipogenic (82%), osteogenic (64%) and neurogenic (79%) differentiation potential.

The majority of clones (17 out of 20) showed even pluripotency along several mesodermal

and neuronal lineages195. These findings indicate that a high percentage of ADAS cells have

multipotential as well as pluripotential capacity in vitro due to self-renewal and creation of

daughter cells with the potency to convert into the major mesodermal and ectodermal line-

ages. Guilak et al. found that at least 81% out of 45 human ADAS cell clones showed unipo-

tency by differentiating into adipocytes, chondrocytes, osteocytes or neuronal cells138. Fur-

ther 52% of the ADAS cell clones converted into two or more lineages. The major number of

clones expressed phenotypes of osteoblasts (48%), chondrocytes (43%), and neuron-like

cells (52%) than of adipocytes (12%) which could be due to the repeated doublings leading

to the loss of adipogenic differentiation capacity amoung ADAS cells. These findings also

support the hypothesis that ADAS cells are capable of undergoing multipotent differentiation

pathways of the mesodermal and ectodermal lineages and do not solely consist of a mixture

of different unipotent stem cells. However, further investigation on incidence and percentage

between multipotent, bipotent and unipotent adipose mesenchymal stem cells remains es-

sential.

25

adipocytesendothelial cells

chondrocytes

osteocytes

myocyteshepatocytes

cardiomyocytes

neuronal cells

macrophages

ADAS

epithelium

Figure 3. Multiple differentiation pathways of human SVF cells

Human SVF cells convert into multiple cell lineages under appropriate culture conditions, including fat tissue, endothelial cells,

chondrocytes, osteocytes, myocytes, epithelial cells, hepatocytes, cardiomyocytes, neuronal cells and macrophages.

26

proliferationdifferentiation

chondrocyte

endothelial cell

osteocyte

myocyte

other tissues

ADAS

Figure 4. Scheme of SVF cell differentiation

SVF cells are capable to perform self-renewal and additionally to differentiate into numerous different cell types. The

lineage-committed cell progresses through several stages of maturation prior to the onset of terminal differentiation,

which is marked by the cessation of proliferation and shift towards increased expression of tissue-specific markers.

27

MESODERMAL DIFFERENTIATION

ADIPOGENESIS

Adipose-tissue derived adult mesenchymal stem cells naturally differentiate into mature adi-

pocytes. In adipogenic induction medium, ADAS cells develop intracellular lipid vacuoles

which coalesce and give rise to a single, cytoplasma-filling vacuole. Besides this definite

marker of adipogenesis349,414, several fat cell specific genes are expressed, comprising glyc-

erol-3-phosphate dehydrogenase (GPDH)216, lipoprotein lipase127,272, peroxisome proliferator-

activated receptor γ (PPARγ)366, leptin246, adipocyte fatty acid binding protein (aP2)11,

CCAAT/enhancer binding protein (C/EBP) and glucose transporter 4 (Glut4) (Figure 5)149.

GPDH is a key enzyme in adipose progenitor cell differentiation335 and has been widely ap-

plied for decades to measure the degree of adipocyte conversion298,368. The occurrence of

the genes given above in ADAS derived fat cells personates a very analogical pattern to that

noted in adipogenesis in MSCs82,308,312.

Adipocyte conversion of ADAS cells is not exclusively mined by miscellaneous growth factors

and the absence of serum (which acts as inhibitor during differentiation), but also by extracel-

lular matrix proteins, like fibronectin and collagen among others369.

ADAS proliferation and differentiation are two intimately connected processes, in adipogene-

sis as well as differentiation into other cell types (compare also Figure 4). Long-term prolif-

eration in fetal calf serum (FCS)-supplemented medium notedly and irreversibly decrease

ADAS cell potency to undergo fat differentiation149,158,160. Contrary to FCS, human autologous

serum (hS) enables long-term proliferation as well as subsequent differentiation, even after

prolonged culture and doubling rates160. Some studies support a composite of FCS with hS,

due to the fact that ADAS cells cultured in hS alone were found to proliferate weekly123, did

not attach to the culture flask75, or were not viable148. Hence the inhibitory effect of prolonged

proliferation periods in FCS-supplemented medium is most likely due to serum factors in the

FCS, varying between sundry charges and unpredictable to change or control. According to

this, a serum-free culture medium would be highly desirable136,202,387.

Optimal differentiation results are achieved when adipogenic induction medium is added at

maximal cell confluence. Also, an age-related decrease in proliferation and differentiation

capacity in ADAS cells has been observed7,147,198.

During acquisition of the adipocyte phenotype chronological changes in the expression of

several genes appear, including early, intermediate and late mRNA and protein markers as

well as triglyceride accumulation (see Figure 5)57,68,247,271,286.

28

Cell confluence and cell-cell contact both improve adipocyte conversion but are not manda-

tory for fat differentiation298. Growth arrest, in contrary, appears to be inevitable for differen-

tiation of ADAS cells into mature adipocytes.

Adipocyte specific genes are mainly transactivated by two transcription factors, PPAR-γ and

CCAAT/enhancer binding protein α (C/EBP-α)72,93,223,366. Both of them are also involved in

growth arrest required for adipocyte conversion. Furthermore C/EBP-α acts as an antimitotic

agent and leads to growth inhibition390,396. PPAR-γ is capable of inducing growth arrest in

fibroblasts and other cells, leading to cell cycle withdrawal9. PPAR-γ and C/EBP-α seem to

act cooperatively to bring about growth arrest that precedes differentiation9.

After withdrawal from the cell cycle in the early stage of adipocyte development, premature

signs of differentiation include the expression of lipoprotein lipase (LPL) mRNA4,83,132,247,272.

LPL is secreted in high amounts by mature fat cells and plays a central role in controlling lipid

accumulation89,129. LPL expression is independent of the addition of adipocyte differentiation

substrates and emerges spontaneously at cell confluence12,13. Therefore LPL occurrence

may display growth arrest rather than an early marker of adipogenesis and is not adipocyte

specific90,115,386. PPAR-γ and C/EBP expression is significantly upregulated during early adi-

pocyte differentiation, although low levels of these factors are also expressed in undifferenti-

ated ADAS cells. Both are subsequently involved in terminal differentiation by transactivation

of adipocyte specific genes and maximal levels of expression are attained in mature fat

cells54,71,72,420. During conversion into adipocytes, ADAS cells change their morphology from a

fibroblast-like shape to a spherical appearance. This process of lipid accumulation also in-

cludes major changes in cytoskeletal components, like actin and tubulin among others88,367,

as well as the level and types of extracellular matrix elements276 221,277. These changes in

cell shape reflect a distinct process in differentiation and are not results of lipid vacuole ac-

cumulation.

Two major events in terminal adipogenesis are de novo lipogenesis and acquisition of insulin

sensitivity. Lipogenic enzyme levels, including mRNA and protein expression, and activity of

triglycerol metabolism, increase 10- to 100-fold during final differentiation into adipocytes.

Maturation comprises mainly glycerol-3-phosphate dehydrogenase, glycerol-3-phosphate

acyltransferase, glyceraldehydes-3-phosphate dehydrogenase, fatty acid (FA) synthase, ace-

tyl-CoA carboxylase, stearoyl-CoA desaturase (SCD1), adenosintriphosphate (ATP) citrate

lyase and malic enzyme306,368,411. Insulin receptor numbers, insulin sensivity and glucose

transporter levels rise as well121,224,324,337. Furthermore, total number of adrenergic receptors

is upregulated during adipogenic differentiation, specially an increase in the β2- and the β3-

subtypes110,111,137,222. Leptin as well as aP2, an adipocyte-specific fatty acid binding protein,

29

are likewise increased during terminal adipogenesis11,40,73,246,368. PPAR-γ and C/EBP-α are

incorporated in the coordinate activation of several of these genes, comprising leptin, aP2,

Glut4 and SCD1 inter alia166,245,256,392,393.

growth arrestpost-confluent mitoses

clonal expansionearly late differentiation

Pref-1C/EBPβ

PPARγ

C/EBPα

adipocytegenes

lipogenicenzymes

FA bindingproteins

secretedfactors

ECM alteration cytosceletal remodelling

Although mature adipocytes were thought to remain in terminal differentiation without pro-

gressing towards dedifferentiation nor re-entering mitosis410,415, several studies support the

capacity of mature fat cells to undergo progressive dedifferentiation back to the primary

ADAS cell type378. Dedifferentiation followed by proliferation15,378 and subsequent differentia-

tion back into adipocytes355,377,379 as well as other lineages like endothelial cells have been

reported (Figure 4)314. A true reversion to an ADAS bi- or even multipotential cell phenotype

could play a role in the regulation of adipocyte and endothelial cell numbers and conse-

quently adipose tissue mass.

CHONDROGENESIS

Under chondrogenic medium conditions ADAS cells establish cellular nodules that produce

cartilage-related extracellular matrix molecules: collagen type II, VI, sulfate proteoglycans,

e.g. chondroitin 4-sulfate and keratan sulfate, proteoglycan aggrecan and proline/ arginine-

rich end leucine-rich repeat protein (PRELP) in vitro61,103,107,175,243,291,312,414,417,433,434. Optimal

conditions for differentiation are high density micro mass cultures supplemented with trans-

forming growth factor beta (TGF-β) and insulin, combined with dexamethasone, ascorbate-2-

phosphate, sodium pyruvate, or combined with transferrin, to name the most common possi-

bilities24,107,175,243.

Figure 5. Adipocyte

differentiation

Chronological changes in

the cellular processes and

transcriptional cascade

during fat cell matura-

tion.

30

ADAS cells seeded on scaffolds and subsequently implanted produce cartilage matrix in

vivo. Chondrogenic differentiation is influenced by different cell-biomaterial interactions, de-

pendent on the material applied. The biomaterial not only influences the chondrogenic differ-

entiation but also the mechanical properties of the construct25. ADAS cells seeded onto algi-

nate carriers followed by implantation into immunodeficient mice posses prolonged synthesis

of cartilage matrix molecules over 12 weeks107. In vivo, engineered cartilage contains ECM

molecules such as aggrecan, collagen II and collagen VI107,243. Besides the chondrogenic

growth factors and the ECM environment, oxygen tension also plays an important role in

chondrogenic differentiation. As cartilage is an avascular tissue, ADAS cells possible un-

dergo hypoxia during cartilage formation in vivo. ADAS cells differentiate in both, normoxic

as well as hypoxic environment, but hypoxia strongly reduces in vitro chondrogenesis248.

It remains controversial whether ADAS cells or MSCs are superior for cartilage engineer-

ing98,179,279,417. However, ADAS cells seem to have a greater chondrogenic potential than

MSCs when applied for engineering of cartilage in vitro as well as in a rabbit osteochondral

defect model98,279.

OSTEOGENESIS

ADAS cell differentiation towards the osteogenic cell lineage is well established for in vitro

as well as for in vivo animal tissue engineering mod-

els102,103,141,145,163,174,194,213,230,248,291,388,421,433,434. A clinical obervation is in part responsible for

the discovery of the osteogenic differentiation capacity of ADAS cells. A rare disorder named

“progressive osseus heteroplasia” together with the capacity of MSCs to convert into the os-

teogenic lineage led to the asssumption that ADAS cells are likewise able to differentiate into

osteocytes36,38,53,134,142,164,179,287,297. Patients with “progressive osseous heteroplasia” develop

calcified nodules representing normal bone which is placed at ectopic soft-tissue locations

including subcutaneous fat depots22,197,257,356,401.

Osteogenic induction of ADAS cells can be achieved by similar culture conditions as used in

MSCs, including supplementation with ascorbic acid together with 1-alpha,25-

dihydroxyvitamin D3 (1,25(OH)2D3), the hormonal metabolite of vitamin D, or

dexamethasone194,213,291. Under osteogenic differentiation medium ADAS cells are capable of

expressing diverse genes and proteins found in the osteoblast’s phenotype: type I collagen,

alkaline phosphatase, osteocalcin, osteonectin, osteopontin, parathyroid hormone (PTH)

receptor, bone morphogenetic protein-2 (BMP-2), BMP-4, BMP receptors I and II, bone sialo

protein and RunX-1141,213,291,433. Furthermore, ADAS cells exhibit a bone-like response to fluid

shear stress in vitro, achieved with mechanical loading by pulsating fluid flow (PFF). The re-

sult is increased nitric oxide (NO) production and cyclooxygenase-2 expression213. This might

indicate a bone-cell specific function during bone remodelling in vivo. ADAS cells are also

31

competent to generate mineralized matrix in vitro in both long term 2D or 3D osteogenic cul-

tures.

Human ADAS cells are able to generate bone tissue in immunodeficient rodent ectopic bone

models when seeded onto several different carrier scaffolds103,145,163,231. Furthermore these

cells have proven to regenerate bone defects in in vivo animal models81,85,104,and even in

humans when applied in a clinical trial for treatment of sizeable bilateral calvarial defects235.

Due to the restricted amount of bone marrow stem cells available in the patient with calvarial

defects, ADAS cells were applied to the defect in a single procedure together with autolo-

gous iliac crest bone and fibrin glue combined with resorbable sheets. CT-scans three

months after the reconstruction showed new ossification as well as nearly complete calvarial

continuity235.

ADAS cells have a capacity for bone formation that is comparable to bone marrow MSCs.

They maintain their osteogenic potential with aging when conferred to juvenile cells354.

SKELETAL MYOGENESIS

Cultured under appropriate myogenic medium conditions, ADAS cells undergo conversion

into a skeletal muscle cell phenotype in vitro263,332,433. Addition of hydrocortisone, dexa-

methasone and 5-azacytidine or coculture with primary skeletal myocytes induces myocyte-

related gene expression. Upregulated are key regulatory master factors like myogenic factor

5 (myf5), myf6, myogenic differentiation 1 (MyoD1) and myogenin232,263,433. ADAS cells also

respond with muscle cell differentiation in a myogenic environment, as they can be incorpo-

rated into muscle fibers of mdx mice after ischemia restoring dystrophin expression signifi-

cantly101.

Evidence exists that ADAS cells are a promising tool for restoration of muscle function in vivo

as well. ADAS cells attached to injectable poly(lactic-co-glycolic acid) (PLGA) spheres in-

jected subcutaneously into the necks of nude mice after culture in myogenic medium, gener-

ated new muscle tissue in vivo210. When transplanted into damaged regions of rabbit skeletal

muscle, ADAS cells increased fiber cross section area and muscle weight, resulting in sig-

nificantly raised contractile force, compared to the injured control group27. Furthermore, there

is striking indication that ADAS cells could be useful for the treatment of Duchenne muscular

dystrophy. When injected into the tibial and gastrocnemius muscle in dystrophin-deficient

mdx mice, an animal model for Duchenne muscular dystrophy, these cells induced a long-

term engraftment and a high expression of human dystrophin332.

CARDIOMYOGENESIS

The use of stem cells to regenerate damaged heart tissue is advocated as the new treatment

for heart failure secondary to heart disease or severe myocardial infarction. There are prom-

32

ising results of in vitro studies achieving myocardial differentiation in ADAS cells112,124,313,320.

When cultured in semisolid methylcellulose medium under low serum conditions and sup-

plemented with 2-mercaptoethanol, insulin, transferrin, interleukin-3 (IL-3), IL-6 and stem cell

factor (SCF), cohesive cell groups form and display branching fibers, sharing tight connec-

tions and beating at a single rate. These cells express cardiomyocyte-specific markers in-

cluding myosin-enhancing factor 2C, sacromeric α-actinin, ventricular and atrial myosin light

chain MLC-2v and MLC-2a, β-myosin heavy chain, atrial natriuretic peptide (NAD) and con-

nexin43 as well as connexin40313. In a murine in vivo model of acute myocardial injury, donor

ADAS cells were injected into the intraventricular chamber instantly after application of cryo-

injury to the myocardium373. Donor ADAS cells were identified exclusively in infarct areas of

animals treated with ADAS cells, but not in animals without myocardial damage or in the con-

trol group without cell treatment. Areas containing donor cells were positive for Heavy Chain,

Nkx2.5 and Troponin I373. So far this is one of the first study demonstrating that ADAS cells

have the potency both to undergo cardiomyocytic conversion and to engraft infracted myo-

cardium in vivo373. These are promising results for future application of ADAS cells in myo-

cardial infarction and other cardiovascular diseases, that are one of the leading causes for

morbidity and mortality in western countries.

IMMUNE CELLS

Preadipocytes as well as adipocytes secrete numerous inflammatory cytokines, such as tu-

mor necrosis factor α (TNF-α), and can be activated by lipopolysaccharide binding which

results in immunomodulatory molecule secretion56,169,207,208,242. Furthermore, PPARγ and aP2,

initially believed to be highly specific for adipocytes, were also found in macrophages309. In-

terestingly, preadipocytes were discovered to be genomically closer related to macrophages

than adipocytes69. Recent studies have demonstrated that ADAS cells display common func-

tional features of monocytes and macrophages by revealing specific phagocytosis activity

including antimicrobial activity via an oxygen-dependent mechanism342. These cells likewise

express CD80, CD86, CD45, anti perivascular monocyte/ macrophage antibody (ED2), anti-

macrophage monoclonal antibody-2 (MOMA-2), F4/80 and macrophage antigen-1 (Mac-1)/

CD11b69,84. In vivo transdifferentiation after grafting ADAS cells into the peritoneal cavity al-

lowed a transdifferentiation rate into macrophages of about 70% of cells69.

33

ECTODERMAL DIFFERENTIATION

NEUROGENESIS

The ability to undergo in vitro and in vivo neuronal lineage differentiation is another property

of ADAS cells that may have impact on CNS tissue regeneration113,195,196,385. With treatment

of different supplement combinations, ADAS cells achieve neurosphere formation when cul-

tured at high density195. Subsequent culture of the spheroid bodies on laminin leads to final

neural differentiation195. Differentiated cells protrude extensive cell processes with a neuronal

morphology and express neural markers, including nestin, neuronal nuclear protein (Neu N),

neurofilament (NF) 160, microtubule associated protein 2 a/b (MAP2ab) and the mature as-

trocyte marker glial fibrillary acidic protein (GFAP)195. While nestin and NF160 belong to the

early markers of the neuronal lineage, MAP2ab, Neu N as well as GFAP are thought to

match with terminal conversion of ADAS cells in neurogenesis195. Several protocols exist in

the literature to induce neurogenic conversion and achieve expression of neuronal-specific

markers20,117,192,339,341,419,423,433. Furthermore, positive staining for functional neuronal tissue,

comprising the neurotransmitter gamma-amino butyric acid (GABA), both subunits of the

glutamate N-methyl-D-aspartate (NMDA) receptor and α-1 voltage gated calcium channels,

was detected in neural differentiated ADAS cells341. Still it remains to be investigated if such

neurons are electrically active and functional with all essential characteristics of mature CNS

neurons364.

So far there are a few promising in vitro studies using ADAS cells for treatment of cerebral

ischemia, spinal cord injury as well as other neurological disorders193,196,340. ADAS cells mi-

grate into injured cerebral lesions after ischemic stroke and improve functional deficits and

motor recovery193. When applied in a rat model of spinal cord injury, ADAS cells migrate into

the damaged regions and significantly improve motor function after 4-5 weeks196.

Although neurogenesis is known to occur in adult life in limited regions of the brain119,385,

adult neuronal stem cells remain a population unattainable for clinical applications3,119. There-

fore ADAS cells display a highly promising source for cerebral injury treatment in the future.

EPITHELIAL DIFFERENTIATION

ADAS cells are known to accelerate epidermal regeneration when seeded together with

keratinocytes in a cutaneous reconstructive model. Furthermore, ADAS cells inhibit keratino-

cyte apoptosis and induced rete ridge- and epidermal ridge-like structure formation14. Fur-

thermore ADAS cells improve dermal regeneration in full thickness skin defects and reduce

wound contraction and subsequent scarring99. Treatment of human ADAS cells with all-trans

retinoic acid (ATRA) initiates cytokeratin 18 expression and the formation of keratin fibers

34

indicating epithelial differentiation. As demonstrated by Brzoska et al, more than 80% of cells

converted to an epithelial lineage cell type after incubation with ATRA55.

ENDODERMAL DIFFERENTIATION

HEPATOGENIC AND INTESTINAL DIFFERENTIATION

Although primary hepatocytes, liver stem cells and bone marrow stem cells represent the

main sources for liver regeneration, ADAS cells have recently been transdifferentiated into

hepatocyte-like cells351. Under treatment with a cytokine mixture and dimethylsulfoxid

(DMSO), ADAS cell conversion into the hepatic lineage was confirmed by the expression of

albumin, α-fetoprotein, as well as functional characteristics comprising low-density lipoprotein

(LDL) uptake and urea production. After transplantation into a hepatic injury mouse model,

ADAS cells differentiated into hepatocytes in vivo351. Other groups applied more complex

differentiation protocols to induce hepatic lineage differentiation in both ADAS cells and

MSCs, again finding upregulation of several liver-markers including albumine, cytochrome

P450 (CYP) isoenzymes, α-fetoprotein, C/EBPβ and hepatocyte nuclear factor 4 alfa

(HNF4α), which confirmed cellular conversion into hepatocytes381-383.

Autologous stem cell transplantation of ADAS cells has also been used for the treatment of

Crohn’s fistula in a phase I clinical trial122. Although a phase I trial does not allow final con-

clusions about effectiveness, the results achieved appear highly promising. After inoculation

of fistulas with autologous ADAS cells, six fistulas out of eight showed coverage with epithe-

lium at the external opening. A healing success was obtained in 75% and no adverse effects

were observed during the follow-up period of in average 22 months122.

VASCULOGENESIS AND ANGIOGENESIS

De novo blood vessel development in the embryo comprehends two different mechanisms,

vasculogenesis and angiogenesis. Vasculogenesis enfolds the primary vascular plexus

formed out of the mesoderm by differentiation of angioblasts followed by the generation of

primary blood vessels329. The fibroblast growth factor (FGF) family is crucial to conduct the

mesoderm in forming of angioblasts as well as hemangioblasts. Vascular endothelial growth

factor (VEGF) and its receptor tyrosine kinase VEGF-Receptor 2 (VEGF-R2/KDR/fetal liver

kinase 1 (flk-1)) represent a signalling system which is highly important for the differentiation

of endothelial cells and the development of the vascular system. Specified cell adhesion

molecules such as vascular endothelial cadherin (VE-cadherin) and platelet endothelial cell

adhesion molecule 1 (PECAM-1/CD31), as well as mechanical forces and vascular regres-

sion with remodelling, participate in the process of endothelial cell conversion, apoptosis, and

35

succeeding angiogenesis. Branching of several capillaries and pruning of others during vas-

culogenesis leads to de novo blood vessel formation and subsequently to a mature vascular

network.

Following the primary vascular plexus formation, endothelial cell differentiation and prolifera-

tion cause sprouting and splitting of blood vessels with subsequent development of new cap-

illary structures, an incident called angiogenesis. Angiogenesis consists of two different

types, sprouting angiogenesis from pre-existing capillaries and non-sprouting angiogenesis

or intussusception. Both require VEGF as a trigger for endothelial cell proliferation and mi-

gration62,63,328. Angiogenesis is a highly complex mechanism and displays an invasive cellular

process which is dependent on several growth factors, proteolytic enzymes, extracellular

matrix proteins and adhesion receptors46,143,144,180.

Primarily, it was adopted that vasculogenesis is a process constricted to embryonal devel-

opment, but recent studies proved that postnatal vasculogenesis takes place in the adult or-

ganism with the help of endothelial progenitor cells (EPCs)353. EPCs have a similar profile

when compared to an embryonic angioblast and these stem cells respond to angiogenic

growth factors with proliferation and migration forming in situ blood vessels via endothelial

differentiation. They incorporate into foci of neovascularization, thereby demonstrating that

postnatal neovascularization consists of both, angiogenesis as well as vasculogenesis16.

ANGIOGENESIS IN ADIPOSE TISSUE

The development and maturation of adipose tissue vascularity are critical to the function of

adipose tissue87,315. In fact adipose tissue begins to develop in well vascularized regions316

where adipocyte development is characterized by the appearance of a number of fat cell

clusters, termed “primitive organs”. Primitive fat organs are vascular structures in presump-

tive adipose tissue with few or no fat cells and increase throughout fetal development in

number and size. Fetal fat cell development is locally and temporarily dependent on capillary

formation. Arteriolar development clearly precedes adipocyte differentiation in fat tissue and

can be used to distinguish adipose tissue depots in the fetus87. Vascular development and

endothelial cell proliferation rate seem to be fat depot-dependent as endothelial cells from

several different rat fat depots have different replication rates after multiple passages226,227.

ADIPOGENESIS AND ANGIOGENESIS

Endothelial cells and preadipocytes share mutual developmental and autocrine as well as

paracrine relationships. Capillary formation in fat depots emerges synchronistically within the

early stages of adipogenesis, characterized by appearance of perivascular preadipo-

cytes152,184. Another evidence is that both endothelial cells and preadipocytes express αvβ3

integrin and secrete plasminogen activator inhibitor-1 (PAI-1), adjusting endothelial cell and

36

preadipocyte migration in vitro86. αvβ3 integrin is critical for vasculogenesis and angiogenesis

during development and mediates EC attachment, spreading and migration239,299. It binds

vitronectin, fibronectin and fibrin. PAI-1 regulates smooth muscle and endothelial cell motility

and migration through the interaction with vitronectin and its receptor (CD51/61) as well.

Preadipocyte-derived PAI-1 production ensures organization and coordination of adipogene-

sis and angiogenesis at local levels. Therefore preadipocytes are able to migrate together

with sprouting capillary endothelial cells in the formation of the primitive fat organ86.

Sprouting of new blood vessels is dependent on basement membrane protein degradation422.

The breakdown and degradation of connective tissue and basement membrane proteins be-

long to the critical phase of angiogenesis. Two major proteolytic enzymes are crucial for ex-

tracellular protein proteolysis, the matrix metalloproteinases (MMP) and the plasminogen

activator (PA)-plasmin system239,299,310. Inhibition of MMP activity leads to reduced or blocked

angiogenesis, indicating the implication of extracellular proteolysis in basement membrane

degradation, cell migration and ECM invasion, capillary lumen formation, regulation of cyto-

kine activity, and release of membrane bound VEGF and TNFα310,311.

MMP-2 and MMP-9 as well as their tissue inhibitors (TIMP) are expressed by ADAS cells

and adipose tissue in a depot-dependent manner51,70,100,120,251. MMP and TIMP significantly

influence adipose tissue development by adipocyte differentiation but may do so in coopera-

tion or independent of angiogenesis51,70,88,240.

Preadipocytes secrete PAI-1 in a depot-dependent manner32. PAI-1 inhibits both plasmino-

gen activators, t-PA and u-PA, and influences the migration of endothelial cells, smooth

muscle cells, and preadipocytes themselves86,311. Additionally, hypoxia induces PAI-1 ex-

pression in endothelial cells311. Elevated PAI-1 levels in the circulation are involved in the

development of the metabolic syndrome with insulin resistance and increased risk for cardio-

vascular disease8. Glitazones (insulin sensitizer) reduce human fat PAI-1 secretion leading to

improved endothelial function in chronic cardiovascular disease states300.

Both VEGF and FGF act as mitogenic agents in endothelial cells and preadipocytes by pro-

moting cell proliferation, migration and pericellular proteolysis (Figure 6). Their interaction on

angiogenesis is closely linked together41,389,422. VEGF acts through two tyrosine kinase recep-

tors, VEGFR-1 (fms like tyrosine kinase 1 (flt-1)) and VEGFR-2 (flk-1/KDR), which are mainly

expressed by endothelial cells, endothelial and hematopoietc progenitor cells and mono-

cytes. VEGF and its receptors both influence EC proliferation, migration, tubulogenesis, sur-

vival, vascular permeability, MMP proteinases, and the PA system. However, it seems to be

primarily VEGFR-2 which is crucial for angiogenesis. Besides, this receptor can be upregu-

lated in ECs by bFGF supplementation, indicating the synergism between these two growth

factors389. FGF may play a role in angiogenesis during tissue repair299 and enhances adipo-

37

cyte differentiation in vivo203. VEGF gene expression is influenced by a number of factors,

comprising hypoxia, insulin, growth factors and several different cytokines239.

VEGF is considered to be the major factor of the angiogenic activity of media conditioned by

adipocytes and adipose tissue432. Fat tissue expression and secretion of VEGF is induced by

hypoxia as well as insulin255,432. Vice versa insulin may be involved in increased adipose tis-

sue VEGF expression during rebound weight gain after diet restriction in rabbits268. Angio-

poietin (Ang)-1, Ang-2, and Tie 1 and Tie 2 receptors are regarded VEGF’s most fundamen-

tal partners (compare Figure 6)299,422. Ang-1 stabilizes blood vessel walls, whereas Ang-2

promotes destabilization. Hypoxia, VEGF, FGF, and cytokines induce Ang-2 expression in

microvascular ECs249,292. Several other studies have demonstrated the possibility that leptin

induces angiogenesis or influences angiogenic factors50,59,128,187,220,289,319,358,374,431. Both adi-

pose tissue and endothelial cells express the receptor for leptin binding49,50 and leptin is pro-

duced and secreted by adipocytes. Adipose tissue leptin and VEGF expression are similarly

regulated by hypoxia and insulin10,422. Therefore, leptin may potentiate VEGF-mediated an-

giogenesis by indirectly influencing VEGF secretion and uptake by endothelial cells302.

Several other growth factors are involved in angiogenesis as well, e.g. PDGF-β, TGF-β, TNF-

α and FGF, promoting vessel wall stability, establishing the integrity of capillary walls during

sprouting, and promoting tube formation in ECs (Figure 6)299,322.

38

Ang-1FGF

TGF-βPDGF

VEGF

EC

SMC

ECM

Strong evidence suggests adipogenesis to be influenced by angiogenesis and vice

versa338,358,360. Adipocyte endothelial cell interaction is thus presumed to be involved in the

development and maintenance of fat tissue. Also, adipose tissue itself is an origin of several

different vasoactive factors116. A close evolutionary relationship between preadipocytes and

ECs can be assumed since preadipocytes have been demonstrated to influence EC prolif-

eration, reduce apoptosis325, and increase mature EC viability and migration274. Impact on

proliferation and subsequent enhancement of tube formation, mainly meditated by secretion

of VEGF, TGF-β, bFGF and HGF by preadipocytes48,77,274,275,325. The expression of

leptin50,358, Ang 1370, Ang 278,139 and PAI-1345,365 by adipose tissue also has impact on angio-

genesis, as mentioned above, and the activity of these factors is interrelated. Leptin for in-

stance induces the expression of Ang 278 and enhances VEGF-mediated angiogenesis59.

Furthermore, preadipocytes are able to increase the amount of VEGF under hypoxia325 and

during fat pad development280. However, not exclusively adipocyte precursors are capable of

Figure 6. Regulation of blood vessel morphogenesis

during neovascularisation

New vessels arise from pre-existing capillaries and post-

capillary venoles. Pericytes start to migrate and blood

vessels dilate. Afterwards the basal membrane and the

extracellular matrix is degraded. Now endothelial migrate

into the perivascular space along the angiogenic stimulus

mediated through VEGF and its receptors, VEGF recep-

tor 1 and 2. Endothelial cells begin to proliferate and form

capillary-like structures including a lumen. TGF-β, FGF

and PDGF function as angiogenic growth factors as well.

FGF and VEGF function as mitogenic agents in ECs by

promoting cell proliferation, migration and pericellular

proteolysis. PDGF possesses a structural affinity with

VEGF. Ang-1 navigates the interaction between endothe-

lial cells and smooth muscle cells as well as pericytes by

stabilising the freshly formed blood vessels and acting as

an anti-permeability factor.

39

enhancing angiogenesis. Mature adipocytes also produce significant amounts of VEGF66.

MMP-9 which is upregulated in differentiated adipocytes51,204 increases the availability of ma-

trix-bound VEGF39. These findings indicate that mature fat cells, which require a very in-

tensely branched vascular network with direct contact to ECs, mediate endothelial cell ex-

pansion, migration and tube formation by VEGF secretion to optimize their oxygen supply

(Figure 7). This is supported by the fact that fat cells have a significant lower tolerance for

hypoxic conditions than preadipocytes and require higher concentration of oxygen406.

hypoxia

leptin

VEGF

TGF-β

FGF

PAI-1

TIMP

MMP-2MMP-9

proteolysis(MMP& PA)

angiogenicsprouting

cytokines Ang 1 ↑

Reciprocally, arteriolar differentiation precedes adipocyte development during fetal fat pat

formation87,151,154. Endothelial cell ECM maturation precedes differentiation of adipocyte

ECM87,153,156 and endothelial cells release ECM components that enhance preadipocyte dif-

ferentiation402. Remodelling of ECM components is important for both adipogenesis241 and

angiogenesis64. Preadipocyte spreading and migration during adipogenesis may be en-

hanced or even induced by blood vessel differentiation155. As VEGF expression can be gen-

erated by cold exposure in BAT18,19 and increased adipogenesis is promoted by enhanced

VEGF expression during rebound weight gain after diet restriction268, angiogenesis also

seems to play a crucial role in postnatal adipogenesis. Furthermore extracellular proteolysis

may be able to adjust fat tissue angiogenesis as well as fat cell maturation and develop-

ment167,267. Fat tissue proliferation is dependent on angiogenesis and can be regulated

through adipose tissue vascularization338. Anti-angiogenic agents leading to weight loss in

mice, as demonstrated by obesity models338, and are further proof of the close relationship

between endothelium and fat tissue. Several in vitro studies abet these findings, as ECs

Figure 7. Adipose angio-

genesis

Tissue hypoxia enhances

preadipocyte and adipocyte

secretion of proangiogenic

factors. VEGF, FGF, TGF-β

together with other cyto-

kines increase capillary

formation through EC

sprouting and Ang-1 ex-

pression. TIMP, MMP,

PAI-1 further support

vessel growth by tissue

proteolysis and tube forma-

tion.

40

promote adipose tissue derived precursor cell proliferation in conditioned medium177. Direct

adhesion of ECs promotes the active development of immature preadipocytes together with

increased growth of mature adipocytes15. These findings support the view that endothelium

closely regulates adipose tissue growth and homeostasis.

In conclusion, modulating the development of the vascular network in adipose tissue in either

an anti-angiogenic or pro-angiogenic manner may provide a strategy to solve problems with

obesity and cardiovascular complications on the one hand67, on the other hand may help

finding solutions for soft tissue engineering approaches to reconstruct defects after trauma or

tumor removal.

41

STUDIES

In this thesis, the differentiation plasticity and capacity of stromal vascular fraction cells to-

wards adipogenic and endothelial lineage was analyzed. The study was divided into the fol-

lowing six tasks:

1. Comparison of SVF cells from excised fat tissue versus liposuction material 2. Analysis of surface protein expression in SVF cells during the culturing process 3. Differentiation of unpurified SVF cells under endothelial medium conditions 4. Splitting of SVF cells in CD31- and CD31+ SVF cells and characterisation of purified frac-

tions 5. Differentiation potential of CD31- and CD31+ SVF cells towards the endothelial lineage 6. Differentiation potential of CD31- and CD31+ SVF cells towards the adipogenic lineage

1. Comparison of SVF cells from excised fat tissue versus liposuction material Two distinct isolation methods for gaining cells from the adipose SVF are established in the

literature, one from excised fat tissue and one from liposuction aspirates. Due to the harvest-

ing procedure liposuction material does not exclusively contain AT but also blood vessels

and fibrous tissue. Isolation of SVF cells from excised AT, in contrast, involves the manual

removal of capillaries and fibrous tissue. In order to prescind the differences in heterogenity

between SVF cells isolated from liposuctioned versus excised AT, freshly harvested SVF

cells from both isolation methods were analysed with flow cytometry. To quantify the amount

of endothelial cells, hematopoietic stem cells, and other cells contained in the two cell frac-

tions, antibodies against CD31, CD51/61, CD34 and KDR were applied.

2. Analysis of surface protein expression in SVF cells during the culturing process Not only the harvesting method but also the culture conditions influence the SVF cell charac-

teristics and their heterogenity. Long-term culturing of SVF cells generates high cell numbers

but changes major preadipocyte characteristics, i.e. protein surface expression and differen-

tiation capacity.

To analyse the changes in surface protein expression during culture of SVF cells in standard

medium, flow cytometry analyses were performed with stem cell markers for CD34, KDR and

S100 after 7 and 14 days.

42

3. Differentiation of unpurified SVF cells under endothelial medium conditions Usually, the harvested SVF is contaminated with ECs. When measuring preadipocyte trans-

differentiation potential to ECs, ECs embodied in the isolated SVF can mimic a successful

differentiation result even though ECs were already present in the applied cell fraction before

differentiation induction. To determine the growth potential of contaminating endothelial cells

after isolation, various growth media were tested including different EC supporting media

(EC-1, -2, -3). These culture media included standard EC media frequently used in the litera-

ture.

The first culture medium (EC-1) applied was an endothelial lineage differentiation medium for

angiogenic progenitor cells and contained Endothelial Growth Medium-2 Microvascular

(EGM-2MV) with 10 ng/ml VEGF and 10-6 mol/l rosiglitazone. The second medium (EC-2), an

endothelial differentiation medium, was Endothelial Cell Growth Medium (ECGM) supple-

mented with 50 ng/ml VEGF, 20 ng/ml IGF-1, and 2% FCS. The third culture medium (EC-3)

tested was a hematopoietic and endothelial progenitor cell medium for peripheral blood pro-

genitor cells containing IMDM, 50 ng/ml VEGF, 100 ng/ml SCGF-ß, 30% FCS, 1% BSA, and

10-4 mol/l ß-mercaptoethanol.

4. Splitting of SVF cells in CD31- and CD31+ SVF cells and characterisation of purified frac-

tions In order to seperately investigate the differentiation potential of both CD31- and CD31+ SVF

cells towards the adipogenic and endothelial lineage, splitting with paramagnetic beads was

performed (see studies 5. and 6.). To confirm the purity of both separated cell fractions, flow

cytometry analyses for CD105, vWF, CD34 and S100 were carried out in addition to vWF

immunofluorescence staining and histological evaluation. HUVECs were used as positive

control, fibroblasts as negative control.

5. Differentiation potential of CD31- and CD31+ SVF cells towards the endothelial lineage To investigate the differentiation potential of CD31- and CD31+ SVF cells towards the endo-

thelial lineage two endothelial differentiation media (EC-2 and EC-3) were applied to both

CD31 fractions separately. Endothelial differentiation was determined by (i) Matrigel assays

and (ii) flow cytometry analyses. 6. Differentiation potential of CD31- and CD31+ SVF cells towards the adipogenic lineage Both cell fractions, CD31- and CD31+ SVF cells, were tested for their differentiation potential

towards adipose tissue. After magnetic-assisted separation SVF cells were cultured in adi-

pogenic differentiation medium. Fat cell differentiation was measured (i) on a molecular level

43

by obtaining the GPDH activity and (ii) by cell counting under light microscopy and Oil Red O

Staining.

44

MATERIAL AND METHODS

TECHNICAL EQUIPMENT

♦ CO2 incubator, 37°C with 5% CO2

and > 95% humidity (Heraeus)

♦ Laminar flow hood (Flow Laborato-

ries GmbH)

♦ Laborwaage/ Laboratory scale (Sar-

torius)

♦ Laborfeinwaage/ Laboratory preci-

sion scale (Sartorius basic)

♦ Centrifuge Minifuge (T-Heraeus)

♦ Laboratory centrifuge (SIGMA)

♦ Centrifuge Biofuge A (Heraeus)

♦ Phasecontrast microscope, Telaval

31 (Zeiss)

♦ Photo device (Nikon D100)

♦ Fluorescence microscope Axioscope

(Leica)

♦ Programme DISCUS (Leica)

♦ Coverplates (Shandon)

♦ Counting cell chamber (Neubauer

improved)

♦ Water shaking bath (GFL

1092/1083)

♦ Shaking device Reax 2000

(Heidolph)

♦ Autoclave (Melag Autoklav 23)

♦ Conical tubes, 10 ml, 50 ml konisch

(Costar)

♦ Pipetting device Pipetboy

(Tecnomara)

♦ Culture flask T-25 (25 cm2/ 5 ml)

(Greiner)

♦ Culture flask T-75 (75 cm2/ 25 ml)

(Greiner)

♦ Culture dish/ plate, 6-well, 12-well,

24-well, 96-well (Falcon)

♦ Petri dish Ø 150 mm (Falcon)

♦ Glass vials 50 ml, 100 ml, 250 ml,

500 ml (Schott Duran)

♦ Pipettes, 1 ml, 2 ml, 5 ml, 10 ml, 25

ml (Greiner)

♦ Eppendorf cups 1.5 ml (Eppendorf)

♦ Disposable syringe Discardit II 5 ml,

10 ml, 20 ml (Becton Dickinson)

♦ Elisa-Reader with HP-Printer (Dynex

Technologies)

♦ Programme Revelation (Dynex

Technologies)

♦ Multi-channel pipette (Eppendorf)

♦ Repeater Pipette and combitips (Ep-

pendorf)

♦ Glass cylinder, Borosilikatglas

(Braun)

♦ Teflon flask (Braun)

♦ Syringe driven filter unit Millex GP

0.22 µm (Millipore)

♦ 70 μm nylon mesh (Verseidag

Techfab GmbH)

♦ 250 µm nylon mesh (Verseidag

Techfab GmbH)

♦ Sucking device Vacusafe (IBS In-

tegra Biosciences)

♦ Pasteurpipettes, 230 mm (Brand)

♦ Tweezers (Aesculap)

♦ Scalpels (Feather)

♦ Scissors (Aesculap)

♦ Clamps (Aesculap)

45

♦ Needles Microlance (Becton Dickin-

son)

♦ Glass slides (Engelbrecht)

♦ Quadriperms (Heraeus)

♦ Magnetic rack Dynal MPC-1 (Dynal

Biotech GmbH)

♦ MiniMACS separation unit (Miltenyi

Biotec GmbH)

♦ MACS Multi Stand (Miltenyi Biotec

GmbH)

♦ MACS columns type MS (Miltenyi

Biotec GmbH)

♦ FACS Calibur apparatus (Beckton

Dickinson)

♦ FACS tubes (Sarstedt)

♦ Programme Cell Quest Pro (Beckton

Dickinson)

♦ Parafilm (Pechiney Plastic Packing)

REAGENTS

♦ Dulbecco´s Modified Eagle Medium

(DMEM)/Ham’s F12 (F12) 1:1 (PAA)

♦ Dulbecco´s Modified Eagle Medium

(DMEM) High Glucose (PAA)

♦ Isocove´s Modified Dulbecco´s Me-

dium (IMDM) with GlutaMAX (Invi-

trogen)

♦ Endothelial Cell Growth Medium with

supplement mix (PromoCell)

♦ Endothelial Growth Medium-2 Mi-

crovascular (EGM-2 MV) with sup-

plement mix (Cambrex Bio Science)

♦ Penicillin (PAA Laboratories)

♦ Streptomycin (PAA Laboratories)

♦ Fetal calf serum (FCS) (Biochrom)

♦ Bovine serum albumin (BSA) (PAA)

♦ BSA for immunhistology (Sigma)

♦ Basic fibroblast growth factor (bFGF)

(Peprotech)

♦ Vascular endothelial growth factor

(VEGF) (Peprotech)

♦ Insulin-like growth factor 1 (IGF-1)

(Peprotech)

♦ Stem cell growth factor beta (SCGF-

β) (Peprotech)

♦ Dexamethasone (Sigma)

♦ Insulin (Roche)

♦ Transferrin (Sigma)

♦ Triiodo-L-thyronine (Sigma)

♦ Isobutylmethylxanthine (IBMX)

(Sigma)

♦ Pioglitazone (Takeda Pharmaceuti-

cals)

♦ Rosiglitazone (Cayman Chemical)

♦ Phosphat buffered saline (PBS2-)

(Biochrom)

♦ Aqua ad injectabile (Delta Select)

♦ Aqua bidest (Pharmacy RWTH

Aachen)

♦ Trypsin 0.02%, 0.05% EDTA in PBS

(PAA Laboratories)

♦ Collagenase CLS Typ I (Biochrom)

♦ Dispase Typ II (Roche)

♦ Fibronectin (Biochrom)

♦ Matrigel™, growth factor reduced

(BD Biosciences)

46

♦ Cell recovery solution (BD Biosci-

ences)

♦ Oil Red O (Merck)

♦ Glycerol-3-phosphat-dehydrogenase

(GPDH) activity assay kit (TaKaRa)

♦ HEPES (Sigma Aldrich Chemie)

♦ NaCl (Merck)

♦ CaCl2 (Merck)

♦ HCl (Merck)

♦ Glucose (Merck)

♦ NH4Cl (Merck)

♦ NaHCO3 (Merck)

♦ KHCO3 (Merck)

♦ β-mercaptoethanol (Fluka)

♦ Paraformaldehyde (PFA) (MERCK)

♦ Acetone (Pharmacy RWTH Aachen)

♦ 4’,6’-Diamidin-2’-phenylindol-

dihydrochlorid (DAPI) (Boehringer

Mannheim)

♦ DAPI working solution (10µl DAPI

stock solution plus 10ml methanol)

♦ Dimethylsulfonoxid (DMSO) (Sigma)

♦ Anti-CD31 paramagnetic Dynabeads

(Dynal Biotech GmbH)

♦ Anti-CD31 MicroBead Kit (Miltenyi

Biotec GmbH)

♦ Mouse anti-human S100 antibody

(DAKO GmbH)

♦ Rabbit anti-human von-Willebrand-

Factor (vWF) antibody (DAKO

GmbH)

♦ Secondary swine anti-rabbit FITC-

antibody, IgG (DAKO GmbH)

♦ Mouse anti-human CD31 antibody

(Dianova GmbH)

♦ Mouse anti-human CD34 antibody

(Dianova GmbH)

♦ Mouse anti-human CD51/61 anti-

body (Dianova GmbH)

♦ Mouse anti-human KDR antibody

(Dianova GmbH)

♦ Secondary goat anti-mouse FITC-

antibody, IgG (Dianova GmbH)

♦ Anti-human CD105 FITC-conjugated

antibody (ImmunoTools GmbH)

♦ Isotype control IgG1 and IgG2 anti-

bodies (ImmunoTools GmbH)

♦ CD31 APC-conjugated antibody

(Miltenyi Biotec GmbH)

♦ DiI acLDL cell tracker (Cell Systems)

♦ Mowiol 4-88 (Calbiochem)

CULTURE MEDIA

Attachment medium/Fibroblast medium

♦ DMEM/Ham’s F12

♦ 10% FCS

♦ 100 U/ml penicillin

♦ 100 µg/ml streptomycin

Table 2.

Endothelial differentiation media

EC-1 EC-2 EC-3

♦ Endothelial Growth

Medium-2 Microvascu-

lar (EGM-2MV)

♦ 10 ng/ml VEGF

♦ 10-6 mol/l rosiglitazone



♦ Endothelial Cell

Growth Medium

(ECGM)

♦ 50 ng/ml VEGF

♦ 20 ng/ml IGF-1

♦ 2% FCS



♦ IMDM

♦ 50 ng/ml VEGF

♦ 100 ng/ml SCGF-β

♦ 30% FCS

♦ 1% BSA

♦ 10-4 mol/l β-

mercaptoethanol



HDMEC medium

♦ EGM-2MV

♦ 15% FCS

♦ 100µg/ml Heparin

♦ 7µg/ml IBMX



HUVEC medium

♦ EGM-2



48

Preadipocyte proliferation medium


♦ 10% FCS

♦ 100 µl bFGF



Table 3.

Adipogenic differentiation media

Adipocyte differentiation medium 1 (d1-5) Adipocyte differentiation medium 2 (≥ d6)


♦ 100 nM dexamethasone

♦ 10 µg/ml transferrin

♦ 66 nM insulin

♦ 1 nM triiodo-L-thyronine

♦ 0.5 mM IBMX

♦ 0.1 µg/ml pioglitazone




♦ 100 nM dexamethasone

♦ 10 µg/ml transferrin

♦ 66 nM insulin

♦ 1 nM triiodo-L-thyronine



Table 4.

Seading of SVF cells

Seeding

density

20.000/cm2

(cell num-

bers)

Seeding

density

30.000/cm2

(cell num-

bers)

Cells at

confluency

(cell num-

bers)

Trypsin/

EDTA (ml)

Growth me-

dium (ml)

6-well 181.400 272.100 1.2 x 106 1 3

T-25 0.5 x 106 0.75 x 106 2.8 x 106 2 5

T-75 1.5 x 106 2.25 x 106 8.4 x 106 5 10-15

49

BUFFERS AND SOLUTIONS

Collagenase buffer

♦ 100 mM HEPES

♦ 120 mM NaCl

♦ 50 mM KCl

♦ 1 mM CaCl2

♦ 50 mM glucose

♦ → pH 7.4

Collagenase solution

♦ 0.494 U/ml collagenase CLS Typ I

♦ 1.5 % BSA

♦ Collagenase buffer

♦ → 1 ml collagenase buffer per 1 g tissue

Erythrocyte lysis buffer

1. Stock solution: For 100 ml Aqua bidest:

♦ 8.02 g NH4Cl

♦ 0.84 g NaHCO3 or KHCO3

♦ 0.37 g EDTA (Di-Na-salt)

♦ 100 ml of this solution plus 900 ml Aqua bidest

♦ → pH 7.4

2. Working solution:

♦ Stock soltution diluted 1:10 with Auqa bidest

MACS buffer

♦ 2 mM EDTA

♦ 0.1% BSA

♦ PBS2-

♦ → degassed by ultrasound

FACS buffer

♦ 2 mM EDTA

♦ 0.2% FCS

♦ PBS2-

50

ISOLATION OF SVF CELLS

SVF cells were isolated from freshly excised human subcutaneous abdominal or mammary

fat tissue at the Department of Plastic Surgery and Hand Surgery - Burn Center from patients

who underwent elective operations (e.g. abdominoplasty, breast reduction, liposuction). In

regards to the amount of human preadipocytes gained from fat tissue, the harvesting method

plays a central role. There are two different surgical isolation methods for fat tissue. Firstly

harvesting of adipose material can be performed by simple excision of adipose tissue during

elective operations such as abdominoplasty, breast reduction and others. The second

method is liposuction in which numerous diminutive fat pieces are isolated. This displays the

most atraumatic version for the patient (Figure 8). Cell yields from liposuction vary enor-

mously in the literature, according to the harvesting procedure, from 1.6 x105 to approxi-

mately 2 x106 cells per gram of lipoaspirated human fat405,434.

Harvesting of excised fat tissue was arranged by isolating the single fat lobules which were

prepared by careful removal of capillaries and connective tissue. After mincing into small

pieces of 2-6 mm in diameter with scissors, adipose tissue was washed with 0.9% NaCl and

digested. Harvesting of adipose tissue by liposuction was performed according to Sydney

Coleman. With the Coleman method, manually applied negative pressure using a 10-cc-

syringe with a blunt tip cannula minces the fat lobules and sucks the small pieces into the

syringe (Figure 8)80. To guarantee cell viability no additional solutions were added. Harvest-

ing sides were predominantly abdomen and breast. Isolation methods were adapted to the

methods of Rodbell330 and Björntorp44,45 with minor modifications. All procedures were per-

formed under sterile conditions under the laminar flow hood. Adipose tissue was either used

directly after harvesting or stored for up to 24 hours at 4°C in DMEM High Glucose. Adipose

Figure 8. Different isolation

methods for fat tissue

Process of adipose tissue harvest

from tissue excision, e.g. abdomin-

oplasty (A.) or lipoasiprates (B.).

A.

B.

51

tissue from aspiration was digested by collagenase treatment after washing with 0.9% NaCl

to remove possible blood contamination.

Digestion of excized and liposuctioned fat tissue was performed with 0.2% collagenase CLS

type I (0.494 U/ml) and 1.5% BSA dissolved in collagenase buffer for 60 minutes at 37°C

under constant shaking. The gained suspension was filtered through a 250 µm filter mesh

and centrifuged at 700g for 7 minutes at room temperature. SVF cells were separated from

mature fat cells by discharging the fat layer on top, containing the lipid-filled adipocytes. SVF

cells located in the cell pellet on the bottom of the tube were resuspended in different culture

media dependent on their further use (Figure 9). For separation of CD31- and CD31+ cells,

the SVF cell pellet was resuspended in EGM-2 MV culture medium in order to enhance EC

numbers and maintain EC characteristics of CD31+ cells in the SVF. For plain flow cytometry

analyses and endothelial differentiation assays of SVF cells, cells were seeded in

DMEM/F12 (1:1) with 10% FCS. Seeding was performed on fibronectin-coated culture flasks

at a density of 30,000 cells/cm2. Cells were washed twice with 0.9% NaCl after 24 and 48

hours to remove contaminating blood cells.

52

adipose SVFin culture

fat tissue fat lobules

digestion

filtration

centrifugation

separation

resuspention

seeding

SVF pellet

CD31 DYNABEAD SEPARATION OF SVF CELLS

To split CD31- from CD31+ cells two widely accepted methods for cell separation were ap-

plied, cell splitting with Dynabeads and with Microbeads. For endothelial cell separation the

most frequently used surface protein is CD31 as it is expressed on all endothelial cells but

also on platelets. To minimize platelet contamination cells were washed thoroughly with NaCl

after harvesting and again with PBS2- before trypsinizing for Dynabead/Microbead separa-

tion. Cells were grown to subconfluence for three to five days, trypsinized after washing, re-

Figure 9. SVF cell isolation from

excised tissue

Excised adipose tissue is minced with

scissors into single fat lobules and digested

with a collagenase solution. The resulting

suspension is then filtered through a nylon

mesh to remove undigested components,

specially fibrous tissue and small vessels.

Subsequent centrifugation deposits the

stromal vascular fraction (SVF) as a pellet

at the bottom of the tube from the floating

lipid layer (mature adipocytes) on top. The

supernatant is removed by pipetting and

followed by resuspension of the cell pellet

which allows seeding of the SVF cells in a

culture dish.

53

suspended in 10 ml cold PBS2-/0.1% BSA with 2 mM EDTA (MACS buffer) and washed by

pipetting the cell suspension carefully. Cells were centrifuged at 400g for 5 minutes, resus-

pended in 8 ml cold MACS buffer and passed trough a 70 µm nylon mesh into a 15 ml tube.

The mesh ensured that only single cells passed through by excluding cell clusters thereby

maximizing the purity and percentage of CD31+ cells. Afterwards cells were counted with a

Neubauer chamber to calculate cell numbers. To ensure optimal binding of paramagnetic

beads cell concentration should not exceed 108 cells/ml sample volume. Before use Dyna-

beads were prepared by washing with cold MACS buffer. Dynabeads were suspended in 8

ml cold MACS buffer in a 15 ml tube and pipetted thoroughly. Next the tube was placed into

the magnetic rack for two minutes until the solution became clear and all beads had moved

to the magnet. The see-through supernatant was discharged, the tube removed from the

rack and the beads were resuspended anew in 8 ml cold MACS buffer by pipetting. This

washing step was performed three times and Dynabeads were resuspended in the original

volume again. To separate CD31+ cells from CD31- cells by anti-CD31 paramagnetic beads,

cells were incubated with 200 µl washed CD31 Dynabeads for endothelial cells (25 µl

beads/ml cell suspension) at 4°C for 30 minutes under constant rotation, allowing for binding

of endothelial cells to the beads. Afterwards the bead-cell suspension was pipetted a few

times to release cell clusters which may have formed during the incubation period. The tube

containing the Dynabead-labeled cells was placed in the magnetic rack (Dynal MPC-1) for 2

minutes until all beads were attached to the magnet (Figure 10). The supernatant with CD31-

cells was transferred to a fresh tube by pipetting, centrifuged at 400g for 5 minutes, washed

once with MACS buffer, and cells were subsequently seeded on fresh culture flasks. For iso-

lating CD31+ cells, three washing steps as described before were performed with Dynabead-

labelled cells. After the falcon tube was removed from the magnetic rack, CD31+ SVF cells

with Dynabeads attached to the surface were resuspended in fresh DMEM/F12.

54

For attachment to culture flasks both cell types were cultivated in DMEM/F12 supplemented

with 10% FCS in fibronectin-coated flasks over night at a density of 80.000 cells/cm2. Seed-

ing at high density was performed in order to keep proliferation times as low as possible until

complete confluence was achieved, necessary for further differentiation experiments towards

the adipogenic and endothelial lineage.

CD31 MICROBEAD SEPARATION OF SVF CELLS

Cells were grown to subconfluence, trypsinized after washing twice with PBS2-, resuspended

in 10 ml MACS buffer and passed trough a 70 µm nylon mesh to remove cell clusters. Cells

were resuspended in 60 µl cold MACS buffer to a maximum concentration of 1x 107 cells/

60 µl buffer. 20 µl FcR Blocking reagent was added to minimise unspecific binding of Micro-

Beads to FcR domains. The gained suspension was vortexed thoroughly and 20 µl of CD31

MicroBeads were added. Cells were incubated for 15 minutes at 4 °C to allow binding of

beads to the cell surface. During the incubation period MACS columns were prepared by

placing two columns in the magnetic fields of each MiniMACS separator. The columns were

rinsed with 1 ml MACS buffer three times. After the incubation time non-attaching beads

were removed from the suspension cells by washing once with 5 ml medium and subsequent

centrifugation at 400g for 5 minutes. The supernatant was discharged, cells were resus-

pended in 500 µl MACS buffer (up to 108 cells) and transferred onto prepared MACS col-

B.

C.

A. Figure 10. Splitting of SVF cells with

paramagnetic CD31 Dynabeads

Dynabeads are uniform, superparamagnetic

polystyrene beads with a diameter of about

4.5 µm. The beads are coated with a mouse

IgG1 monoclonal antibody specific for the

CD31 cell surface antigen PECAM-1. Re-

quired material for Dynabead separation

includes the Dynal magnet, CD31 Dyna-

beads and a conical tube (A.). Washed Dyna-

beads are mixed with the cell sample and the

beads ligate to the CD31+ target cells (red)

after incubation (B.). After binding the posi-

tive cells are separated from the negative cells

by a magnet, whereas the CD31+ cells attach

to the magnet and the CD31- cells remain in

the cell suspension (C.).

55

umns. CD31- cells were collected after passing through the column without attachment in a

15 ml tube and columns were washed three times with 1 ml MACS buffer, each time once

the column reservoir is empty. The total effluent was collected and the gained CD31- cell

suspension was centrifuged at 400g for 5 minutes, washed once with 10 ml medium and

seeded onto fresh culture flasks. CD31+ SVF cells, which attached to the columns, were de-

tached by removing the columns from the magnetic filed first and placing the columns in two

fresh 15 ml tubes. Cells were immediately flushed out by adding 1 ml of medium to the col-

umns and firmly applying the plunger supplied together with the columns (Figure 11).

Cells were washed once with 10 ml medium and resuspended in fresh DMEM/F12 (1:1) after

centrifugation. Both CD31+ and CD31- cells were cultivated in DMEM/F12 (1:1) supple-

mented with 10% FCS at a density of 80.000 cells/cm2 over night for adherence to culture

dishes. Seeding was performed at high density to keep proliferation times until complete con-

fluence was achieved as low as possible, necessary for further differentiation experiments

towards the adipogenic and endothelial lineage.

B.

C. D.

A.Figure 11. Splitting of SVF cells with para-

magnetic CD31 MicroBeads

MACS MicroBeads are superparamagnetic

particles composed of a biodegradable matrix

and have a diameter of about 50 nm. For direct

magnetic cell labelling the cell suspension is

incubated with CD31 MicroBeads giving

enough time for the beads to bind to the CD31+

cells (red) (B.). Subsequently the cell suspen-

sion is transferred onto a MACS MS Column

placed in a MiniMACS Separator (A.). The

separator is a strong permanent magnet and

generates a high-gradient magnetic field on the

column matrix. MicroBead-labelled cells are

retained in the magnetic field whereas unla-

belled cells pass through (C.). They are collected

in a conical tube as the CD31- cell fraction.

After the column is removed from the magnet

the retained CD31+ cells are eluted by pushing

the plunger down and flushing out the posi-

tively selected cells (D.).

56

CULTURING OF SVF CELLS

Culture media were changed for both CD31- and CD31+ to either endothelial or adipogenic

differentiation medium 12 hours after Dynabead or MicroBead separation, respectively (see

Figure 12).

To evaluate differentiation plasticity towards endothelium, three types of differentiation media

were tested (Table 2). The first culture medium (EC-1) used was an endothelial differentiat-

zion medium for angiogenic progenitor cells according to Wang et al.409 and contained Endo-

thelial Growth Medium-2 Microvascular (EGM-2MV) with 10 ng/ml VEGF and 10-6 mol/l

rosiglitazone. The second medium (EC-2), an endothelial differentiation medium, was Endo-

thelial Cell Growth Medium (ECGM) supplemented with 50 ng/ml VEGF, 20 ng/ml IGF-1, and

2% FCS, according to Miranville et al.260 with minor modifications. The third culture medium

(EC-3) tested was a hematopoietic and endothelial progenitor cell medium for peripheral

blood progenitor cells according to Gehling et al. 125, containing IMDM, 50 ng/ml VEGF, 100

ng/ml SCGF-ß, 30% FCS, 1% BSA, and 10-4 mol/l ß-mercaptoethanol.

To perform adipogenic differentiation, cells were cultured in DMEM/F12 (1:1) supplemented

with bFGF (10 ng/ml) and 10% FCS (referred to as proliferation medium) to obtain cell con-

fluency again. Lipid droplet development was then promoted for 21 days by changing me-

dium to DMEM/F12 (1:1) without serum addition, supplemented with 66 nM insulin, 100 nM

dexamethasone, 0.5 mM IBMX, 0.1 µg/ml pioglitazone, 1 nM triiodo-L-thyronine, and 10

µg/ml human transferrin (adipocyte differentiation medium 1, Table 3). After 5 days of incuba-

tion, medium was used as before but without IBMX and pioglitazone for 16 more days (adi-

pocyte differentiation medium 2, Table 3). Culture media were changed every three days.

57

10% 90%CD31 Dynabead/ Microbead

splitting

SVF culture in EC medium

CD31- SVFCD31+ SVF

adipose differentiation► GPDH assay

capillary structures► Matrigel

Extracellular Matrix

CD31- SVFCD31+ SVF

CELL SPLITTING

Medium was removed from the culture flasks and cells were washed twice with PBS2- to

eliminate residual medium. According to the size of the culture flask trypsin/EDTA was added

and cells were incubated at 37°C for 2-3 minutes. Serum-containing medium was supple-

mented to the trypsin/EDTA containing the detached cells in order to stop the enzymatic

process. Cells were centrifuged at 1350 rpm for 5 minutes, resuspended in 5 to 10 ml fresh

culture medium and counted with a Neubauer cell counting chamber under the phasecon-

trast microscope. Cell numbers were calculated as follows: Counted cells / 4 x 104 = cell

numbers / ml. Subsequently cells were seeded at the desired density and medium was

changed the following day to remove non-vital cells.

Figure 12. Experimental design of

Dynabead/Microbead splitting of

SVF cell culture and separate

differentiation of subpopulations

towards adipogenic and andothe-

lial lineage

SVF cells were cultured in EC me-

dium to enhance EC numbers prior to

cell separation. After splitting with

paramagnetic beads CD31- and

CD31+ SVF cells were both cultured

in endothelial and adipogenic differ-

entiation medium. Endothelial differ-

entiation was measured via capillary

structure formation on Matrigel

ECM and flow cytometry analysis.

Adipogenic conversion was deter-

mined by GPDH activity and Oil Red

O staining.

58

FIBRONECTIN AND MATRIGELTM COATING Fibronectin (1 mg) was diluted to a concentration of 0.02 mg/ml by adding 50 ml aqua ad

injectabile (fibronectin working solution). The bottom of the culture flask was covered with

fibronectin working solution and air-dried under sterile conditions.

MatrigelTM Basement Membrane Extracellular Matrix is made out of Engelbreth-Holm-Swarm

(EHS) mouse tumor. Major components of Matrigel include laminin, collagen IV, heparan

sulfate proteoglycanes, entactin and nidogen212. Further it contains TGF-β, FGF, tissue

plasminogen activator254 and other growth factors which naturally exist in the EHS tumor.

Matrigel is known to support and provide several factors necessary for the study of angio-

genesis283,284. For testing the angiogenic potential of SVF cells in vivo the thin gel method

was used. Culture flasks are coated with Matrigel in a thickness of about 0.5 mm and cells

are plated on top. Matrigel was diluted 1:2 with serum-free medium (EGM2-MV without sup-

plement mix), placed in aliquots and stored at -20°C. To handle Matrigel exclusively pre-

cooled material should be used as Matrigel gels at temperatures over 5-10°C. One day prior

to use Matrigel was thawn on ice at 4°C overnight and pipettes and culture plates were

placed in the freezer at -20°C for precooling. Next day a bucket was filled with ice and Ma-

trigel, pipettes and culture plates were placed on ice to keep them cold. All experiments with

Matrigel coating were perfomed with 6-well plates. About 500 µl of Matrigel were added per

well of a 6-well plate (50µl/cm2) and culture plate was placed for one hour at 37°C to allow

Matrigel to harden. Afterwards cells were seeded at a density of 30,000/cm2 (see Table 4) in

EC medium to examine EC differentiation and capillary formation capacity.

HUMAN DERMAL MICROVASCULAR ENDOTHELIAL CELL (HDMEC) ISOLATION

HDMECs were isolated from freshly excised human skin at the Department of Plastic Sur-

gery and Hand Surgery - Burn Center from patients who underwent elective operations (e.g.

abdominoplasty, breast reduction, flap surgery). The unprocessed skin was placed on a ster-

ile Petri dish with the epidermal, smooth side facing upward, and was cut into small pieces of

5 x 5 mm2 with a scalpel and digested overnight with dispase II at 4°C. Afterwards the skin-

dispase mixture was incubated under constant shaking at 37°C for 30 minutes. Dispase II

enables the separation and removal of the epidermis from the dermis. With sterile tweezers

skin pieces were transferred in a fresh tube with 20 ml cold PBS2- to stop the dispase diges-

tion. After transferring onto a petri dish the epidermis was separated from the dermis by care-

fully peeling back the epidermal layer using sterile tweezers. The dermis was minced with

scissors into small pieces in a 50 ml tube. The dermal layer was then digested in prewarmed

0.2% collagenase CLS type I with 1.5% BSA in collagenase buffer (sterile filtered) for 90

minutes at 37°C under constant shaking. Cell suspension was filtered through a 250 µm

59

mesh to remove contaminating fibrous tissue and afterwards culture medium was added.

After centrifugation at 400 g for 10 minutes the cell pellet was resuspended in HDMEC me-

dium and cells were seeded on fibronectine-coated culture dishes. Cells were washed twice

after 24 hours with 0.9% NaCl to remove possible blood contamination and cultured to sub-

confluency. At subconfluency cells were washed twice with PBS2- and incubated with Tryp-

sin/EDTA for two to three minutes and stopped with medium containing 10% FCS. Detached

cells were counted with a Neubauer counting chamber and separated with CD31-Dynabeads

and CD31-Microbeads as described earlier. Afterwards the CD31 positive HDMECs were

seeded on fibronectin-coated chamber slides and 6-well plates in HDMEC medium. They

were used as positive control for endothelial differentiation assays on MatrigelTM Basement

Membrane Extracellular Matrix as well as flow cytometry analysis.

HUMAN DERMAL FIBROBLAST ISOLATION

Fibroblasts were isolated following the same procedure as with HDMECs from human skin.

After separation with CD31 Dynabeads/Microbeads, the CD31 negative fibroblasts were

seeded on chamber slides, 6-well plates, and T75 culture flasks in fibroblast medium as

negative control.

HUMAN UMBILICAL VEIN ENDOTHELIAL CELL ISOLATION

HUVECs were isolated from human umbilical veins that had been received as a gift from the

Department of Pathology. The injured ends of each vein were removed and a feeding needle

was carefully inserted into the vein and fixed with a clamp. Afterwards the vein was rinsed

with 20 ml HEPES/6 mM EDTA via a syringe to remove possible blood contamination. An-

other feeding needle was inserted at the other end of the vein, fixed and the flow through

was checked with another 20 ml of HEPES/6 mM EDTA. A second syringe was filled with 5

ml trypsin/EDTA (37 °C warm) to replace the first one. Trypsin/EDTA was injected into the

umbilical vein and the cord was inserted in U-shape into a 500 ml vial filled with 37°C pre-

warmed PBS2-. After incubation for 20 minutes at 37°C in the water bath, the cord was

placed on a sterile operation cloth and massaged for five minutes to release HUVECs from

the vessel wall. 8 ml HEPES/6 mM EDTA was filled into another syringe and the syringe was

connected to the feeding needle. The cord vein was flushed thoroughly and the eluted fluid

was collected in a 50 ml tube containing 2 ml FCS to stop the digestion. After centrifugation

at 400g for 5 minutes at room temperature cells were resuspended in HUVEC medium and

subsequently seeded on fibronectin-coated T75 culture flasks. They were used as positive

control for endothelial differentiation assays on MatrigelTM Extracellular Matrix as well as flow

cytometry analysis.

60

FLOW CYTOMETRY ANALYSIS

To perform flow cytometry analyses, freshly isolated cells were incubated for 15 minutes with

erythrocyte lysis buffer at 4°C. For staining with DiI, cells were washed with PBS2-, and incu-

bated with DiI-ac-LDL (acLDL) (10 µg/ml) for one hour at 37°C. Afterwards cells were

washed twice with plain medium, once with PBS2-, and fixed with 4% paraformaldehyde

(PFA). Next, cells were centrifuged at 400g for 10 minutes and resuspended in PBS2- con-

taining 0.2% FCS and 2 mM EDTA. For incubation with primary anti-human CD31, CD34,

KDR, CD51/61, vWF, S100, and CD105-FITC, cells were also fixed with 4% PFA, resus-

pended in PBS containing 0.2% FCS and 2 mM EDTA, and kept at 4°C for 30 minutes.

Washing and incubation with secondary FITC-conjugated antibodies was performed for 30

minutes at 4°C with all samples except for the already FITC-labelled CD105. For additional

staining with CD31 APC-conjugated antibodies to generate double-labelling, cells were incu-

bated with CD31-APC for 10 minutes at 4 °C, washed and resuspended in PBS containing

0.2% FCS and 2 mM EDTA. Labelled cells were analyzed by flow cytometry using a FACS

Calibur apparatus (Beckton Dickinson) and CellQuest Pro analysis (Figure 14).

FL 3

SSC

FSC

FL 1

FL 2

FL 4 APC

PE

FITC

488nm

633nm

90° light scattergranularity

2°-16° light scattersize

ArgonLaser

Side scatter(SSC)

Forward scatter(FSC)

Light source

A.

B.

Figure 13. Flow cytometry

analysis

Flow cytometry analysis allows

simultaneous measuring of relative

cell size (FSC), cell granularity and

internal complexity (SSC) and

fluorescence intensity (A.). The

major components comprise a fluid

system, an optical system with

laser, reflector and filter device.

According to the fluorescence

staining applied wavelengths differ

between 450 nm and 700 nm. FITC

requires a laser wavelength of about

488 nm in the green range, con-

trariwise APC has its optimum at

633 nm in the red spectrum (B.).

61

Isotype controls for IgG1 and IgG2a were used as recommended by the supplier to exclude

non-specific binding of antibodies and were always negative. Each labelling with FITC-CD

markers was compared to control cells exclusively stained with the secondary FITC-

conjugated fluorescence antibody. The percentage of positively stained cells was determined

after correction for the percentage of cells reacting with the secondary FITC antibody. Stain-

ing of CD31+ as well as CD31- SVF cells was performed on the day of adipogenic and endo-

thelial differentiation induction.

IMMUNOSTAINING WITH VON-WILLEBRAND-FACTOR

For immunostaining with vWF cells were seeded on glass slides in chamber slides culture

dishes. Preceeding the staining procedure culture medium was removed from chamber

slides containing adherent cells and glass slides were washed twice with PBS2-. Cells were

fixed in cold acetone for 20 minutes at 4°C, acetone was removed again and glass slides

were dried at room temperature for a few minutes until completely parched. Glass slides

were placed into PBS2- and positioned into coverplates. Coverplates were rinsed once with

fresh PBS2- and each incubated with 100 µl primary anti-vWF antibodies for one hour at room

temperature. Antibodies against vWF were diluted 1:200 with PBS2-/1% BSA prior to use.

After washing three times with PBS2-, slides were stained with 100 µl FITC-conjugated sec-

ondary antibodies each for one hour at 37°C. FITC antibodies were diluted 1:30 with PBS/1%

BSA before use as well. After washing three times with PBS2-, staining of the cell nucleus

was performed by applying 100 µl DAPI solution for each coverplate and incubating for 15

minutes at room temperature. After washing three times with PBS2- glass slides were re-

moved from coverplates, coated with two drops of Mowiol 4-88 and covered with small glass

slides. Mowiol was hardened for a few hours at 4°C in the dark. Detection of vWF expression

was performed by fluorescence microscopy. vWF staining of CD31+ and CD31- SVF cells

was performed on the same day as adipogenic and endothelial differentiation induction to

ensure purity of cells used for differentiation assays.

DETERMINATION OF GLYCEROL-3-PHOSPHATE DEHYDROGENASE (GPDH) ACTIV-ITY

To evaluate preadipocyte differentiation on a molecular level, the activity of GPDH was

measured with a kit system from Takara. GPDH is a representative molecular key marker of

adipogenic conversion335, which has been used for decades to measure preadipocyte differ-

entiation298,368.

62

NADH/H+ NAD+

GPDH

cytosol

Dihydroxyaceton-phosphat

Alfa-Glycero-phosphat

GPOXFADFADH2

IMM

Qox Qred

To perform GPDH assays, culture medium was removed and cells were washed twice with

PBS2-. Cell lysis was induced by adding 1 ml “Enzyme Extraction Buffer” per T25 culture

flask. Culture flasks were incubated for three to five minutes at 37°C until cell lysis was com-

pleted. Afterwards the lysate was transferred to a 15 ml tube and diluted to two-fold by add-

ing 1 ml “Dilution Buffer”. To gain the desired four-fold dilution, 250 µl of the cell suspension

were added to another 250 µl “Dilution Buffer” in a fresh 1.5 ml Eppendorf cup. The samples

are stable for one hour on ice if necessary. 25 µl of each four-fold diluted sample was trans-

ferred into each 96-well. GPDH activity was determined by measuring absorbance at 340 nm

with an ELISA reader. The 96-well plate was inserted into the reader and 100 µl “Substrate

Solution” was pipetted to each sample via multi-channel pipette. As the reaction starts imme-

diately after mixing the cell suspension with “Substrate Solution”, it was substantial to add the

“Substrate Solution” quickly to each 96-well. This was achieved by the application of a multi-

channel pipette. Next, the ELISA reader was started at once to measure the GPDH activity.

The enzyme activity was measured ten times during the redox reaction. GPDH activity was

calculated with the following formula supplied by Takara:

GPDH activity (units / ml) = ΔOD340 x 2.06 x dilution ratio of the sample

coenzyme for the reversible conversion between both substrates and can be measured by the GPDH assay through the secondary

decrease of NADH. GPDH activity is calculated by measuring the reduction of NADH via declining absorbence at 340nm.

Dihydroxyacetone phosphate + NADH + H+ → Glycerol-3-phosphate + NAD+

Figure 14. Enzymatic reactions in glycerol-3-

phosphate dehydrogenase (GPDH) assay

Sequential binding of the fatty acids activated as

acyl-CoAs to glycerol-3-phosphate is a major step in

lipid biosythesis. In the liver glycerol-3-phosphate is

directly synthesised from glycerol with the help of

the glycerol kinase. Due to the absence of the glycerol

kinase in adipose tissue und muscle, glycerol-3-

phosphate is produced by the reduction of dihy-

droxyacetone phosphate in combination with

NADH. As dihydroxyacetone phosphate is an inter-

mediate product in the glycolytic pathway, lipid

synthesis in adipose tissue and muscle cannot pro-

ceed without glucose. The enzyme which catalyses

the conversion between dihydroxyacetone phosphate

and glycerol-3-phosphate is the glycerol-3-phosphate

dehydrogenase (GPDH). GPDH requires NAD as a

63

OIL RED O STAINING

Oil Red O staining is an established staining method for intracellular lipid vacuoles. This

method stains lipid droplets with a red colour so that even small droplets can be identified in

the cytoplasma and the percentage of lipid-containing cells can be validated. Besides lipid

droplets intracellular stress vacuoles are stained slightly red as well when incubated with Oil

Red O. The staining intensity of stress vacuoles is minor to that of lipids and these vacuoles

are extremely small in diameter so that they can be distinguished from lipid droplets in light

microscopy examination.

Oil Red O dye (0.5 g) was diluted in 100 ml of 99% isopropanol. This stock solution was

mixed 6:4 with aqua bidest and stored for 24h at room temperature. Afterwards the solution

was filtered to remove small dye particles. Monolayer cultures were washed twice with PBS2-

and fixed with 10% cold PFA and incubated for five hours at 4°C. After removing the PFA Oil

Red O working solution was added to culture flasks and samples were kept at room tempera-

ture for at least two hours. After washing twice with PBS2- the stained cells were kept in 10%

PFA and examined by light microscopy to count lipid-filled adipocytes.

CELL COUNTING

To dertermine (i) the percentage of differentiated to undifferentiated cells, (ii) to detect those

cells attached to Dynabeads as well as (iii) cells stained for vWF under the fluorescence mi-

croscopy, cell counting was performed under the phasecontrast or fluorescence microscope

with 100x magnification.

Cells were counted in six randomly chosen fields (six fields per culture flask). Differentiated

and undifferentiated cells were counted to determine the percentage of both cell types and

compare the differentiation level between different wells. The same was done with cells car-

rying Dynabeads on their surface and vWF positive cells appearing green under fluores-

cence light.

STATISTICAL EVALUATION

Data of expression of surface markers and GPDH activity were expressed as mean value ±

standard deviation. The significance of differences between CD31+ and CD31- cell differen-

tiation was evaluated using the t-test. Differences with p<0.05 were considered significant

and p<0.01 highly significant.

64

RESULTS

COMPARISON OF EXCISED FAT TISSUE VERSUS LIPOSUCTION MATERIAL

The surface expression of freshly isolated SVF cells from liposuction and excision was char-

acterized by flow cytometry without performing any previous purification. SVF cells harvested

from excised fat tissue show strong labelling for CD34 and a less intensive staining for KDR

(Figure 15 A., Table 5). These cells also comprise a small population of 5-10% of CD31 and

CD51/61 positive cells. SVF cells derived from liposuction aspirate (Figure 15 B.) are positive

for CD34, similar to cells from excised tissue (compare Table 5). CD31 and CD51/61 fluores-

cent labeling was stronger in liposuctioned SVF cells with up to 40% CD31 positive cells

(p=0.05 compared to CD31 expression in excised tissue). KDR was much more positive in

lipoaspirate than in excised fat (p<0.01).

Following standard culture conditions for two weeks in DMEM/F12 containing 10% FCS, cells

from lipoaspirate remained positive for CD34 in the same percentage as freshly isolated cells

while CD34 expression in excised fat decreased. Endothelial cell marker CD31 decreased in

liposuction and excised material by about 50%, CD51/61 shows a similar tendency (Table 5).

During cell cultivation, KDR expression or selection, respectively, increased in excised tissue

cells but decreased in liposuction material.

Fibroblasts, HDMECs and HUVECs were used as controls. Fibroblasts were negative for

CD31, vWF, CD51/61, KDR, S100, acLDL, positive for CD105, and slightly positive for

CD34. HDMECs showed positive results for CD31, vWF, acLDL, CD34, and KDR, as well as

slightly positive staining for CD51/61. HUVECs were positive for CD31, acLDL, CD34, KDR,

and negative for CD51/61 (Figure 21). Isotype controls for IgG1 and IgG2a were always

negative (Figure 15).

65

CD51

A. CD31 CD34

KDR

CD31

CD51

CD34

KDR

B.

FITC control IgG1 IgG2a

FITC control IgG1 IgG2a

Figure 15. Characterisation of SVF cells from excised versus liposuctioned adipose tissue

SVF cells were isolated from excised (A.) and liposuctioned (B.) adipose tissue by collagenase digestion and immedi-

ately analyzed by flow cytometry without any previous purification or culturing. Cells were screened for the expres-

sion of CD31, CD34, CD51/61, and KDR. IgG1 and IgG2a were used as isotype controls. Also, staining with secon-

dary FITC-conjugated fluorescence antibodies (FITC control) only was monitored.

66

Table 5. Expression of surface markers on adipose tissue derived SVF cells

SVF cells were isolated from excised adipose tissue or liposcution material, respectively, as described in Figure 15, and cul-

tured in DMEM/F12 plus 10% FCS for 14 days, followed by flow cytometry analysis. Given is the expression of the surface

markers CD31, CD34, CD51/61, and KDR in percentage. Fibroblasts were used as negative, HDMECs and HUVECs as

positive control. Data are derived from three individual experiments.

excision

day 0

excision

day 14

liposuction

day 0

liposuction

day 14

CD31+ 10% ± 2% 6% ± 3% 24% ± 15% 12% ± 5%

CD34+ 57% ± 3% 36% ± 10% 57% ± 20% 57% ± 4%

CD51/61+ 4% ± 1% 5% ± 2% 11% ± 8% 7% ± 3%

KDR+ 14% ± 5 52% ± 8 43% ± 23% 23% ± 7%

67

CHARACTERISATION OF SURFACE MARKERS AND MORPHOLOGICAL CHANGES IN SVF CELLS IN CULTURE

The developement and changes in surface marker expression under standard culture condi-

tions were measured after one and two weeks (Figure 16). Preadipocytes loose their differ-

entiation capacity when cultured in standard medium for several passages. Here we focused

additionally on stem cell markers (CD34, KDR and S100) to evaluate the differentiation po-

tential after long term culture (compare also Table 5).

Unpurified SVF cells from liposuctioned and excised fat tissue were cultured in DMEM/F12

plus 10% FCS for either one or two weeks and flow cytometry was performed. After one

week CD34, KDR and S100 were still fully expressed by SVF cells (Figure 16 A.), after two

weeks all three stem cells markers decreased (Figure 16 B., Table 5).

A.

B.

CD34 KDR

CD34 KDR

S100

S100

Figure 16. Flow cytometry analysis of preadipocytes during culture

SVF cells without prior purification were cultured in DMEM/HAM´s F12 with 10% FCS. Changes in surface

protein expression during proliferation were measured by flow cytometry for CD34, KDR and S100 on day 7 (A.)

and day 14 (B.).

68

IN VITRO DIFFERENTIATION OF UNPURIFIED SVF CELLS UNDER ENDOTHELIAL ME-DIUM CONDITIONS

Freshly isolated preadipocytes responded significantly different to the cultivation in the vari-

ous EC media (Figure 17). SVF cells were tested for their EC development on plain plastic

culture dishes and were stained with vWF and DAPI. In medium 1 (EC-1) only minor prolif-

eration and no formation of capillary structures was detected in vWF positive cells (Figure 17

A.-D.). Medium 2 (EC-2), in contrast, induced substantial proliferation and enhanced tubular

network formation of vWF positive cells (Figure 17 E.-H.). Further positive stained cells

showed capillary structures with widespread network connections (Figure 17 F.). Medium 3

(EC-3) also allowed significant cell proliferation but no network formation (Figure 17 I.-M.).

Standard proliferation medium for preadipocytes showed equal results in proliferation and

tubular structure formation as medium EC-1 (data not shown).

69

EC medium 1 EC medium 2 EC medium 3

A.

B.

C.

D.

E.

F.

G.

H.

I.

K.

L.

M.

Figure 17. Preadipocytes in different EC media without prior purification

Adipose progenitor cells were screened for EC development on plastic culture dishes by immunofluorescence staining with

vWF and DAPI. All pictures are vWF images except for D., H., M. (vWF-DAPI merged), A.-D. Medium 1, E.-H. Medium

2, I.-M. Medium 3. Magnification in A., E., I. is x20, in B., F., K. is x50, in C., G., L., D., H. and M. is x100.

70

SPLITTING AND PURITY ANALYSIS OF CD31- AND CD31+ SVF CELLS

To further characterize the unpurified SVF population, von-Willebrand factor immunofluores-

cence staining was performed 12h after cell isolation and culture in DMEM/F12, 10% FCS.

Freshly harvested SVF cells from excised fat tissue contain a small amount of approximately

10% of vWF positive (Figure 18 A.) and CD31 positive cells, as revealed by microscopical

counting and flow cytometry analyses (compare Table 5). Cultured under endothelial differ-

entiation conditions (EC-2) in the presence of VEGF and IGF-1, these SVF cells develop

tubular structures on plastic without Matrigel™. This was demonstrated by vWF-

immunofluorescence staining after culturing for two weeks (Figure 18 B.). Following separa-

tion into CD31- and CD31+ cells by Dynabeads or MicroBeads, respectively, CD31- SVF cells

show no expression of vWF or CD31 (Figure 18 C., E.), neither after one nor after two weeks

culturing in proliferation medium. Furthermore, CD31- cells do not reveal endothelial network

formation on plastic culture dishes (Figure 18 C.). Analyzing expression of S100, an early

marker of adipogenesis, these cells are found almost 100% positive in flow cytometry (Figure

18 D.-E.). CD31+ cells, in contrast, demonstrate strong vWF expression (97% ± 2% stained

cells) and CD105 staining after one week. The Dynabead isolation procedure demonstrates

to be effective since 95% ± 5% of the cells carry Dynabeads, confirming that they are posi-

tive for CD31 (Figure 18 F.-I., Figure 18 E.). Cells isolated with CD31 MicroBeads show

strong staining for CD31 in flow cytometry (Figure 18 L., N., P.). Furthermore CD31+ Mi-

croBead-isolated cells reveal the same surface expression characteristics for CD105 and

vWF as after isolation with Dynabeads. During proliferation, magnetic beads initially remain

bound to the cells and resist washing steps, however, cell proliferation reduces bead num-

bers continuously.

71

S100

E.

CD105

vWF

CD31 Dynabead/ MicroBead splitting

CD31- SVF

freshlyisolatedSVF cells(excisedfat tissue)

CD31+ SVF

cultured in EC mediumwithoutCD31 splitting

A. B.

C.

F.

90% 10%

D.

G.

H.

I.

CD31+ SVF

CD31+ SVF

CD31+ SVF

L.

M.

S100

CD

31

CD105

CD

31C

D31

vWF

CD34

K.

CD

31N.

CD34

P.

O.

Figure 18. Scheme of the application of CD31 Dynabead separation for SVF cells

Freshly isolated SVF cells from excised tissue were analyzed by fluorescence microscopy after isolation and transferred to

tissue culture dishes (A.). Cells were then either cultured in EC medium (EC-2) without Dynabead treatment and analyzed

again after two weeks (B.) or separated by CD31 Dynabeads or MicroBeads into a CD31- (C.-E.) and a CD31+ fraction (F.-

P.). To confirm the purity of the isolation procedure, flow cytometry was performed with CD31- cells double-stained for

72

CD31 and S100 expression (D. and E.) and with CD31+ cells double-labelled for CD31 in combination with CD105, vWF,

or CD34 (K.-P.). For fluorescence microscopy, cells were stained with vWF and DAPI. All pictures except for F. (vWF alone)

and I. (bright field and vWF, merged) are vWF-DAPI merged images. F.-I., K., M. and O.: Dynabead cell sorting, L., N. and

P.: MicroBead cell sorting. Arrows in H. and I. indicate Dynabeads attached to cell surfaces. Magnification in A., B., C., F.,

G. and I. is x200, in H. x400.

DIFFERENTIATION POTENTIAL OF CD31- AND CD31+ SVF CELLS TOWARDS EC LINEAGE

To evaluate endothelial differentiation potential of CD31- SVF cells morphologically, this cell

fraction was seeded onto Growth Factor Reduced Matrigel™ in EC-2 or EC-3 culture medium,

respectively, and tube formation was assayed. Cells form three-dimensional tube-like struc-

tures in both media (Figure 19 A.-D.), while EC-2 medium also allows formation of extensive

intercellular tube networks (Figure 19 C.-D.). CD31+ SVF cells showed equal results com-

pared to CD31- SVF cells when seeded on Matrigel™ ECM (data not shown). HUVECs, which

were used as a positive control, also formed networks but were not able to grow three-

dimensionally (Figure 19 E.-F., compare to Figure 19 C.).

Next, endothelial differentiation of CD31- cells grown in EC-2 or EC-3, respectively, was ana-

lyzed by flow cytometry to determine the quantitative expression of the surface markers

CD34, KDR, and acLDL (compare Table 6). In EC-3 medium CD31- SVF cells express CD34

by up to 46% and KDR up to 90%, demonstrating the capacity of this medium to enhance

stem cell characteristics of SVF cells (Figure 20 A.). In EC-2 medium CD34 and KDR ex-

pression is significantly lower (p=0.003 for CD34, p=0.006 for KDR) (Table 6, Figure 20 B.).

In both media, cells were positively stained for acLDL confirming differentiation towards an

endothelial phenotype (Figure 20).

HDMECs and HUVECs were used as positive (Figure 21), fibroblasts as negative control.

73

A. B.

C. D.

E. F.

►

►

►

►

200µm

200µm

200µm

200µm

200µm

200µm

►

►

►

Figure 19. Morphological analysis of endothelial differentiation of CD31- SVF cells and HUVECs

CD31- cells were isolated from the SVF by Dynabeads. Endothelial differentiation was performed in a stem cell medium (EC-

3, A.-B.) and an endothelial differentiation medium (EC-2, C.-D.) on Matrigel™. As control human umbilical vein endothe-

lial cells (HUVECs) were cultured in EGM-2 medium on Matrigel™ (E.-F.). Endothelial differentiation was analyzed by

light microscopy. Shown are bright field micrographs at a magnification of x100. ►, capillary tubules; , three-dimensional

networks.

74

A.

B.

CD34 KDR

acLDL

CD34 KDR

acLDL

Figure 20. Flow cytometry analysis of CD31- cells differentiation under optimized medium conditions

CD31- cells were isolated from the SVF and endothelial differentiation was induced by applying a stem cell medium (EC-3,

A.) or an endothelial differentiation medium (EC-2, B.), respectively. Quantitative expression of the surface markers CD34,

KDR, and acLDL was determined in flow cytometry.

75

culture medium EC-2 culture medium EC-3

CD34+ 2% ± 1% 37% ± 9%

KDR+ 19% ± 15% 78% ± 12%

acLDL 70% ± 5% 77% ± 7%

Table 6. Expression of surface markers on CD31- SVF cells under endothelial medium conditions

To evaluate differentiation of CD31- cells towards endothelium, CD31- cells were isolated from the SVF by Dynabeads and

grown in EC-2 or EC-3, respectively, followed by quantitative analysis of the expression of the surface markers CD34, KDR,

and acLDL (compare Figure 20). Data are from three individual experiments.

CD34 KDR

CD31 CD51/61

acLDL

Figure 21. Flow cytometry analysis of HUVECs

HUVECs were used as control in flow cytometry analysis for the presence of endothelial cell markers as CD31, CD51/61,

CD34, KDR and acLDL uptake.

76

IN VITRO DIFFERENTIATION OF SVF CELLS UNDER ADIPOGENIC MEDIUM CONDI-TIONS

Unseparated SVF cells from excised tissue differentiate under adipogenic culture conditions

to almost 90% into adipocytes showing large lipid vacuoles after 21 days in adipogenic me-

dium (data published elsewhere). Regarding the separated SVF cell fractions, the CD31-

cells could differentiate into mature adipocytes by nearly 100%, as demonstrated by light

microscopy and Oil Red O staining in Figure 22 A.-D.. Interestingly, purified CD31+ mature

endothelial cells from the SVF also convert into adipocytes after 21 days under adipogenic

culture conditions. Undergoing differentiation, cells are viable and healthy with no signs of

apoptosis present. From microscopical cell counting, approximately 65% of CD31+ cells pre-

sented accumulation of lipid vacuoles equivalent to an adipocyte phenotype. As can be seen

from Figure 22 E.-G., CD31+ cells have CD31 Dynabeads attached to their cell membranes

which confirms their CD31 surface expression. When comparing GDPH activity of both CD31

subsets, CD31+ cells showed a GPDH activity of 64% ± 11% compared to GPDH levels of

CD31- cells set 100% (Figure 23). Lipid droplets appeared approximately 2-3 days later dur-

ing differentiation in CD31+ than in CD31- cells. Applying flow cytometry, the proliferating

CD31+ cell fraction was screened for the presence of the surface marker CD34 before induc-

ing adipogenic differentiation. As shown in Figure 18 O.-P., approximately 45% of the cells

express CD34.

77

CD31- CD31+

A.

B.

C.

D.

E.

F.

G.

H.

200µm

200µm

200µm

200µm

200µm

100µm

100µm

200µm Figure 22. Morphological analysis of in vitro differentiation of SVF cells under adipogenic medium conditions

SVF cells were separated into CD31- (A.-D.) and CD31+ (E.-H.) by Dynabead treatment. Both fractions were independently

differentiated towards mature adipocytes. All pictures, except for D. and H. which shown oil red staining results, are bright

field images. Shown in the left upper corner of G. is a high-power magnification of a differentiating cell with Dynabeads

attached. Arrows in E., F., and G. indicate Dynabeads attached to cell surfaces. Magnification in A.-D., E. and H. is x100, in

F. and G. is x200.

78

0

20

40

60

80

100

120

CD31- Diff CD31+ Diff

Ext

ento

f diff

eren

tiatio

nre

lativ

e to

CD

31- c

ells

[%]

*Figure 23. Quantitative analysis of in vitro

differentiation of SVF cells in adipogenic

medium

SVF cells were separated into CD31- and

CD31+ cells by Dynabeads, independently

cultured in proliferation medium, and differen-

tiated at confluence. Differentiation of the

CD31+ fraction as determined by GPDH activ-

ity was measured after 3 weeks relative to

CD31- cells. Results are from three individual

experiments. *, p=0.01 compared to standard

differentiation of CD31- cells.

79

DISCUSSION

The stromal vascular fraction (SVF) of adipose tissue is a heterogenous cell population and

therefore very hard to characterize. Although there are similar findings about the surface pro-

tein expression of SVF cells, much controversy remains. The cell fractions that have been

used for surface analyses and differentiation assays cannot be compared since some re-

search groups apply liposuction aspirates60,252 133 or excised fat tissue260,314; others use more

purified, cell-sorted fractions260,350, while still others have identified cell surface structures

from cells not before culturing for several passages60. This leads to cell fractions with differ-

ent surface characteristics and different proliferation and differentiation behaviours. In this

thesis SVF cells from lipoaspirate and excised fat were screened for the presence of CD34,

KDR, CD31 and CD51/61. The rational for this was threefold: to (i) illustrate the differences

between SVF cells isolated from excised versus liposuction material, (ii) to determine the

relative amount of endothelial and hematopoietic progenitor cells in the freshly isolated SVF,

and (iii) to be aware of potential cellular contaminations, especially endothelial cells, which

might cause a false-positive differentiation result. Additionally, cells were cultured for two

weeks in standard medium to promote proliferation and to evalutate possible changes of sur-

face protein expression under long-term culture conditions.

After isolation of SVF cells the cell pellet is naturally contaminated with endothelial cells.

When measuring the differentiation potential of SVF cells into endothelial cells, already in-

corporated ECs might mimic a successful transdifferentiation of SVF cells. Therefore, it is not

possible to evaluate whether endothelial cells were transdifferentiated from SVF cells or if

they were already present in the applied SVF cell fraction before starting the experiment. To

determine the growth potential of contaminating ECs after isolation of SVF cells, different EC

growth media and a standard proliferation medium for preadipocytes were tested.

To analyze the differentiation plasticity of SVF cells towards endothelial structures and adi-

pose tissue, exclusively purified cell fractions were used to exclude cell contamination and

allow a solid analysis. Therefore, the cell pellet was splitted into a CD31+ and a CD31- cell

fraction using paramagnetic beads. After confirming purity of both cell fractions, CD31+ and

CD31- SVF cells were cultured in EC media or adipogenic differentiation medium. For endo-

thelial differentiation only EC-2 and EC-3 medium were applied as these two media showed

best potential for EC proliferation and differentiation according to Study 3.

80

TISSUE ENGINEERING WITH ADIPOSE DERIVED ADULT STEM CELLS

Soft tissue defects remain a challenge and often difficult problem in plastic and reconstruc-

tive surgery. There is high clinical need for adequate reconstruction of extended soft tissue

defects as found after deep burns, trauma, or tumor resections.

The application of autologous fat tissue to repair soft tissue defects seems most reasonable

in its approach as excess amounts of adipose tissue are found all over the human body. Adi-

pose tissue can be easily obtained through liposuction or elective operations and has been

used for transplantation or in soft tissue tissue engineering projects. Although there is re-

markable progress in this research field, serious problems remain. Transplantation of autolo-

gous adipose tissue to a defect site often results in 40-60% of graft volume loss over time.

One reason for significant tissue resorption is that large fat grafts never acquire sufficient

vascularization. Inadequate tissue vascularization restrains the supply of oxygen and nutri-

ents to the graft followed by consecutive tissue destruction406 and long-term tissue resorpti-

on304,303. Therefore the use of mature fat tissue for transplantation has not been consistently

successful in patients58,95,108,201,304,305,362.

More promising in the context of soft tissue reconstruction is the use of adipose tissue de-

rived progenitor cells seeded on biomaterials. By harvesting viable cells from patients healthy

autologous preadipocytes can be expanded through cell culture replication. Cells may then

be seeded onto scaffolds which need to support cell growth, proliferation and differentiation

of the progenitor cell fraction into mature fat cells. The scaffold construct is normally ab-

sorbed over time due to its biodegradable material and ideally, exclusively mature, well-

vascularized healthy adipose tissue remains.

Preadipocytes are deemed more advantageous as cell source for adipose tissue engineering

than mature adipocytes because they can be easily obtained, cultured, and expanded. Due

to the ability to generate large cell numbers and finally differentiate into various cell types

other than adipocytes97,157,162,433,434, preadipocytes are thought to have high potential in the

future development of soft tissue engineering.

PRESENT PROBLEMS AND HURDLES

Though several strategies using polymeric materials seeded with preadipocytes have been

successful to induce adipose tissue formation in vivo, one often quoted shortcoming with

these strategies is delayed growth in volume of the tissue constructs. This has been attrib-

uted mainly to slow vascularisation of the tissue leading to tissue resorption, oil cysts and

fibrosis253,307,363. Oxygen and nutrients supply are the limiting factors for the survival of trans-

planted cells. Especially adipose tissue engineering fails due to inadequate blood supply as

mature fat cells face cell death if they do not get in contact with surrounding capillaries by

81

inosculation within the first four days363,408. In conclusion, sufficient vascularisation and ap-

propriate cellular differentiation are two key problems in soft tissue engineering.

A further problem in organ and tissue transplants which will come into place once cell differ-

entiation and vascularisation have been addressed is the immunological rejection of the do-

nated tissue by the patient in non-autologous methods or cell culture using FCS. Strategies

in tissue engineering ascertain possible solutions to these problems225, including the applica-

tion of preadipocytes instead of adipocytes due to better proliferation capacity34,304 and hy-

poxia tolerance in precursor cells406, the exertion of autologous serum during in vitro cell cul-

ture160,161 and improvement of applied biomaterials159 among others.

HOW TO MANAGE INSUFFICIENT VASCULARISATION

Current methods to induce neovascularisation, the formation of a new vascular network,

within a tissue-engineered construct reveal limitations. Methods for adipose tissue angio-

genesis include co-culture of adipose cells with ECs in tubular scaffolds to induce vessel

formation. Although growth factors help in the formation of capillary structures, the generated

vessels are not long lasting304,303. The creation of vasculature prior to implantation allows the

tissue to receive nutrients faster because it does not have to form its vasculature. However,

the tissue may still not receive the required nutrition47. Other vascularisation methods com-

prise ingrowing of host vessels into the tissue construct leading to revascularisation of the

implanted material, but again insufficient nutrient supply remains a problem for the center of

the scaffold304. The relationship between ADAS cells and ECs as well as the role of certain

growth factors are not sufficiently understood to proceed towards a viable graft of relevant

size. Additionally, the potential of ADAS cells to differentiate into adipocytes and ECs upon

availability of the proper stimuli could be the key for confecting a vascularised fat graft with

sustained and increased volume. Therefore, this thesis studied the differentiation potential of

cells to convert from adipose tissue to endothelium and vice versa via progenitor cells.

CHARACTERIZATION OF THE SVF

Taking the broad mixture and combinations of cell surface CD markers, it is logical that the

SVF is not a unique population, but rather a pool of various progenitor cells including mature

ECs, too.

ADAS cells exhibit differentiation plasticity similar to that of bone marrow stem cells, as men-

tioned in the introduction. The cells are multipotent adult stem cells and have the ability for

multilineage differentiation429,433,434. Further similarities between ADAS cells and BMSCs

have been described in the literature focussing on growth kinetics and expansion capacity,

cell senescence, and protein surface expression (compare Table 1).

82

As shown by flow cytometry, freshly isolated SVF cells from excised as well as liposuctioned

adipose tissue comprise a significant population of progenitor cells as shown by strong ex-

pression of CD34 and KDR (Figure 15). However, the SVF also contains a population of

CD31, CD51/61, and vWF positive cells, demonstrating a contamination with mature ECs.

CD31 and CD51/61 expression was stronger in liposuction SVF cells, with up to 40% CD31

positive cells, emphazising the higher amount of contaminating ECs in lipoaspirates than in

excised fat (5-10% ECs, Table 5). The vWF expression of 10% for excised tissue as deter-

mined by microscopical cell counting (Figure 18 A.) completely matches the findings on

CD31 and CD51/61 expression in flow cytometry (compare Table 5).

After cell culture in DMEM/F12 with 10% FCS for two weeks, cells from lipoaspirate remain

positive for CD34 to nearly the same percentage as freshly isolated cells while cells from

excised tissue show a significant reduction (p<0.05) (Table 5). The missing reduction in

CD34 after cell adherence in liposuction material is surprising and the cause of this still is

under investigation. Endothelial surface characteristics, such as CD31 and CD51/61, how-

ever, decrease during culturing (Table 5) in all conditions, which might be due to selection

effects from culture medium as well as lower proliferation capacity of CD31+ SVF cells. CD31

expression in Figure 15 A. appears rather homogenous compared to Figure 15 B., however,

this is due to the two different preparation methods. In Figure 15 A., excised fat lobules were

carefully isolated to only gain preadipocytes. Since this is macroscopically not possible, there

is a very small but homogenous fraction of endothelial cells displaying CD31 characteristics.

The presence of these two cell fractions causes a double-peaked flow cytometry curve. In

Figure 15 B., liposuction material was used, which contains a much higher amount of endo-

thelials cells due to its origin and the method of yield which generates the two-peak appear-

ance of the flow cytometry.

KDR expression in liposuction material is initially 43% ± 23% and decreases to 23% ± 7%,

perhaps due to culturing conditions. This is in accordance with the reduction of endothelial

markers. In excised adipose tissue, however, the findings of KDR expression remain contro-

versial since KDR expression immediately after isolation is surprisingly low and its increase

due to cultivation is not in agreement with liposuction results (Table 5).

In general stem cell markers (CD34, KDR and S100) on SVF cells decrease during long-term

culture in standard medium as demonstrated in Figure 16. This is in agreement with the find-

ing that preadipocytes loose their differentiation capacity after multiple passages.

To analyze CD31+ and CD31- differentiation characteristics and surface markers independ-

ently, unpurified SVF cells from excised adipose tissue were split by CD31 Dynabeads. Al-

ternatively, CD31 MicroBeads were applied for the cell separation if double-labeling was per-

formed.

83

Whether or not Dynabead attachment to the cell surface influences the differentiation poten-

tial of preadipocytes has not been analyzed so far. In order to minimize any impact of the

Dynabeads, concentration of beads was kept low to allow proper attachment of cells to cul-

ture dishes which is otherwise impaired. As Dynabeads might bind non-specifically to adher-

ent and phagocytic cells, all steps with Dynabeads were carried out at 4°C. At this tempera-

ture, cell activity is low and non-specific binding reduced to a minimum. To further minimize

non-specific binding, the bead-cell suspension was pipetted a few times after incubation

(prior to separation on the magnet) to loosen any non-target cells adhering to the beads. All

these procedures reduced non-specific binding of Dynabeads which was finally evaluated by

staining Dynabead-carrying cells for vWF to ensure the endothelial phenotype. Cell counting

with fluorescence and light microscopy revealed that 95 ± 5% of the cells carried Dynabeads

(Figure 18 F.-I.). Of this fraction, 97 ± 2% were positive for vWF. This is supported by flow

cytometry after MicroBead cell sorting (Figure 18 M.-N.). These results reveal the beneficial

effects of the above shown technical modifications in Dynabead usage and show that 92 ±

4% of CD31+ cells are vWF positive. Together with the high expression of CD105 in CD31+

cells (Figure 18 K.-L.), this confirms the mature endothelial cell phenotype and the unique-

ness of the CD31+ fraction.

Freshly harvested CD31- SVF cells, in contrast to CD31+ cells, show no expression of vWF

and CD31 as ECs would do, neither after one nor after two weeks of culture in proliferation

medium (data not shown). Furthermore, CD31- cells do not present endothelial network for-

mation on plastic culture dishes (Figure 18 C.). Flow cytometry analyses reveal that this cell

fraction is 100% positive for S100 (Figure 18 D.-E.). Even though S100 labels nervous tis-

sue, skeletal and cardiac muscle, malignant melanoma cells and some other cell types, it is

well accepted as an early marker of adipogenesis21. It can therefore be assumed that the

CD31- fraction consists of adipocytes in a precursor state, the original preadipocytes. Thus,

separation of SVF cells by CD31 Dynabeads generates one cell fraction with preadipocyte

characteristics (CD31-) and one with endothelial properties (CD31+). SVF cells have recently

been shown to dedifferentiate from mature fat cells into a progenitor cell type and to differen-

tiate from there into ECs314. This has high implications for angiogenesis-related and cardio-

vascular topics.

In conclusion the surface protein expression of SVF cells demonstrates characteristics of

hematopoietic stem cells, early adipogenesis, ECs and endothelial progenitor cells (see Fig-

ure 24).

84

S100

CD34

KDR CD31

CD51/61

CD133

CD144

CD105

earlyadipogenesis

endothelial cell

hematopoietic stem cell

SVF

ADAS CELL DIFFERENTIATION TOWARDS ENDOTHELIAL CELLS

SVF cells have the capacity to differentiate into multiple cell phenotypes including all three

germ sheet lineages as mentioned earlier2,55,69,84,107,291,314,433,434. In addition to the remarkable

differentiation capacity of SVF cells, these cells have recently been differentiated into an en-

dothelial phenotype when cultured under EC conditions60,260,314. The first study demonstrating

the capacity of SVF cells to differentiate into capillary-like structures was already accom-

plished in 1999 by Zangani et al.429. When cultured on reconstituted basement membrane

(RBM) from Engelbreth-Holm-Swarm mouse sarcoma, mammary stromal cells rapidly turned

into capillary structures containing an internal lumen. The network formation was intensified

by VEGF supplementation and inhibited by the anti-angiogenic agent suramin. Differentiated

mammary stromal cells expressed KDR as well as ZO-1, a tight-junction-associated protein.

Conversion into mature adipocytes was still possible when cultured under adipogenic condi-

tions. More recent studies used other ECM components than RBM, such as methylcellulose

or Matrigel Extracellular Matrix as well as the presence of VEGF in combination with miscel-

laneous other growth factors for the differentiation of ADAS cells into ECs. When cultured in

the presence of ECM proteins and VEGF, ADAS cells develop branched tubular structures

and express various EC markers, such as CD31, CD34, vWF, CD144, and eNOS60,260,314.

The differentiation of mesenchymal stem cells into mature ECs has great impact on clinical

applications as well as tissue engineering research, as diffusion can supply cells just over a

distance of about 150 µm with nutrition, which is insufficient in ischemic disease and also in

larger three-dimensional fat tissue constructs in tissue engineering26. New capillaries from

the surrounding tissue invade after a delay of about five days and are not able to reach the

Figure 24. Cluster of

Differentiation com-

parable to bone

marrow stem cells

Freshly isolated SVF

cells have nearly the

same surface character-

istics as stem cells

from bone marrow and

display mesenchymal

stem cells.

85

centre of the construct. This leads to cell necrosis especially in highly vascularized tissues

like adipose tissue31.

THERAPEUTIC ANGIOGENESIS

In 2004, Planard-Bernard et al. demonstrated a common origin for adipose tissue and endo-

thelium314. They postulated that adipocytes and ECs have a common progenitor; however,

the interlinking pathways between adipose tissue and endothelium and the differentiation

potential from one tissue into the other via progenitor cells have not been analyzed. So far it

has only been shown that SVF cells are able to perform metaplasia from mature fat cells into

endothelial cells314. This thesis focused on this concept and aimed to further analyze the

interlinking pathways between adipose tissue and endothelium.

ADAS cells show a remarkable potential to form capillary-like structures in vitro. Further,

ADAS cells resemble EC characteristics in in vivo models as well. Angiogenic potential of fat

tissue has been known and clinically applied for treating wounds and ischemic organs for

more than 400 years (first mentioned in 1585)360,432. Applied for therapeutic angiogenesis,

ADAS cells have a wide range of possible clinical applications in neovascularization266,380,

including peripheral vascular disease, ischemic cardiomyopathy and myocardial infarction262,

ischemic stroke193, diabetic retinopathy, acute tubular necrosis91,189 and transplant related

ischemia. ADAS cells have first demonstrated enhanced angiogenesis and improved vascu-

larization in hindlimb ischemia models in mice60,260,266,274,275,314,325. Two potential but nonex-

clusive mechanisms could be involved in the pro-angiogenic effect of ADAS cells. Injected

cells could directly contribute to incorporation into the vessel wall60,260,314, or cells release

angiogenic growth factors which improve angiogenesis48,274,314. The interaction of both is

most likely the cause for blood flow enhancement, increased capillary density, and improved

angiogenesis induced by ADAS cells.

To study the angiogenic potential of adipose tissue derived cells, Cao et al. used CD34-

CD31- Flk-1+ cells60 that displayed features of EPCs. Miranville et al. applied CD34+ CD31-

ADAS cells260. Most other studies utilised unpurified adipose mesenchymal stem

cells274,314,325. Regardless of the applied subpopulation, all ADAS cell fractions improved an-

giogenesis, vascularization and blood flow, although CD34+ cells seem to be superior to

CD34- cells in increasing capillary density in hindlimb ischemia260. CD34- cells induce CD34

expression in ischemic tissue and vessels60 supporting the fact that CD34 is a crucial marker

for neo- or revascularization. Administration of ADAS cells can potentially affect revasculari-

zation to a degree similar to the application of BMSCs260.

A recent study has introduced a new method to approach decreased cardiac function after

myocardial infarction with transplantation of monolayered ADAS cells onto the scarred myo-

cardium four weeks after ischemic injury262. After application of ADAS cells with cell sheet

86

technology, the engrafted sheet gradually grew shaping a thick layer of tissue and compris-

ing freshly emerging blood vessels and a couple of cardiomyocytes. Additionally, angiogene-

sis was initiated in a paracrine manner and cardiac function improved subsequently in rats

with myocardial infarction262.

Another hint for the close relationship between adipose tissue and blood vessels is given by

preadipocytes that transdifferentiate into smooth muscle cells (SMCs)1. By overexpression of

aortic carboxypeptidase-like protein (ACLP) in adipogenic precursor cells a smooth muscle-

like cell phentotype can be induced in SVF cells and clonal preadipocyte cell lines1. ACLP is

expressed in ECM components of collagen-rich tissues229 and is up-regulated during vascu-

lar SMC differentiation228. Under smooth muscle (SM) differentiation environment, with β-

mercapto-ethanol and ascorbic acid, preadipocytes express anti SM22a, SM myosin heavy

chain, caldesmon, SM α-actin and develop stress fibers1.

To further verify the theory on mesenchymal differentiation plasticity, we performed morpho-

logical studies on CD31- SVF cells seeded onto growth factor reduced Matrigel™ in different

culture media, EC-2 or EC-3, respectively. EC-2 and EC-3 were chosen according to Study 3

as these two media showed the best results in inducing endothelial cell morphology in SVF

cells. The first medium (EC-2) is a culture medium to induce differentiation towards endothe-

lial lineage260. The second culture medium (EC-3) used has been applied as a hematopoietic

and endothelial progenitor cell medium for peripheral blood progenitors125. CD31- cells, which

were initially 100% positive S100 (compare Figure 18 D.-E.), were able to form three-

dimensional tube like structures in both media (Figure 19 A.-D.). EC-2 medium additionally

allowed the formation of extensive intercellular tube networks (Figure 19 C.-D.). HUVECs

used as a positive control also formed networks but were not able to form three-dimensional

capillary-like structures (Figure 19 E.-F., compare to Figure 19 C.). Analyzing endothelial

differentiation of CD31- cells grown in EC-2 or EC-3 by flow cytometry, we found that in EC-3

medium CD31- SVF cells express CD34 up to around 40% and KDR of 80% (Figure 20 A.,

Table 6). EC-3 medium therefore seems to support endothelial precursor characteristics in

the CD31- cells. In EC-2 medium, in contrast, CD34 and KDR expression was significantly

lower (Figure 20 B., Table 6) demonstrating the capacity of this medium to enhance endothe-

lial differentiation to mature ECs and diminish endothelial precursor cell characteristics of

SVF cells. In both media cells were positively stained for acLDL confirming differentiation

towards an endothelial phenotype (Figure 20, Table 6). AcLDL labels endothelial progenitor

cells, vascular endothelial cells, and macrophages.

Adipose tissue contains a subtype of mesenchymal stem cells that owes the ability to differ-

entiate into mature ECs in vitro, migrate into peripheral blood vessels, enhance angiogenesis

87

and revascularization by differentiating into ECs in vivo. Cells with the same abilities derived

from bone marrow as well as from blood circulation have been described since 199717 and

were termed “endothelial progenitor cells” for their EC potential. EPCs from bone marrow are

positive for CD34, VEGFR-2/KDR, CD133 and have the ability to take up

acLDL17,74,125,171,181,301,333,397,398,424. During mobilization and subsequent blood circulation they

lose CD133 and start expressing CD31, vascular endothelial cadherin, and vWF. Although

most subpopulations of ADAS cells have unequal surface protein patterns, various unique

markers were detected on them that resemble EPC characteristics. But not only the surface

characteristics reveal similarities between EPCs und ADAS cells. ADAS cells seem to par-

ticipate in endothelial repair, neovascularisation and angiogenesis in almost the same man-

ner as EPCs. Indeed, ADAS cells and EPCs seem to be involved in the repair of damaged

endothelium60,260,314. However, little is known about the signals associated in directing cells to

the centre of injured endothelium. Both ADAS cells and EPCs are capable of transdifferentia-

tion into smooth muscle cells1,114 underlining the ability of vessel wall integration. EPCs may

participate in remodelling of ischemic cardiac tissue and regeneration of ischemic myocar-

dium by modulation of angiogenesis and myogenesis190,205,215,348 in an almost comparable

manner as ADAS cells262,373.

EPCEC

smooth muscle cell

damaged EC

ADASEC

smooth muscle cell?

damaged EC

Figure 25. Vessel wall repair

through EPCs and ADAS cells

Similar to endothelial progenitor

cells ADAS cells may take part

in the repair of injured vessel

walls. Some of the ADAS cells

may integrate into the endothe-

lial monolayer after transdiffer-

entiation into endothelial and

even smooth muscle cells.

88

Some research groups aimed to identify a cell population that exhibits EPC characteristics in

adipose tissue. A recent study has isolated KDR positive cells from human lipoaspirates,

enhanced KDR expression through incubation in a hematopoietic stem cell medium and sub-

sequently differentiated the KDR+ subpopulation towards the endothelial lineage252. The

KDR+ subpopulation of LPA cells, pronounced as adipose EPCs, differentiated into ECs of

about 95%, being positive for VE-cadherin, vWF, LDL-lectin and formed tubular structures.

Although the KDR+ cells had striking characteristics of EPCs, an initial phenotyping and

characterization of the EPC fraction (KDR+ cells) was not performed. Flow cytometry for this

cell fraction was only performed for KDR alone, CD34, CD133 and acLDL uptake were not

obtained. Therefore it remains questionable whether the KDR+ subpopulation contained adi-

pose derived EPCs.

In another study, KDR+ ADAS cells were isolated and induced to perform endothelial lineage

conversion by plain culturing in EC medium. Differentiated ECs expressed CD31, CD34, VE-

cadherin and eNOS and were capable of tubular structure formation on Matrigel extracellular

matrix60. Even though the results so far display tremendous insights into the specific differen-

tiation potential of various cells, further investigations are needed to reveal the true potential

of the different mesenchymal stem cell fractions from adipose tissue to undergo EPC differ-

entiation.

ENDOTHELIAL CELL DIFFERENTIATION TO ADIPOCYTES

CD31+ and CD31- were separately cultivated under adipogenic medium conditions to analyze

their ability to convert into adipocytes. As expected, CD31- cells fully differentiated into ma-

ture adipocytes (Figure 22 A.-D.). Interestingly, purified SVF CD31+ cells also differentiated

into mature adipocytes after 21 days under adipogenic culture conditions. As can be seen

from Figure 22 E.-G., CD31+ preadipocytes and maturing adipocytes have CD31 Dynabeads

attached to their cell membranes confirming their CD31 surface expression. Further they

were positive for CD105 and vWF, additionally confirming their origin from endothelial line-

age (Figure 18 F.-N., Figure 26). While CD31+ and CD31- cells present a different morphol-

ogy during proliferation (compare Figure 22 A., 22 E.), they look identical in differentiated

state as adipocytes (Figure 22 D., 22 H.). Numbers from microscopic cell counting well

match with the more precise findings from GPDH assays in which CD31+ cells show a GPDH

activity of 64% ± 11% compared to CD31- cells set 100% (Figure 23). Thus, these findings

present for the first time a CD31+ progenitor cell type that can either convert into EC or adi-

pocyte, depending on the culture conditions (Figure 26). The CD31+ cell fraction is also

CD34+, since CD34 was expressed by about 45% of cells in flow cytometry (Figure 18 O.-P.).

The fact that CD34 and GPDH percentages do not completely match is due to the incongru-

ency between cell number (represented by CD34-level) and cell volume (GPDH-level).

89

Preadipocytes

Adipose Tissue EC

Endothelial diff.

Adipogenic diff.

MatrigelTM

Adipogenic diff.

MatrigelTM

Endothelial diff.

CD31- S100+

CD31+ CD105+ vWF+

Figure 26. Morphological analysis of in vitro differentiation of CD31- and CD31+ adipose progenitor cells under

endothelial and adipogenic differentiation conditions

Adipose progenitor cells were separated by Dynabeads for ECs in a CD31- and CD31+ cell fraction, both fractions were inde-

pendently differentiated towards endothelial structures on Matrigel as well as towards mature adipocytes.

In conclusion, the results presented here that adipose tissue is a source of mesenchymal

progenitor cells with broad differentiation plasticity. It is not exclusively the CD31- S100+ cell

population in adipose tissue, the classical preadipocytes, which can undergo fat differentia-

tion, but also the CD31+ CD105+ vWF+ cell fraction, phenotypically adipose tissue derived

ECs (compare Figure 26). In turn, preadipocytes even though predetermined to naturally

differentiate to adipocytes, have the potential to mature to endothelial network structures.

Conclusively, the adipose SVF appears to be a pool of multipotent cells with high plasticity

and differentiation capacity towards endothelium as well as adipose tissue. This might give

rise to innovative strategies in further tissue engineering applications where the adipose SVF

is applied for both, engineering of fat tissue and vascular structures.

90

ARTERIOSCLEROSIS AND ADIPOSE TISSUE

Arteriosclerosis affects large and medium-sized muscular and elastic arteries. Pathological

conditions, such as endothelial dysfunction, vascular inflammation, accumulation of lipids,

cell death and calcification of the intima lead to an atherosclerotic plaque of the vessel wall

(Figure 27). By protruding into the internal artery, plaques minimize the luminal diameter

available for blood flow, which results in abnormalities of blood flow, vascular remodelling,

acute and chronic obstruction and subsequent decreased oxygen supply.

In the past atherosclerosis was seen as a bland lipid storage disorder, however, substantial

advances in basic and experimental science have highlighted the role of inflammation as well

as cellular and molecular components supporting plaque evolvement.

Blood leukocytes, mediator cells of inflammation and host defence, play a major role in local-

ising the earliest lesions of arteriosclerosis and initiate the process of chronic plaque forma-

tion. Healthy endothelium does physiologically not allow leukocyte binding (Figure 27 A.).

However, after endothelial injury at sites of shear stress in the circulation and atherogenic

diet, inclusion of lipoproteins in the artery wall with accumulation of lipids in the subendothe-

lial space takes place (Figure 27 B.). Augmented expression of selective surface adhesion

molecules on arterial ECs promotes enhanced leukocyte recruitment237,273. When vascular

endothelium becomes inflamed, increased expression of adhesion molecules as vascular cell

adhesion molecule 1 (VCAM-1) and intercellular adhesion molecule 1 (ICAM-1) lead to bind-

ing of leukocytes to the activated ECs and integrins mediate firmer attachment (Figure 27

C.). Further blood flow disturbance promotes production of proteoglycans in the vessel wall,

that can bind lipoprotein particles, easing their oxidative modification and therefore advantag-

ing an inflammatory response234. Proinflammatory cytokines expressed within the artheroma

provide a chemotactic stimulus to the adherent leukocyte and promote penetration into the

intima. Infiltration of immune cells together with proteoglycan changes result in fatty streak

accumulation. Certain inflammatory mediators (MCP-1, M-CSF) support expression of

macrophage scavenger receptors, promoting the adhesion of circulating monocytes entering

the intima. Monocytes convert into macrophages when taking up the modified lipoproteins

located in the intima. These lipid-laden macrophages together with T-lymphocytes form a

lipid core that is isolated from the blood flow by a fibrous cap generated by migrated

SMCs371. Activated leukocytes and intrinsic arterial cells release mediators regulating this

process, comprising growth factors and cytokines such as TGF-β, PDGF BB, TNF-α and -β

and interferon-γ42,361. Fibrogenic growth factors stimulate SMC replication and the production

of a dense ECM leading to a more advanced atherosclerotic lesion (Figure 27 D.)336. But

inflammatory mediators can further evoke the expression of collagenase by foam cells in the

intimal plaque which comprehends thinning of the fibrous cap, leaving it weak and suscepti-

ble to rupture. Final evolvement of the end-stage plaque implies also further accumulation of

91

inflammatory cells, specially macrophages and T-lymphocytes, apoptosis and necrosis of the

central part of the lesion and calcification, resulting in evolvement of a necrotic core (Figure

27 E.)371. Cross-talk between macrophages and T-lymphocytes heightens the expression of

the potent pro-coagulant tissue factor238. When thinning of the fibrous cap and blood flow

disturbance lead to plaque rupture, the tissue factor triggers fatal thrombus formation that

causes arterial occlusion and a subsequent acute ischemic event of the target organ.

92

A.

B.

C.

D.

E.

endothelial cell

smooth muscle cell,contractile

normal ECM

hypertrophic ECM/fibrous cap

LDL/ lipoproteins

lymphocyte

monocyte/blood macrophage

foam cell

lipid core with necrosis

smooth muscle cell,synthetic

Figure 27. Stages of atheroscle-

rosis

A. Healthy endothelium. Blood

leukocytes adhere poorly to normal

endothelium. B. Inclusion of lipo-

proteins in the artery wall after

biochemical injury and subsequent

lipid accumulation. C. Leukocyte

recruitment to the nascent athero-

sclerotic lesion. When the endothe-

lial monolayer becomes inflamed, it

expresses adhesion molecules that

bind leukocytes. Proinflammatory

cytokines expressed within

atheroma provide a chemotactic

stimulus to the adherent leuko-

cytes, directing their migration into

the intima. Inflammatory mediators

can augment expression of macro-

phage scavenger receptors leading

to uptake of modified lipoprotein

particles and formation of lipid-

laden macrophages, foam cells. D.

Lymphocytes join macrophages in

the intima during lesion evolution.

Leukocytes, together with resident

vascular wall cells, secrete cyto-

kines and growth factors that can

promote the migration and prolif-

eration of smooth muscle cells. E.

Inflammatory factors can inhibit

collagen synthesis and evoke the

expression of collagenase by foam

cells. These alterations thin the

fibrous cap, leaving it fragile and

susceptible to rupture.

93

Numerous studies suggest that adipose tissue may have a strong impact on most obesity-

related cardiovascular complications, since an interrelation between adipocyte secreted pro-

teins and insulin resistance, hypofibrinolysis, inflammation, thrombosis and artherosclerosis

becomes more and more evident. Adipose tissue controls not only energy metabolism, but

further displays a secretory tissue for adipocytokines, which modify vascular response, for

proinflammatory cytokines, and for PAI-1, abetting fibrin accumulation. Most of these factors

produced by AT illustrate a major clue in the evolvement of the metabolic syndrome in obe-

sity. The metabolic syndrome is defined as cumulative cooccurrence of visceral obesity,

dyslipoproteinaemia with increased LDL and triglycerides but decreased high density lipopro-

tein (HDL) and essential hypertension. The beginning of metabolic syndrome is dominated

by an insulin resistance of the insulin-dependent tissues and impaired glucose tolerance,

requiring an increased insulin level for cellular glucose utilisation. This hyperinsulinaemia

heightens the feeling of hunger, resulting in obesity and progression of atherosclerosis323.

Recently, even more constituents of the metabolic syndrome have been revealed, compris-

ing subclinical inflammation, hyperuricaemia and microalbuminuria.

Among the adipocytokines, leptin as well as adiponectin play a role in the development of

cardiovascular risk factors. Leptin is a glycoprotein produced by mature adipocytes and acts

as a central part in controlling body fat storage. As adipose mass increases, its plasma level

increases as well, resulting in constricted appetite and enhanced energy expenditure. But the

mechanism of leptin does not work as efficient as it should since obese subjects remain

obese despite raised leptin concentration, possibly due to leptin resistance comparable to

insulin resistance in obesity.

Adiponectin is also secreted by adipocytes and correlates positively with insulin sensitivity

levels. This supports a negative association with body weight as plasma levels decrease with

the degree of insulin resistance413. Adiponectin acts as an anti-inflammatory and anti-

atherosclerosis adipokine and accumulates in an injured vessel wall by specifically binding to

collagen type I, III and IV293. Further adiponectin plays a role as a scaffold of newly formed

collagen in myocardial remodelling after ischaemic infarction182 and reduces the expression

of adhesion molecules on endothelial including VCAM-1 and ICAM-1296, which allow initial

leukocyte recruitment in atherosclerosis237,273. It inhibits TNF production by proliferation inhi-

bition of myelomonocytic lineages and mature macrophage functions425, reduces oxidized

LDL-induced EC proliferation270, and suppresses macrophage conversion into foam cells295.

Lower risk of myocardial infarction comes therefore together with high levels of circulating

adiponectin, whereas subjects with atherosclerosis seem to have low adiponectin concentra-

tions217,278. In contrary, resistin produced by adipose tissue induces the expression of endo-

thelial adhesion molecules VCAM-1 and ICAM-1206,404.

94

THE IMPACT OF PROGENITOR CELLS IN ATHEROSCLEROSIS

Besides the classical hypothesis of the atherosclerotic process recent research findings indi-

cate the involvement of stem and progenitor cells from the blood circulation in the develop-

ment of arteriosclerotic lesions. Progenitor cells were found to take part not only in endothe-

lial repair but also in smooth muscle cell accumulation in the vessel wall. In animal models

the injection of EPCs or BMSCs resulted in an aggravation of atherosclerosis and unstable

plaque evolvement126,426. Atherosclerosis turns out to be a pathological process initiated by

endothelial cell death, for instance due to trauma, which is followed by EC replacement

through progenitor cell aggregation. LDL can easily accumulate in the intima during the dif-

ferentiation of progenitor cells, a process which takes several days to weeks. Further blood

mononuclear cells adhere to freshly differentiated ECs and migrate into the subendothelial

space. Meanwhile, blood and resident progenitor cells harboured in the adventitia migrate

into the intima, followed by proliferation and differentiation into SMCs. According to that pro-

genitor cells seem to be accountable for the formation of atherosclerotic lesions as a major

cell source by giving rise to undue SMC accumulation172,173,347. In conclusion, stem cells

seem not simply to participate in vascular repair, but further in atherosclerotic lesion forma-

tion via pathological remodelling of vessel walls183,250.

Additionally Daub et al. proved that human CD34+ progenitor cells differentiate into foam

cells and ECs through the regulation of platelets. A changed balance of platelet-mediated

differentiation of CD34+ progenitor cells into foam cells and ECs may thus play a critical role

in atherogenesis as well94.

Taking these research findings and the differentiation plasticity of adipose SVF cells demon-

strated in this thesis, it might be possible that not only blood progenitor cells, such as EPCs,

take part in atherosclerosis, but also progenitor cells from adipose tissue (compare Figure

25).

The broad differentiation plasticity of CD31+ SVF cells could be explained by the common

mesenchymal origin of endothelium and adipose tissue and might have further implications

on the pathogenesis of atherosclerosis. Previous literature findings link the development and

advancement of atherosclerotic lesions to endothelial cells which are thought to be involved

in LDL binding and metabolism391,400, thereby displaying adipocyte characteristics. The

pathogenic processes in atherosclerosis might be not exclusively performed by endothelial or

hematopoietic progenitor cells, but also in part by CD34+ adipose SVF cells as well as CD31+

cells showing adipocyte-related metabolic activities.

95

CONCLUSION

This thesis demonstrates that the term “preadipocyte” can be misleading if purification by cell

sorting or equivalent methods have not been applied. Depending on the method of yield the

amount of endothelial cells in the SVF varies between 10% ± 2% in excised fat and 24% ±

15% in lipoaspirate. The SVF is therefore not a unique cell fraction but a mixture of various

cells with varying surface protein expressions, one of which is a CD31-, S100+ cell type which

has the capacity to differentiate into adipocyte as well as endothelial precursor cell and ma-

ture EC, depending on the involved differentiation-inducing factors. In addition, the SVF fur-

ther contains a CD31+, CD34+, S100- cell fraction with an endothelial phenotype which can

be converted into adipose tissue under adequate differentiation conditions. These results

encourage new approaches in adipose tissue engineering, since the use of purified multipo-

tent SVF cells with adipogenic and angiogenic potential instead of applying the unpurified

SVF cell pool might allow better vascularisation and tissue growth results. Furthermore, the

findings of this thesis might have relevant impact on the pathogenesis of atherosclerosis

since the development and advancement of atherosclerotic lesions seems to involve not only

progenitor cells, but also endothelial cells displaying both, endothelial and adipogenic proper-

ties.

Preadipocytes Adipose EC

Adipocytes Capillaries

Figure 28. Plasticity of hu-

man stromal vascular cells to

perform adipogenic and endo-

thelial differentiation

Besides their natural branch of

differentiation to fat cells and

tubular structures, SVF preadi-

pocytes and endothelial cells can

also differentiate into other

phenotypes. Preadipocytes

(CD31-, S100+) can form tubular

structures on Matrigel, and

endothelial cells (CD 31+,

CD34+, S100-) can aquire a fat

cell phenotype expressing fat-

specific enzymes.

96

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PUBLICATIONS

FULL PAPERS

1. H. Ott, M. Wosnitza, H.F. Merk. Immunologic control parameters during specific immuno-

therapy, Hautarzt – 2008 Jul;59(7):551-6

2. M. Wosnitza, C. Blazek, M. Megahed. Pemphigus herpetiformis, Hautarzt – 2008

Jun;59(6):459-460

3. M. Wosnitza, K. Hemmrich, A. Groger, S. Gräber, N. Pallua. Plasticity of human adipose

stem cells to perform adipogenic and endothelial differentiation, Differentition - 2007

Jan;75(1):12-23

4. K. Hemmrich, G.P.L. Thomas, M. Wosnitza, D. von Heimburg, W. A. Morrison, N. Pallua.

The Isolation and Expansion of Primary Preadipocytes, International Journal of Adipose

Tissue – 2007;1(1):24-33

PUBLISHED ABSTRACTS

1. K. Hemmrich, M. Wosnitza, A. Gröger, N. Pallua. Aus Präadipozyten werden Endothel-

zellen, aus Endothelzellen werden Fettzellen – die Plastizität der Fettgewebsvorläuferzel-

len, Plastische Chirurgie 09/2006; 6, Suppl. 1; 46

2. K. Hemmrich, M. Wosnitza, A. Groger, S. Gräber, N. Pallua. Plasticity of human adipose

stem cells to perform adipogenic and endothelial differentiation, JPRAS 09/2006; 59; S1-

12

SCIENTIFIC PRESENTATIONS

1. M. Wosnitza, C. Blazek, M. Megahed, Aachener Dermatologen Abend, Aachen,

06/2008: Pemphigus herpetiformis

2. M. Wosnitza, K. Hemmrich, A. Gröger, N. Pallua, 14th International Congress of the In-

ternational Confederation for Plastic, Reconstructive and Aesthetic Surgery (IPRAS), Ber-

lin, 06/2007: Adipose tissue is a source of endothelial cells that have the potency to un-

dergo adipogenic differentiation – benefit or pitfall?

3. M. Wosnitza, K. Hemmrich, A. Gröger, N. Pallua, 124. Kongress der Deutschen Gesell-

schaft für Chirurgie, München, 05/2007: Humane Fettgewebsvorläuferzellen ermöglichen

die kombinierte Züchtung von Fettgewebe und endothelialen Strukturen

111

4. K. Hemmrich, M. Wosnitza, C. Gummersbach, D. von Heimburg, N. Pallua. Bernard

O’Brien Institute of Microsurgery Alumni Meeting, Melbourne, Australia, 03/2007: The use

of preadipocytes for adipose tissue engineering

5. K. Hemmrich, M. Wosnitza, A. Gröger, N. Pallua, 37. Jahrestagung der Vereinigung der

Deutschen Plastischen Chirurgen und 11. Jahrestagung der Vereinigung der Deutschen

Ästhetischen Chirurgen, Aachen, 09-10/2006: Aus Präadipozyten werden Endothelzellen,

aus Endothelzellen werden Fettzellen – die Plastizität der Fettgewebsvorläuferzellen

6. K. Hemmrich, M. Wosnitza, A. Gröger, S. Gräber, N. Pallua, European Conference of

Scientists and Plastic Surgeons (ECSAPS), London, England, 09/2006: Plasticity of hu-

man adipose stem cells to perform adipogenic and endothelial differentiation

7. M. Wosnitza, K. Hemmrich, A. Gröger, N. Pallua, Würzburg, 2nd International Conference

"Strategies in Tissue Engineering", Würzburg, 05-06/2006: Human adipose stromal vas-

cular fraction cells allow combined engineering of fat tissue and endothelial structures

8. M. Wosnitza, K. Hemmrich, A. Gröger, S. Gräber, N. Pallua, 41. Congress of the Euro-

pean Society of Surgical Research, Rostock, 05/2006: Potency of human adipose stro-

mal vascular fraction to perform adipogenic as well as endothelial differentiation

112

ACKNOWLEDGEMENT

During the time of my studies working on this thesis I had the fortune to be supported by

many wonderful people whom I owe thanks and gratitude.

Without the grateful support of Prof. Dr. Dr. Norbert Pallua my work on this thesis would not

have been possible at all. His good judgement and patience guided me during the work on

this thesis. Gratitude to him for being a wonderful teacher, for always showing much interest

in my work and for his grateful support. As this is not for granted I owe him special thanks for

all the encouragement and help.

Further I want to thank Christoph Suschek and Karsten Hemmrich for supplying special ad-

vice, supporting my ideas and for making it possible to perform all these experiments. They

were the major force behind this work.

I would also like to acknowledge the wonderful laboratory staff, Tanja Wollersheim and An-

drea Fritz, who thaught and trained me in the basic methods of cell culture and laboratory

methods. They were always available to help me to achieve my goals with excellent advice

and very helpful experience.

I want to thank my parents for always supporting me, for their constant motivation and en-

couragement. Without their help I would never have been able to achieve all this.

113

ERKLÄRUNG § 5 ABS. 1 ZUR DATENAUFBEWARUNG

Hiermit erkläre ich, dass die dieser Dissertation zu Grunde liegenden Originaldaten

bei mir,

Melanie Wosnitza, Wallstr. 15-17, 52064 Aachen,

hinterlegt sind.

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CURRICULUM VITAE Persönliche Daten geboren am 08.07.1980 in Düsseldorf, deutsch, ledig

Eltern Sigrid Wosnitza, geborene Dierdorf, PA, Ass. Geschäftsleitung, Unifrax GmbH, Düsseldorf Achim Wosnitza, Abteilungs- und Projektleiter, RWE AG, Essen/ Ruhestand

Schulausbildung Grundschule Montessorie-Grundschule, Düsseldorf 1987-1991 Gymnasium Städtisches Gymnasium Koblenzer Straße, Düsseldorf 1991-2000 Abitur 06/2000 Universitäre Ausbildung Studium der Elektrotechnik RWTH Aachen WS 2000/01 Studium der Humanmedizin RWTH Aachen WS 2001/02-

WS 2007/08 Examina Physikum 09/2003 Zweiter Abschnitt der Ärztlichen Prüfung 10/2007 Famulaturen Klinik für Plastische Chirurgie, Hand- und Verbrennungschirurgie,

Universitätsklinikum RWTH Aachen 03-04/2004

Hautklinik Universitätsklinikum RWTH Aachen 03-04/2005 Department of Dermatology, University of Tartu, Tartu, Estland 08/2005 Department of Dermatology, University of Melbourne,

Royal Melbourne Hospital, Melbourne, Australien 03-04/2006

Praktisches Jahr 1. Tertial Department of Internal Medicine, Box Hill Hospital,

Monash University, Melbourne, Australien 08-12/2006

2. Tertial Chirurgische Klinik, Stadtspital Waid, Zürich, Schweiz 12/2006-04/2007 3.Tertial Hautklinik Universitätsklinikum RWTH Aachen 04-07/2007 Forschung Dissertation Klinik für Plastische Chirurgie, Hand- und Verbrennungschirurgie,

Universitätsklinikum RWTH Aachen, Univ.-Prof. Dr. Dr. med. Prof. h.c. (RC) N. Pallua

03/2004-08/2006

Forschungstätigkeit Forschung in der Klinik für Plastische Chirurgie, Hand- und Ver-

brennungschirurgie/ IZKF BIOMAT als studentische Hilfskraft, Uni-versitätsklinikum RWTH Aachen, Univ.-Prof. Dr. Dr. med. Prof. h.c. (RC) N. Pallua

07/2005-12/2006

Forschung in der Hautklinik, Universitätsklinikum RWTH Aachen,

Prof. Dr. med. J. M. Baron 11/2005-08/2006

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Ärztliche Tätigkeit Approbation als Ärztin

Universitätsklinikum RWTH Aachen 14.11.2007

Assistenzärztin Hautklinik Universitätsklinikum RWTH Aachen, Univ.-Prof. Dr. med. H. F. Merk

Seit 11/2007

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