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Doctoral Thesis

Design of artificial microtissues

Author(s): Kelm, Jens Michael

Publication Date: 2005

Permanent Link: https://doi.org/10.3929/ethz-a-005064702

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DISS. ETH No. 16188


A dissertation submitted to the

SWISS FEDERAL INSTITUTE OF TECHNOLOGY ZURICH

For the degree of

Doctor of Natural Sciences

Presented by

JENS MICHAEL KELM

Dipl. Biotech. TU Braunschweig

born 24.06.1971

Citizen of

Germany

Accepted on the recommandation of

Prof. Dr. Martin Fussenegger, examiner

Prof. Dr. Sabine Werner, co-examiner

Zurich, 2005

Summary VII

Summary

Design of artificial microtissues (100-400 µm in diameter) by self-assembling of

monodispersed primary or neoplastic/engineered cell lines is gathering momentum in

regenerative medicine, developmental biology and the design of more reliable cell-based

drug-discovery initiatives. In this work, we have refined the hanging drop cultivation

technology, to generate artificial microtissues under controlled conditions and evaluated their

potential to (i) maintain tissue-specific functionality, (ii) produce therapeutic proteins, (iii)

provide complex feeder structures for difficult-to-differentiate cell types, iv) induce

neovascularization and (v) support integration of implants into the host tissue and vascular

network.

Hepatocytes originating from a human hepatocellular carcinoma adopted a polarized

in vivo-like morphology, expressing a number of hepatocyte-specific transcripts

involved in liver metabolism and detoxification (Chapter 2, 8).

We developed the first 3D cell-based high-throughput-compatible in culture ELISA

for VEGF profiling (Chapter 2).

Neonatal rat (NRCs) and mouse cardiomyocytes (NMCs) were assembled to

contracting myocardial microtissues, which retained cardiomyocyte-specific

morphology. For the first time, we were able to grow dissociated adult rat

cardiomyocytes (ARCs) in a 3-dimensional environment (Chapter 3).

Recombinant protein production of lentiviral transduced NRCs and NMCs could be

increased up to 6-fold in microtissue cultures, compared to their monolayer

counterparts (Chapter 3).

Self-patterning to an in vivo-like morphology could be observed in cultures of two and

more cell types according to the differential adhesion hypothesis. In mixed cultures of

human umbilical endothelial cells (HUVECs) and human fibroblasts/hepatocytes,

HUVECs enveloped the microtissues and assembled a barrier between the surrounding

liquid and the tissue (Chapter 2, 5). Sensory neurons originating from embryonic

mouse dorsal root ganglions (DRGs), mixed with mouse-derived fibroblasts and

Schwann cells segregated to the outer surface and formed ganglia-like cap structures.

Outgrowing axons were aligned with myelinating Schwann cells (Chapter 4).

Summary VIII

Microtissues of several cell types were found to express endogenous vascular

endothelial growth factor (VEGF). This was utilized to engineer vascularized

microtissues without addition of any proangiogenic factor (Chapter 5).

Myocardial microtissues placed onto the embryonic chicken chorioallantoic

membrane (CAM) integrated into the CAM and connected to the host vascular

network without supply of any proangiogenic factors (Chapter 6).

Myocardial microtissues composed of neonatal rat cardiomyocytes transplanted into

the pericardial cavity of adult rats entirely integrated within 7 days into the

myocardium (Chapter 6).

Microtissues were successfully used as building blocks to assemble larger-sized tissue

constructs (heart muscle, cartilage, connective tissue, skeletal muscle). Integrating

HUVECs considerably improved connective microtissue assembly, indicating that

endothelial cells might play an important role for tissue integrity (Chapter 6, 7).

Human fibroblast-composed macrotissues placed onto the CAM connected to the

chicken vascular network only when HUVECs had been incorporated, whereas

HUVEC-free macrotissues completely failed to integrate into the CAM (Chapter 7).

Summary IX

Zusammenfassung

Die Entwicklung von artifiziellen Mikrogeweben (100-400 µm im Durchmesser) mittels

zellulärer Reaggregation gewinnt immer mehr an Bedeutung für die regenerative Medizin, in

der Entwicklungsbiologie und der Entdeckung neuer Medikamenten durch verlässlichere Zell

basierender Untersuchungen. In dieser Arbeit haben wir artifizielle Mikrogewebe in

Tropfkulturen unter kontrollierten Bedingungen produziert und deren Potential zur i)

Erhaltung Gewebe-spezifischer Strukturen, ii) Produktion von therapeutischen Proteinen, iii)

Induzierung eines Gefäßsystems in vitro, iv) Induktion von Neovaskularisierung und v)

Integration/Anbindung an ein bestehendes Gefäßsystem untersucht.

Leberzellen, isoliert von einem Leberkarzinoma, behielten die für Hepatozyten

charakteristische polarisierte Zellmorphologie und exprimierten Leber spezifische

Transkripte die für den Leberstoffwechsel wichtig sind wesentlich stärker als

Hepatozyten in klassischen Monolayer Kulturen (Kapitel 2, 8).

Basierend auf die Produktion von Mikrogeweben haben wir einen „in culture“ ELISA

zur Detektion von VEGF entwickelt (Kapitel 2).

Neonatale Ratten und Maus Herzmuskelzellen reaggregierten zu funktionellen

Herzmuskelmikrogewebe in denen die Kardiomyozyten ihre spezifische Morphologie

aufrecht erhielten. Wir konnten zum ersten Mal zeigen, das sich adulte

Kardiomyozyten in einem 3-dimensionalen Verbund kultivieren lassen (Kapitel 3).

Protein Produktion von lentiviral transduzierten, rekombinanten, neonatalen

Kardiomyozyten, konnte in Mikrogeweben im Vergleich zu Monolayer Kulturen um

das 6-fache gesteigert werden (Kapitel 3).

Mischkulturen von zwei oder mehr Zelltypen organisierten sich selbst zu in vivo

ähnlichen Strukturen, so wie es von der differentiellen Adhäsions-Theorie beschrieben

wird. Mischkulturen von humanen Nabelschnur Endothelzellen und humanen

Fibroblasten/Hepatozyten umschlossen jeweils die Endothezellen das Mikrogewebe

und bildeten eine Barriere zwischen dem umgebenden Flüssigkeit und dem Gewebe

(Kapitel 2, 5). Sensorische Nerven isoliert von embryonalen Maus dorsalen

Wurzelganglions vermischt mit Maus Fibroblasten entmischten sich und die Neuronen

bildeten Ganglia ähnliche Strukturen. Die auswachsenden Axone waren assoziiert mit

Schwann’schen Zellen und teilweise myelinisiert (Kapitel 4).

Summary X

Mikrogewebe von verschiedenen Zelltypen produzierten endogenen vaskulären

Endothelzell Wachstumsfaktor (VEGF). Diese Eigenschaft konnte genutzt werden, um

Mikrogewebe zu vaskularisieren ohne Zugabe von proangiogenen Faktoren (Kapitel

5).

Herzmuskelmikrogewebe platziert auf der chorioallantoischen Membran eines

embryonalen Huhns, wanderte ins Membrangewebe ein und induzierte

Revaskularisierung ohne Zugabe zusätzlicher proangiogenen Faktoren (Kapitel 6).

Herzmuskelmikrogewebe welches in den Herzbeutel injiziert wurde, integrierte sich

innerhalb von sieben Tagen ins Herzmuskelgewebe des Empfängers (Kapitel 6).

Mikrogewebe ( m3 Maßstab) konnten erfolgreich als kleinste mögliche

Gewebeeinheiten verwendet werden um größere Gewebe (mm3 Maßstab) zu

generieren. Dabei hat sich herausgestellt, dass Endothelzellen die Reaggregation von

Mikrobindegewebe erheblich verbessern konnten Kapitel (6, 7).

Aus humanen Fibroblasten bestehende Makrogewebe verbanden sich mit dem

Gefäßsystem der chorioallantoischen Membran nur wenn Endothezellen mit ins

Gewebe eingearbeitet wurden. Gewebe ohne Endothezellen wurden abgestoßen und

starben ab (Kapitel 7).

I

Table of Contents

SUMMARY.......................................................................................................................................................VII

ZUSAMMENFASSUNG....................................................................................................................................IX

Chapter 1:

Impact of 3D Cell Culture Technology

2D VS 3D CELL CULTURE................................................................................................................................2

3D CELL CULTURE SYSTEMS ........................................................................................................................4

REFERENCES...................................................................................................................................................... 7

Chapter 2:

Microscale Tissue Engineering Using Gravity-Enforced Cell Assembly

ABSTRACT ......................................................................................................................................................... 11

IMPACT OF MICROTISSUE DESIGN ON REGENERATIVE MEDICINE............................................. 11

SCAFFOLD-FREE MICROTISSUES – HANGING DROP TECHNOLOGY REVISITED...................... 13

DESIGN OF ARTIFICIAL HEPATIC TISSUES............................................................................................ 16

ARTIFICIAL MYOCARDIAL MICROTISSUE ............................................................................................ 18

MICROCARTILAGE......................................................................................................................................... 20

BEYOND TISSUE ENGINEERING – THE FUTURE OF MICROTISSUES IN

BIOPHARMACEUTICAL MANUFACTURING AND HIGH THROUGHPUT DRUGDISCOVERY....22

PERSPECTIVES................................................................................................................................................. 25

ACKNOWLEDGEMENTS................................................................................................................................ 25

REFERENCES.................................................................................................................................................... 25

II

Chapter 3:

Design of Artificial Myocardial Microtissues

ABSTRACT ......................................................................................................................................................... 31

INTRODUCTION............................................................................................................................................... 32

MATERIALS AND METHODS........................................................................................................................ 34

Isolation and three-dimensional cultivation of neonatal rat and mouse cardiomyocytes.................................. 34

Video microscopy............................................................................................................................................. 35

Fluorescence-based characterization of cell morphologies............................................................................... 35

Confocal light microscopy................................................................................................................................ 36

Lentivirus-based transduction technology ........................................................................................................ 36

Quantitative expression analysis of the secreted -amylase (SAMY) ................................................................ 37

RESULTS............................................................................................................................................................. 37

Production of myocardial microtissues............................................................................................................. 37

Immunohistological characterization of myocardial microtissues.................................................................... 41

Characterization of the extracellular matrix of myocardial microtissues.......................................................... 43

Expression of vascular endothelial growth factor (VEGF) by myocardial microtissues .................................. 44

Lentiviral infection of cardiomyocytes............................................................................................................. 46

DISCUSSION ...................................................................................................................................................... 48


REFERENCES.................................................................................................................................................... 51

Chapter 4:

Self-Assembling of Sensory Neurons to Ganglia-like Structures

ABSTRACT ......................................................................................................................................................... 56

INTRODUCTION............................................................................................................................................... 56

MATERIAL AND METHODS.......................................................................................................................... 58

Isolation of mouse embryonic dorsal root ganglia (DRG) and fibroblasts ....................................................... 58

Cell culture and microtissue production ........................................................................................................... 59

Macrotissue assembly....................................................................................................................................... 59

Immunofluorescence-based cell characterization ............................................................................................. 59

Histology .......................................................................................................................................................... 60

III


Transmission Electron Microscopy .................................................................................................................. 61

Gas-inducible ifn- expression ......................................................................................................................... 61

RESULTS............................................................................................................................................................. 62

Cellular re-organization .................................................................................................................................... 62

Development of 3D neuronal structures ........................................................................................................... 63

Long-term cultivation of DRG:MEF microtissue cultures ............................................................................... 66

Assembly of innervated macrotissues............................................................................................................... 67

DISCUSSION ...................................................................................................................................................... 68


REFERENCES.................................................................................................................................................... 71

Chapter 5:

VEGF Profiling and Angiogenesis in Human Microtissues

ABSTRACT ......................................................................................................................................................... 76

INTRODUCTION............................................................................................................................................... 76

MATERIAL AND METHODS.......................................................................................................................... 78

Isolation of primary human aortic fibroblasts................................................................................................... 78

Cell culture ....................................................................................................................................................... 78

Microtissue production ..................................................................................................................................... 79

Fluorescence-based characterization of cell morphologies............................................................................... 79

Toluidine blue staining and immunohistochemistry of paraffin-embedded microtissue sections .................... 80


Transmission Electron Microscopy .................................................................................................................. 81

ELISA-based VEGF quantification .................................................................................................................. 81

RESULTS............................................................................................................................................................. 81

VEGF production profiling of human cell-derived monolayer and microtissue cultures ................................. 81

Self-organization potential of different cell phenotypes in a microtissue format ............................................. 84

VEGF profiling of microtissues assembled from different cell types............................................................... 85

Angiogenesis-based capillary formation in microtissues.................................................................................. 86

Inhibition of angiogenesis in HAF-HUVEC microtissues................................................................................ 92

DISCUSSION ...................................................................................................................................................... 92

IV

ACKNOWLEDGMENTS................................................................................................................................... 95

REFERENCES.................................................................................................................................................... 95

Chapter 6:

Improved Tissue-Transplant Fusion and Vascularization of Myocardial

Micro- and Macrotissues Implanted into Chicken Embryos and Rats

ABSTRACT ....................................................................................................................................................... 101

INTRODUCTION............................................................................................................................................. 101

MATERIAL AND METHODS........................................................................................................................ 103

Preparation of primary cells.............................................................................................................................103

Microtissue Production....................................................................................................................................104

Macrotissue Assembly.....................................................................................................................................104

Immunofluorescence analysis..........................................................................................................................104

Confocal light microscopy...............................................................................................................................105

Transmission electron microscopy ..................................................................................................................105

Microchip-based electrophysiology.................................................................................................................105

Chicken chorioallantoic membrane (CAM) assay ...........................................................................................106

Transplantation of myocardial microtissues into rat hearts .............................................................................106

RESULTS............................................................................................................................................................107

Microtissues assembled from adult cardiomyocytes .......................................................................................107

Microchip-based electrophysiologic analysis of myocardial microtissues ......................................................109

Design and neo-vascularization of higher-order macrotissues assembled from individual myocardial

microtissues .....................................................................................................................................................110

Inter-species vascularization crosstalk enables connection of myocardial microtissues to the chicken embryo

vasculature .......................................................................................................................................................113

Integration of implanted myocardial microtissues into rat hearts ....................................................................115

DISCUSSION .................................................................................................................................................... 118

ACKNOWLEDGMENTS................................................................................................................................. 120

REFERENCES.................................................................................................................................................. 120

V

Chapter 7:

Design of Custom-Shaped Vascularized Tissues Using Microtissue

Spheroids as Minimal Building Units

ABSTRACT ....................................................................................................................................................... 125

INTRODUCTION............................................................................................................................................. 125

MATERIAL AND METHODS........................................................................................................................ 127

Preparation of primary cells.............................................................................................................................127

Cell Culture......................................................................................................................................................127

Microtissue Production....................................................................................................................................127

Macrotissue Assembly.....................................................................................................................................128

Immunohistochemistry ....................................................................................................................................128

Transmission electron microscopy ..................................................................................................................129

Chicken chorioallantoic membrane (CAM) assay ...........................................................................................129

RESULTS........................................................................................................................................................... 130

Microtissue assembly to larger-sized macrotissues .........................................................................................130

Neo-vascularization of scaffold-free macrotissues ..........................................................................................134

Implantation of HMF-HUVEC macrotissues into chicken embryos ...............................................................136

DISCUSSION .................................................................................................................................................... 139

ACKNOWLEDGEMENTS.............................................................................................................................. 141

REFERENCES.................................................................................................................................................. 141

Chapter 8:

Synergies of Microtissue Design, Viral Transduction and Adjustable

Transgene Expression for Regenerative Medicine

ABSTRACT ....................................................................................................................................................... 146

INTRODUCTION............................................................................................................................................. 146

DESIGN OF ARTIFICIAL MICROTISSUES............................................................................................... 147

Directed Cell Differentiation ...........................................................................................................................149

Gene-function analysis ....................................................................................................................................150

Animal-free drug testing and drug discovery ..................................................................................................151

VIRAL TRANSDUCTION............................................................................................................................... 152

VI

Viral vectors for gene therapy .........................................................................................................................153

Gene regulation................................................................................................................................................157

Perspectives .....................................................................................................................................................158

ADJUSTABLE TRANSGENE EXPRESSION .............................................................................................. 158

Key characteristics of an ideal gene regulation system ...................................................................................159

Antibiotic-controlled gene regulation systems ................................................................................................159

Transgene control by chemically induced dimerization ..................................................................................160

Hormone-inducible gene expression................................................................................................................161

Quorum sensing-based transgene modulation .................................................................................................161

Temperature-dependent gene regulation..........................................................................................................162

Gene regulation systems in drug discovery .....................................................................................................162

Use of gene-control systems in biopharmaceutical manufacturing .................................................................163

Outlook ............................................................................................................................................................163

REFERENCES.................................................................................................................................................. 166

ACKNOWLEDGEMENTS.............................................................................................................................. 181

CURRICULUM VITAE ................................................................................................................................... 183

Chapter 1

Impact of 3D Cell Culture Technology

3D Cell Culture Technology 2

2D vs 3D Cell Culture

In vitro cultivation of mammalian cells is predominantly carried out growing cells on

adhesive cell culture surfaces as flat monolayers. However, in their natural environment cells

not only adhere to each other, but are also embedded in an extracellular matrix (ECM)

containing proteins such as collagens, intergrins, laminin, and fibronectin, which affect cell

shape (Goldmann 2002), polarity (Boudreau 2003), tension (Tarone et al. 2000),

differentiation (Bokel et al. 2002) and help to organize communication between the cells

(Schenk et al. 2003) (Figure 1). Local disruption of ECM by pharmacologic or genetic means

results in selective programmed cell death (apoptosis) among adjacent cells (Boudreau et al.

1995). The cell shape of endothelial cells control whether individual cells grow or die,

representing a fundamental mechanism for cell fate regulation within a tissue environment

(Chen et al. 1997). Given this complex mechanical and biochemical interplay of cells in a

tissue, it is no surprise that cells grown in flat monolayers miss biological subtleties (Abbott

2003).

3D structures and interaction between cells and their microenvironment are already

essential in the earliest stages of embryonic development for organization, differentiation and

proliferation (Mathis et al. 2002). For example, the requisite step of cellular condensation

during mesenchymal chondrogenesis is mimicked in vitro in chondrocyte cultures where high

cell densities results in the formation of 3D spheroid structures that are cartilaginous in nature

and associated with the upregulation of ECM components such as type II collagen and

cartilage link protein (Denker et al. 1995). Studies of embryonic chick (calvarial or limb-bud)

cells also confirm the cell density-mediated induction of chondrogenesis (Wong et al. 1995;

Woodward et al. 1999) and demonstrate requirement for cell-cell interactions in this process

(Woodward et al. 1999). Bone development requires the concerted action of several

microenvironmental signals. During osteogenesis cells differentiate into pre-osteoblasts and

then undergo cellular condensation, a process, which precedes osteoblast differentiation and

matrix mineralization (Dunlop et al. 1995). The similarity between these systems of chondro-

and osteogenesis and their concordance with similar processes in the developing embryo

strongly suggest that cell organization into 3D structures is essential for ex vivo tissue

formation (Kale et al. 2000).

Due to the lack of 2-dimensional (2D) cell culture technologies to display tissue-like

phenotypes, biologists are turning more and more to 3-dimensional (3D) cell culture


technologies. Radiation biologists have used multicellular tumor spheroids (MCTS) for

around 25 years and their utility is now receiving a wider appreciation. MCTS reproduce the

tumor microenvironment more accurately than 2D cultures (Sutherland 1988; Mueller-Klieser

1997; Hamilton 1998; Kunz-Schughart et al. 1998; Desoize et al. 2000), which has profound

implications for tumor biology, particularly with regard to altered gene expression and

sensitivity to chemotherapeutic agents (multicellular resistance) (Dubessy et al. 2000). Cancer

cells have shown to respond differently to anticancer drugs in 2D and 3D configuration.

Breast cancer cells treated with antibodies against the cell surface receptor 1-integrin

changed their abnormal shape and growth behaviour in 3D culture whereas the effect couldn’t

be observed in monolayer cultures (Weaver et al. 1997). Jacks and Weinberg went as far as to

quote the study of cancer cells in monolayers without including their ECM environment and

neighbouring cells as quaint if not archaic (Jacks et al. 2002) (see also Figure 1).

For tissue engineering initiatives, coaxing cells to form artificial tissues in a reliable

manner is the quintessential engineering design problem that must be accomplished. Tissue

engineering exploits living cells in a variety of ways to restore, maintain or enhance tissues

and organs. It conjures up visions of organs built from scratch in the laboratory with the

potential impact to reduce the need for organ replacement and accelerate the development of

new drugs. Cell-based testing is well established in drug discovery with well-described

models that exist for cancer (Johnson et al. 2001), intestinal absorption (Le Ferrec et al. 2001)

and diabetes (Reed et al. 1999). A cell-based model that is faithful to its in vivo behaviour

offers obvious advantages, such as predictability, savings of time and cost. However, current

models fall short of this ideal (Bhadriraju et al. 2002). Even genetically normal cells, such as

hepatocytes or endothelial cells placed into in cell culture quickly loose their differentiated

gene expression pattern and phenotype (Berthiaume et al. 1996; Kelm et al. 2004). For

example, the hepatitis C virus has infected more than 170 million people worldwide, but

infecting liver cells in vitro is extremely difficult as human hepatocytes quickly loose their

susceptibility to viral infection. In vitro engineered liver tissue may provide a cheaper system

with better control of variables for studying viral infection compared to animal model systems

(Griffith et al. 2002). The more closely in vitro models mimic the morphology and

biochemical processes in the body the more it will allow researchers to reduce the use of

experimental animals even if it will never replace in vivo trials.


Figure 1 Implication of 3 dimensional shape , tension , external factors and extracellular matrix (ECM)

on migration, differentiation, proliferation, homeostasis, and apoptosis (adapted from Vogel et al. 2003).

3D Cell Culture Systems

Seeding scaffolds, on which cells can re-establish their 3D structure, is currently the

standard technology. An important part in scaffold-based tissue engineering strategies is taken

by the biomaterials used for the scaffolds. They serve as substrate on which cell populations

can attach and migrate, be implanted with combinations of specific cell types, as a cell

delivery vehicle and be utilized as a drug carrier to activate specific cellular functions in a

localized region (Shin et al. 2003). Four types of biomaterials are used as scaffold material for

tissue engineering applications: (i) synthetic organic materials (such as aliphatic polyesters,

polyethylene glycol), (ii) synthetic inorganic scaffold material (such as hydroxyapatite,

tricalciumphosphate, glass ceramics), (iii) organic materials of natural origin (such as

collagen, fibrin glue, matrigel) and (iv) inorganic materials of natural origin (such as coralline

hydroxyapatite) (Hutmacher 2001; Vats et al. 2003). However, such matrices bear biological

External factors

ECM

Redistribution of

transmembrane receptors

Reconfiguration of

cytoskeleton

Change of physical

tension

Remodeling of ECM

Differential

gene

expression

Protein

secretion

Alternate

signaling

pathways

ApoptosisMigration

Homeostasis Differentiation

Cell Cycle

Proliferation

Physical Tension


information and elicit biological response, which might differ from the response found in the

natural microenvironment (Hunziker 1999). The inability of biomaterial scaffolds to

functionally integrate into surrounding tissue is one of the major roadblocks to developing

new biomaterials and tissue-engineering scaffolds (Vogel et al. 2003). Third generation

biomaterials (biomimetic materials) are capable of eliciting specific cellular response and

directing new tissue formation mediated by specific interactions which can be manipulated by

altering design parameters instead of non-specifically adsorbed ECM proteins (Hench et al.

2002). Despite considerable advances, current approaches to engineering cell-surface

interactions fall short in mimicking the complexity of signals through which surrounding

tissue regulates cell behavior such as induction of angiogenesis (Vogel et al. 2003). However,

one may ask why cells originating from a tissue environment should need a scaffold to rebuild

a tissue-like community.

Figure 2 Illustration of a scaffold without (A) and seeded with cells (B) (photographed by Carnegie

Mellon University, Bone Tissue Engineering Initiative)

An alternative to scaffold-based concepts is the cellular reaggregation, an attempt to

achieve a more or less complete regeneration of tissues from dispersed cells of a particular

origin under controlled conditions (Chapter 2). In contrast to the use of scaffolds, there is only

marginal experience in tissue engineering concerning cellular viability and integration into

host tissue of cellular reaggregates but in general it will reduce post-implantational side

effects to a solely cell-based problem. At least good integration of chondrocytes cultured as

spheroids on human condyle cartilage has been observed so far (Anderer et al. 2002). One of

the principle constraints of the size of tissues engineered in vitro that do not have their own

blood supply is the short distance over which oxygen can diffuse before being consumed

(Griffith et al. 2002). To control the cell number/composition reducing the tissue size to a

minimum, we made use of the hanging drop technology to accumulate cells into a tissue-like

environment (Figure 3, 4). This study exploits the potential of smallest possible tissue units


(microtissues) assembled in hanging drops for (i) tissue generation, (ii) cellular organization,

(iii) angiogenic properties and (iv) microtissue graft integration.

Figure 3 Gravity enforced assembly of microtissues in hanging drops. Droplets of a single cell

suspension are placed onto a surface and cultivated upside down. After 1-4 days depending on the cell type, cells

forme a cellular reaggregate

Figure 4 Generation of a multilayer micortissue configuration. After accumulation of a feeder spheroid,

a second cell type of interest is added into the hanging drop. The added cells form an additional cell layer around

the feeder spheroid (Chapter 2, 4).

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Chapter 2

Microscale Tissue Engineering Using

Gravity-Enforced Cell Assembly

Kelm J.M. and Fussenegger M., (2004) Trends in Biotechnology 22, 201-214

Microscale Tissue Engineering 11

Abstract

Design of artificial microtissues by reaggregation of monodispersed primary cells or

neoplastic/engineered cell lines are gathering momentum in regenerative medicine and

provide insight into the third dimension of cell-cell interactions and underlying regulatory

networks. Recent advances in microtissue production have substantiated the potential of

scaffold-free cell aggregates to (i) maintain tissue-specific functionality, (ii) support seamless

integration of implants into host tissues, (iii) provide complex feeder structures for difficult-

to-differentiate cell types, (iv) be amenable to therapeutic and phenotype-modulating

interventions using latest-generation transduction technologies, (v) produce therapeutic

transgenes at increased levels and (vi) offer tissue-like assay environments to improve drug-

function correlations in current discovery programs. Focusing on liver (liver-specific

detoxification characteristics), heart (interconnection of contractile units) and cartilage

(mechanical properties) we cover the latest on scaffold-free microtissue design.

Impact of microtissue design on regenerative medicine

To some extent, regeneration takes place in the human body throughout life. For

example, blood and skin are continually restored while liver, bone, muscle and blood vessels

have a limited capacity for self-repair (Petit-Zeman 2001). Yet, after traumatic injury or (age-

related) disease including osteoporosis, diabetes, cardiovascular and neurodegenerative

disorders, extensive tissue damage/degeneration exceeds the tissue-encoded repair capacity

resulting in the formation of functionally impaired scar tissue. Irreparably damaged tissue

may either be replaced by medical devices or organ (xeno-) transplantation. However,

medical devices often lack durability and an extensive functional repertoire, donor organs are

in limited supply and ongoing concerns about provirus dissemination and hyperacute rejection

reactions limit the scope of current xenotransplantation protocols (Bouche 2002).

Since most current pharmaceutical interventions may at best retard but not revert

pathologic tissue degeneration, they could only be considered stopgaps as tissue engineering-

based regenerative medicine moves from the realms of science fiction to de novo creation of

artificial organs. As natural organ development is characterized by complex processes

orchestrating assembly of different cell types and integrating a near-infinite number of signals

in space and time design of artificial organs appears like a mission impossible. Until a


system’s view on organs will be available, an optimal balance of harnessing cellular self-

repair programs combined with specific pharmacologic/genetic interventions will dominate

the clinical reality in the immediate future.

Unlimited supply of generic therapeutic cell phenotypes resulting from expansion of

desired primary cells or rational differentiation of multipotent (stem) cells by growth factors

or genetic interventions, managing shape and function, timely coordination of multi-cell type

assembly as well as mastering neoplastic cell expansion are among current challenges of

tissue engineers. Capitalizing on recent pharmacologic advances, proliferation-modulating

factors have become a cornerstone of tissue engineering either to expand cells for implant

production or to functionalize biomaterials for tissue regeneration (Lutolf et al. 2003).

However, the risk of eliciting neoplastic cell characteristics following extended expansion

procedures remains imminent. Positive proliferation control to enable expansion as well as

precise growth arrest to prevent neoplastic outgrowth following implantations are vital issues

which will require increased attention (Fussenegger et al. 1998; Fux in press).

Although expanded (stem) cell populations of desired phenotypes have often been

reported to bear significant therapeutic potential following direct injection into tissue lesions

(heart, cartilage), such therapeutic scenarios lack the scope associated with tissue implants

(Mangi et al. 2003; Rafii et al. 2003). Shaping functional 3D tissue from monodispersed

expanded cell cultures is an ongoing challenge (Abbott 2003). Tissue engineers have thus

relied on material science to provide (functionalized) scaffolds on which tissue cells may

grow and differentiate. Latest-generation scaffolds are branched to enable adequate feeding of

cells in the central layers (Kim et al. 1998; Zandonella 2003), provide biological and

mechanical functions of a native extracellular matrix (Kim et al. 1998), degrade once the

organ or tissue becomes established in the body, and may be designed to release growth

factors or transgene-encoding vectors in response to physiological cues (Lee et al. 2000;

Richardson et al. 2001; Lutolf et al. 2003). Although scaffolds offer unique clinical

opportunities in tissue engineering strategies which require a strict combination of shape and

function (e.g., bladder, bone, cartilage, intestine/stomach, liver, skin) shape-supporting

matrices could be expendable or even less suited for the design of brain and heart structures

(Ochoa et al. 2002) (Table 1). Exemplified by recent advances in the production of artificial

liver, heart and cartilage structures we are covering the latest trends in designing scaffold-free

artificial microtissues.


Table 1 Potential advances of gravity-enforced microtissue design

1) Precise tissue size control owing to a strict correlation between cell number and spheroid

diameter to avoid oxygen and nutrient limitations of the in vitro culture (Kelm et al.

2003).

2) The ease to generate microtissues from different cell phenotypes mimicking natural cell-

type composition (Itskovitz-Eldor et al. 2000).

3) Cell mobility during assembly ensuring intercellular organization including polarization

(Rothermel et al. 2001).

4) Development of an extracellular matrix (Anderer et al. 2002).

5) Compatibility with high-throughput assay systems as well as robotic liquid handling

devices (Layer et al. 1992).

6) Applicability to small volumes and cell numbers (Layer et al. 2002).

7) Mild and natural assembly forces unlikely to interfere with cell regulatory networks.

8) Compatibility with a wide variety of cell types (see Table 2).

Scaffold-free microtissues – hanging drop technology revisited

Strategies harnessing the natural reaggregation potential to assemble monodispersed

cells in a tissue-mimicking manner represent a valuable extension of current scaffold-based

tissue engineering initiatives. Scaffold-free reaggregation of cells to microtissues may occur

following (i) cultivation in shake flasks, gyratory shakers and roller bottles (Furukawa et al.

2001; Kelm et al. 2003; Kelm et al. in press) or on non-adhesive surfaces (Kale et al. 2000),

(ii) centrifugation-based compression (Muraglia et al. 2003), (iii) maintenance in cell culture

inserts (Watzka et al. 2000), or (iv) gravity-enforced assembly of microspheres in hanging

drops (Kelm et al. in press) (Tables 2 and 3).

Originally pioneered for production of embryoid bodies and blastocysts to study the

differentiation potential of stem cells (Wobus et al. 2000), gravity-enforced assembly of

microtissues in hanging drops was found to be compatible with a variety of cell types and

became increasingly popular among tissue engineers (i) to assess tumor-related resistance to

chemotherapeutics in tissue-like cancer models (Bjerkvig et al. 1997; Kunz-Schughart 1999),

(ii) for gene-function analysis of differentiation phenomena and development (Itskovitz-Eldor

et al. 2000) and (iii) the design of functional microlivers, microhearts and microcartilage

(Kelm et al. 2003; Kelm et al. 2003; Kelm et al. in press) (Figure 1; Tables 2 and 3).


Figure 1: Microtissues produced by gravity-enforced assembly of monodispersed cells of a single cell

type. Phase-contrast micrographs of microtissues reaggregated from human aortic fibroblasts (A), human dermal

fibroblasts (B), neonatal rat cardiomyocytes (C), and primary rat hepatocytes (D). (scale bar = 20 µm)

A key benefit of gravity-enforced microtissue design is the mobility of cells during

microtissue formation. Tissues consist of an organized assembly of several cell types a fact,

which should be considered for microtissue design. Although forces orchestrating proper

positioning of different cell types within a tissue remain elusive, cell movements during

development or following implantation of (stem) cells are well established (Brazelton et al.

2000; Clarke et al. 2000). Design of organotypic structures will require detailed understanding

of intercellular crosstalk in cocultures.

In order to get insight into organotypic positioning effects of different cell types, we

have evaluated gravity-enforced microtissue assembly from cocultures of (i) HepG2-HUVEC

(human umbilical vein endothelial cells), (ii) HAF (human aortic fibroblast)-HUVEC as well

as (iii) rat heart-derived cell mixtures. Although completely mixed following seeding in

hanging drops, HUVEC cells always move to the periphery which is reminiscent of the

concentric structures shaped during vascularization (Kelm et al., unpublished; Figure 2A and

B). Also, when cell mixtures reflecting the natural cell-type composition of rat hearts are

cultivated in hanging drops, muscle-specific cell phenotypes were predominantly found at the

periphery of beating microtissues (Kelm et al. in press) (Figure 2C;

http://www.biotech.biol.ethz.ch/martinf/staff/jens.html). These findings exemplify the

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compatibility of gravity-enforced microtissue design with the forces shaping organotypic

structures.

Microtissues could also be used as feeder spheroids, which provide a cell-based 3D

matrix for organotypic microcultivation of difficult-to-study cell types or polarized assembly

of different cell layers (Kelm et al. in press). Primary human keratinocytes (endothelial cells)

assembled tight and seamless cell layers when coated onto fibroblast (hepatocyte)-derived

feeder microtissues in hanging drop cultures (Figure 2C-E).

Owing to the wide variety of different tissues, their in vitro production and

engineering strategies, we prefer to focus on microscale tissue engineering initiatives

currently developed to design artificial (i) hepatic tissue (maintaining liver-specific

detoxification and metabolism), (ii) functional myocardial microtissues beating at human-

compatible frequencies (maintenance of contractile units and specialized cell-cell contacts)

and (iii) cartilage (managing mechanical properties).

Figure 2: Confocal analysis of organotypic microtissue structures assembled from two different cell

types by gravity-enforced reaggregation in hanging drops. Microtissues produced by monodispersed cell

A

D

B

C

E F

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EE FF


mixtures (A-C). Hepatocyte (HepG2) – human endothelial cell (HUVEC) (A), human aortic fibroblast (HAF) –

HUVEC (B) mixed cardiomyocyte populations reflecting the natural cell-type composition of the myocard (C)

were cultivated in hanging drops. The entire microtissue was visualized using F-actin-specific staining (red) and

HUVECs were stained for von Willebrand factor (vWF) (green). Cardiomyocytes were monitored by using a

sarcomeric alpha-actinin-specific antibody (green). Premanufactured feeder spheroids were coated with a second

cell type by cultivation in hanging drops (D-F). HepG2 (D) and HAF (E) feeder spheroids were coated with

HUVECs and normal human dermal fibroblasts (NHDF) were coated with keratinocytes (HaCaT) (F). Feeder

tissues were stained for F-Actin (red), HUVECs for vWF (green), HaCaTs for keratin (green). (scale bar =

10 µm)

Design of artificial hepatic tissues

The healthy liver is able to regenerate after injury. However, once damaged by fibrosis

and cirrhosis, resulting from a variety of chronic conditions including alcohol abuse or

infection with hepatitis virus B or C, the liver’s regeneration capacity is compromised. Liver

transplantation is a routine treatment for end-stage liver disease, yet donor organ shortage

continues to be a serious problem (Strain et al. 2002). The liver has a panoply of crucial

functions including production of clotting factors, bile, the regulation of carbohydrate, fat and

protein metabolism, detoxification and breakdown of alcohol and drugs. Most of these

functions are carried out by hepatocytes, which constitute up to 70% of a liver’s cellular

content. Although hepatocytes appear to be the prime candidate for the design of artificial

hepatic tissues, they rapidly lose liver-specific gene expression and become phenotypically

unstable following removal from the complex architecture of the liver (Berthiaume et al.

1996). However, in vitro cultivation of hepatocytes will be key to produce artificial hepatic

tissues for replacement therapies and extracorporal liver-assist devices. Progress in

maintaining the liver-specific phenotype of hepatocytes has been achieved by modulating

tissue culture conditions, providing an extracellular matrix and cocultivating hepatocytes with

other liver cell types (Bhatia et al. 1999; Harimoto et al. 2002).

A variety of different strategies have been pioneered to arrange hepatocytes in 3D

configurations which exhibit epithelial polarization and retain some liver-specific functions

(Hench et al. 2002): (i) Seeding onto preformed matrices (Ambrosino et al. 2003), (ii)

cultivation in soluble matrices (Kamihira et al. 1997), (iii) stacking of monolayers (Harimoto

et al. 2002) and (iv) reaggreagation of monodispersed cells (Kelm et al. 2003) (Tables 2 and

3). Ultrastructural analysis of hepatic HepG2-derived microtissues produced following

gravity-enforced reaggregation in hanging drops revealed seamless integration of single cells

into a compact microliver-like structure (Figure 3A). Individual hepatocytes were embedded


in an extensive collagen-containing extracellular matrix (Figure 3B). Transmission electron

micrographs of microliver crossections showed cubic and polarized hepatocytes characterized

by the presence of bile canaliculi-like structures which are known to promote bile secretion in

the liver (Kelm et al. 2003) (Figure 3C). Besides adopting liver-like cell phenotypes and

structures, detailed expression profiling of hepatocytes assembled by gravity-enforced

reaggregation demonstrated increased production of the detoxifying proteome compared to

isogenic 2D monolayer cultures (Kelm et al. 2003). Managing and harnessing the

detoxification potential of liver cells is a key advancement for the design of artificial liver-like

tissues as well as extracorporal liver-assist devices.

Figure 3: Electron micrographs of HepG2-derived microliver tissues. Scanning electron micrograph of

reaggregated HepG2 spheroids at different magnifications (A, B). Transmission electron microscopy of liver

C

A B

C

A BA B


spheroids revealed intact cubic cells showing hepatocyte-characteristic polarity exemplified by bile canaliculi-

like structures (C).

Artificial myocardial microtissue

Heart diseases including myocardial infarction and heart failures are the most

prevalent pathologies in industrialized countries. Loss of cardiomyocytes accounts for

decreased myocardial function, which may result in total organ failure or trigger

compensatory mechanisms like hypertrophy of the remaining myocardium, activation of

neurohumoral systems and/or autokrine/parakrine stimulation by various growth

factors/cytokines (Zimmermann et al. 2003). The ultimate treatment of end-stage heart failure

remains heart transplantation. However, since available donor organs do not match the

increasing number of patients with heart failures, alternative strategies for restoration of heart

function are a current clinical priority (Miniati et al. 2002). Implantation of functional

cardiomyocytes and other cell types including stem cells has been shown to improve

contractile function in myocardial infarction models. Yet, ongoing clinical trials will have to

confirm the therapeutic impact of these strategies (Barbash et al. 2003; Leobon et al. 2003;

Mangi et al. 2003). An alternative to infusion of single-cell suspensions is the design of

artificial cardiac muscle tissues ex vivo followed by implantation into the diseased heart.

The prevailing 3D cultivation technology in cardiac tissue engineering will have to

unite several key characteristics including (i) long-term maintenance of the contractive

capacity, (ii) multi-cell type cultivation, (iii) potential for self-organization, polarization and

microstructure formation between different cell types, (iv) production of an extracellular

matrix, (v) vascularization, including induction of vascular vessel development and

connection to the host capillary system following implantation, (vi) development of seamless

inter-tissue superstructures and (vii) compatibility with high-efficiency stable gene transfer

technologies to enable cell phenotype-modulating and/or therapeutic interventions. Several

strategies have been developed to produce engineered cardiac tissues including rigid/soluble

matrix-based approaches (Akins et al. 1999; Polonchuk et al. 2000; Teebken et al. 2002)

(particularly successful for shaping heart valves and vessels) and scaffold-free initiatives

(Tables 2 and 3).

We have recently used gravity-enforced reaggregation of pure primary rat and mouse

cardiomyocytes as well as mixed cell populations reflecting the cell type composition of

rodent hearts to design beating heart microstructures ((Kelm et al. in press);


http://www.biotech.biol.ethz.ch/martinf/staff/jens.html). Interestingly, cardiomyocytes

expressed a high degree of organotypic heart tissue phenotypes when arranged in such a

scaffold-free 3D environment. Phenotypic characterization combined with detailed analysis of

muscle-specific cell traits, extracellular matrix components as well as endogenous VEGF

(vascular endothelial growth factor) expression profiles of heart microtissues revealed (i) a

direct cell number - microtissue size correlation (up to 320 m), (ii) inter-microtissue

superstructures, (iii) retention of key cardiomyocyte-specific cell qualities, (iv) a sophisticated

extracellular matrix, (v) a high degree of self-organization exemplified by the tendency of

muscle structures to assemble at the periphery of these myocardial spheroids (Figure 2C) and

(vi) high lentiviral transduction rates for genetic engineering of microhearts. (vii)

Furthermore, myocardial spheroids supported endogenous VEGF expression in a size-

dependent manner, which will likely promote vascularization of heart microtissues produced

from defined cell mixtures, as well as enable connection to the host vascular system following

implantation. Retention of heart-like rod-shaped cardiomyocytes was particularly prominent

when cardiomyocytes were coated onto myofibroblast feeder spheroids. This observation

exemplifies the power of 3D feeder structures for induction and maintenance of specific cell

shapes and phenotypes (Figure 4) (Kelm et al. in press).

Which one of the different myocardial microtissue design concepts will prevail in

future therapies or whether (stem) cell transplantations will succeed remains to be seen in

current clinical trials.


Figure 4: Rat heart fibroblast (RHF) feeder spheroids coated with neonatal rat cardiomyocytes (NRC).

Cardiomyocytes were stained for sarcomeric alpha-actinin (green) while the entire microtissue is visualized by

beta-catenin (red) (A, crossection; B and C, 3D projections). At higher magnification the on-top view of

microtissues reveals that the cardiomyocytes at the periphery exhibit heart-like phenotypes exemplified by their

rod shape and development of intercalated discs between individual cardiomyocytes. (scale bar = 10 µm)

Microcartilage

Osteoarthritis and rheumatoid arthritis, the most prevalent disorders of the

musculoskeletal system, result from disturbance of tissue homeostasis in articular joints and

are diagnosed by joint pain, tenderness, movement limitations, as well as effusion and

variable degrees of inflammation. Rheumatoid arthritis is characterized by chondrocytes

producing inflammatory signals and matrix metalloproteinases, which result in thinning of the

collagen network, decrease of proteoglycan aggregates and reduction of biomechanical

resistance. By contrast, osteoarthritis results from dysregulation of tissue turnover in the

weight-bearing articular cartilage and subchondral bone (Aigner et al. 2002).

Like the myocardium hyaline cartilage is devoid of any self-repair or regeneration

capacity. Since long-term evaluation of conventional surgical interventions for the treatment

of osteoarthritis including joint resurfacing (abrasion, drilling, debridements, microfracture

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techniques or arthroscopic shaving) were of limited success (insufficient repair resulting in

the formation of inadequate resident fibrocartilage), cell-based cartilage regeneration came

into the limelight of current arthritis therapies. As one of the first clinically licensed cell-

based therapies, the Food and Drug Administration (FDA; http://www.fda.gov) approved the

biological production known as autologous chondrocyte implantation (ACI) in 1997 (FDA

Reference No: 96-0372). ACI was based on enzymatic disintegration of cartilage biopsies

followed by selective expansion of chondrocytes and reimplantation into the damaged

cartilage. Unfortunately, chondrocytes lost their differentiated phenotype during in vitro

expansion since the cells were cultured on an inappropriate substrate owing to lack the

requisite characteristic extracellular matrix environment. Although many exercising surgeons

emphasize the good to excellent clinical results (60%-90% of the patients report pain relief

and improved joint functionality), fate and redifferentiation of autologous chondrocyte

implants remain elusive (Hunziker 2002). ACI implants are typically fixed to the defect joint

surface using fibrin glue or resorbable pins but only 8% of the implanted chondrocytes could

be identified within the repaired tissue (Grande et al. 1989). Also, management of a solid

connection between cartilage and bone tissues remains a current clinical focus.

In contrast to ACI-based monolayer cultures, chondrocytes retain their differentiation

status in a 3D environment typically provided by scaffolds supporting initial mechanical

stability and even cell distribution (Risbud et al. 2002). To date, there are only few scaffold-

free approaches available for 3D cultivation/assembly of primary chondrocytes to artificial

cartilage (Tables 2 and 3). Fixation of artificial microcartilage onto native cartilage resulted in

rapid formation of focal adhesion points and seamless integration of the microtissues into the

target cartilage. 3 weeks post fixation, microcartilage-derived cells were located on the native

cartilage surface and showed de novo synthesis of extracellular matrix (Anderer et al. 2002).

Immuno-based confocal analysis of microcartilage (produced by gravity-enforced assembly

of primary human and pig chondrocytes) in hanging drops revealed two distinct cell layers, an

outer one qualified by fibroblast-like cell morphologies and an inner core consisting of a loose

assembly of tubular chondrocytes embedded into an extracellular matrix (Figure 5 C, D; Kelm

et al., unpublished).

Successful therapies for repair and regeneration of cartilage will require assembly of

chondrocytes in a 3D microcartilage configuration compatible with surgical implantation and

fixation, genetic engineering, optimal differentiation and production of an extracellular

matrix. Although gravity-enforced production of scaffold-free microcartilage is expected to

meet with these criteria at a high standard clinical confirmation remains imminent.


Figure 5: Microcartilage produced by gravity-enforced reaggregation of pig and human chrondrocytes

in hanging drops. Phase-contrast micrographs of microcartilage produced from 1200 pig (A) and human (B)

articular chondrocytes. F-Actin-specific staining of pig (C) and human (D) chondrocyte derived microtissues

illustrates the morphological structure of the microcartilage. (scale bar = 20 µm)

Beyond tissue engineering – the future of microtissues in

biopharmaceutical manufacturing and high throughput drug

discovery

Monodispersed cells growing in suspension and protein-free media are currently the

golden standard for large-scale manufacturing of protein therapeutics (Chu et al. 2001).

Although key biopharmaceutical manufacturing parameters such as growth rate, cell density

and specific productivity can be optimized by advanced bioprocess control or specific

molecular interventions in production cell lines, the question remains, whether specific

productivity of suspension cells typically reached in classical bioreactor operation compare

favourably with nature’s well-evolved protein production capacity.

Recent evidence suggested that cells cultivated in microtissues, embedded in a tissue-

like 3D environment or engineered for proliferation-arrested terminal differentiation reach

higher specific productivities compared to proliferation-competent monodispersed suspension

cultures: (i) reprogramming of CHO-K1-, DG44- and NSO-derived cell lines for

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proliferation-controlled terminal differentiation by conditional overexpression of the cyclin-

dependent kinase inhibitors p21Cip1

and p27Kip1

and/or the CAAT/enhancer-binding protein

alpha (C/EBP ) significantly increased specific productivity of (model) product proteins by

up to one order of magnitude (Fussenegger et al. 1998; Meents et al. 2002; Ibarra et al. 2003).

(ii) Encapsulation of mammalian production cell lines in biocompatible hydrogels including

alginate or sodium cellulose sulphate - poly[diallyldimethylammonium chloride]

(PDADMAC) copolymers, results in high-density microspheres showing increased

production compared to isogenic 2D cultures (Weber et al., unpublished). (iii)

Cardiomyocyte-derived microtissues transduced with a secreted reporter gene-encoding

lentivirus produced 6-fold more heterologous protein than control monolayers consisting of

identical cell numbers (Kelm et al. in press). While controlled proliferation technology is

ready for industrial application, microtissue-based biopharmaceutical manufacturing is

currently in an up-scale process. Yet, hardware for industrial manufacturing of

microencapsulated production cell lines is available (Gugerli et al. 2002; Koch et al. 2003)

and gravity-enforced design of microtissues is compatible with standard liquid-handling

robotics.

In addition to provide tissue-mimicking clinical implants or high-performance

aggregates for biopharmaceutical manufacturing, microtissues may reveal a systems’ view on

tissue formation, drug testing and drug discovery. Reaggregated microtissues of tumor cells

have long played a pivotal role in cancer research since multicellular tumor spheroids show

increased proliferative activity and drug resistance similar to solid tumors (Kelm et al. 2003).

Microtumors enable precise analysis of growth constraints (e.g. oxygen and nutrient

consumptions), sensitivities to drugs or radiation, infiltration into non-cancerous tissues as

well as the angiogenic potential. Microtissues are becoming increasingly popular as models

for neurodegenerative disorders toxicology, pharmacology, nutrition and environmental

monitoring (Bhadriraju et al. 2002; Layer et al. 2002).

Drug discovery and diagnostics typically include screening of chemical or metabolic

libraries for therapeutic compounds (antigens, antibodies, nucleotides and peptides) in a

microscale format. Replacing 2D cultures used in classical drug discovery by microtissues

may (i) increase the precision of therapeutic readout, (ii) enable early drug validation at the

tissue level in a cost-efficient animal-free setting and (iii) expand the discovery window into a

yet unknown dimension. Figure 6 exemplifies a prototype high-throughput-compatible ELISA

format which combines target molecule quantification (e.g., vascular endothelial growth

factor; VEGF) with microtissue design (e.g., human aortic fibroblasts, HAF). Multiwell plates


were coated with a VEGF-specific capture antibody and individual wells incubated with

different HAF cell concentrations (500, 2500, 5000, 10000 cells per well). For seven days,

HAF-derived microtissues of different sizes were produced by gravity-enforced reaggreation

in hanging drops. Hypoxia-induced VEGF production correlated directly with the size/cell

number of HAF-derived microtissues and could be quantified using an ELISA-type protocol.

Microtissues are thus compatible with high-throughput profiling of desired proteins.

Figure 6: Microtissue-based high-throughput ELISA-type assay for quantification of desired proteins.

(i) Individual wells of a multiwell plate were coated with a capture antibody specific for the vascular endothelial

growth factor (VEGF) capture antibody. (ii) Monodispersed cell suspensions of human aortic fibroblasts (HAF)

were seeded at various concentrations into each well and the plates were cultivated upside down to enable

formation of hanging drops and gravity-enforced microtissue production. (iii) Microtissues were discarded and

medium removed to quantify microtissue-based VEGF production using an ELISA-type protocol and a

chromogenic readout. (iv) Owing to the direct microtissue size – cell number correlation hypoxia-induced VEGF

production increases with the microtissue diameter.

Coating with the

Capture Antibody

Cultivation of

Microtissues in

Hanging Drops

Developing the

ELISA

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Coating with the

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Cultivation of

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Developing the

ELISA

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Perspectives

“We are a self-assembling organism. That information is there to be captured and

used” says William Haseltine, chairman and chief executive officer (CEO) of Human

Genome Sciences (Rockville, MD) who was among the first to coin the term “regenerative

medicine” to describe new ways of teaching the body to heal itself (Wade 2001). Microtissue

design consisting of reagreggation of biopsy-derived (stem) cell populations or (stem) cell

lines harnesses the organism’s self-assembling programs to provide regenerative medicine

initiatives with clinical tissue implants and new insight into regulatory networks underlying

complex disease phenotypes. “Further in the future”, Haseltine says, “biologists may learn

how to fashion new organs outside the body” (Wade 2000). In that sense, the future has just

begun with the design of artificial microtissues.

Acknowledgements

We are grateful Lars K. Nielsen for providing Figure 3 micrographs. We thank David

Fluri, Beat P. Kramer and Shizuka Hartenbach for critical comments on the manuscript. Work

in the laboratory of M.F. is supported by the Swiss National Science Foundation (grant no.

631-065946).

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Chapter 3

Design of Artificial Myocardial Microtissues

Kelm J.M., Ehler E., Nielsen L.K., Schlatter S., Perriard J.C. and Fussenegger M., (2004)

Tissue Engineering 10, 201-207

Myocardial Microtissue 31

Abstract

Cultivation technologies promoting organization of mammalian cells in three

dimensions are essential for gene-function analyses as well as drug testing and represent the

first step towards the design of tissue replacements and bioartificial organs. Embedded in a

three-dimensional environment, cells are expected to develop tissue-like higher order

intercellular structures (cell-cell contacts, extracellular matrix) which orchestrate cellular

functions including proliferation, differentiation, apoptosis and angiogenesis with unmatched

quality. We have refined the hanging drop cultivation technology to pioneer beating heart

microtissues derived from pure primary rat and mouse cardiomyocyte cultures as well as

mixed populations reflecting the cell type composition of rodent hearts. Phenotypic

characterization combined with detailed analysis of muscle-specific cell traits, extracellular

matrix components as well as endogenous VEGF (vascular endothelial growth factor)

expression profiles of heart microtissues revealed (i) a linear cell number - microtissue size

correlation, (ii) inter-microtissue superstructures, (iii) retention of key cardiomyocyte-specific

cell qualities, (iv) a sophisticated extracellular matrix, and (v) a high degree of self-

organization exemplified by the tendency of muscle structures to assemble at the periphery of

these myocardial spheroids. (vi) Furthermore, myocardial spheroids support endogenous

VEGF expression in a size-dependent manner which will likely promote vascularization of

heart microtissues produced from defined cell mixtures as well as support connection to the

host vascular system following implantation. As cardiomyocytes are known to be refractory to

current transfection technologies we have designed lentivirus-based transduction strategies to

lead the way for genetic engineering of myocardial microtissues in a clinical setting.


Introduction

Cardiac-related insufficiencies are the major cause of morbidity and associated

mortality in industrialized countries affecting over sixty million patients per year in the USA

alone (Claycomb 1991; Gheorghiade et al. 1998; Hennekens 1998). Despite global initiatives

to foster advances in cardiomyocyte-related tissue engineering this cell type turned out to be

largely refractory to state-of-the-art gene transfer and expansion technologies, particularly in

its terminally differentiated adult phenotype (Datwyler et al. 2001). Yet, cardiac tissue

engineering remains a high priority as transplantations and artificial hearts only cover about

10% of current clinical needs and/or fail to offer any long-term therapeutic perspectives. Even

engineered heart cell suspensions and undersized cell aggregates may be sufficient for the

treatment of smaller heart lesions (Akins 2002).

Although studies on primary cardiomyocyte monolayer cultures produced from

enzymatically dispersed fetal, neonatal, or adult vertebrate hearts have substantiated

intracellular structure-function correlations (Rothen-Rutishauser et al. 1998) and revealed

molecular pathways shaping this complex cell phenotype, two-dimensional cultivation of

cardiomyocytes is likely limited in providing clinically relevant information required for

sophisticated tissue engineering. In contrast to monolayer cultures, three-dimensional

cultivation technologies mimic cardiac tissue-like morphologies and provide a suitable

environment for coordination of cell-cell interaction, self-organization, differentiation and

electrical properties, all of which are essential qualities for the identity and integrity of heart

structures (Eschenhagen et al. 1997; Ross et al. 2001). Also, only three-dimensional

cultivation enables development of an extracellular matrix (ECM) which plays a pivotal role

in differentiation, proliferation and apoptosis regulatory networks (Chen et al. 1997; Aplin et

al. 1999; Giancotti et al. 1999).

The majority of cardiomyocyte-adapted three-dimensional cultivation technologies are

based on artificial scaffolds consisting of (i) titanium dioxide ceramics (Polonchuk et al.

2000), (ii) cell/collagen mixtures (Eschenhagen et al. 1997), (iii) polystyrene microcarrier

beads (Akins et al. 1999), (iv) alginate polymers (Magyar et al. 2001; Dar et al. 2002), and (v)

polyglycolic acid structures maintained in microgravity bioreactors (Freed et al. 1997; Carrier

et al. 1999). Although scaffolds enable artificial organs to grow in a desired shape they may

cause post-transplantation side effects resulting from toxic degradation products, induction of

inflammatory reactions and poor resorption (Yang et al. 2001). Alternative strategies for


production of scaffold-free artificial tissues are taking advantage of the aggregation of

mammalian cells following (i) cultivation in shake flasks or on non-adhesive surfaces

(Sperelakis 1978), (ii) centrifugation-based compression (Armstrong et al. 2000), (iii)

maintenance in cell culture inserts (Watzka et al. 2000), or (iv) gravity-enforced assembly in

hanging drops (Keller 1995). To date, most attempts to shape artificial cardiac-like cell

aggregates have been based on physical assembly of isolated primary cardiomyocytes since

the emergence of cardiomyocytes in spheroid cultures of human embryonic stem cells

remains stochastic and strategies for rational differentiation of this pluripotent cell type into

cardiac cells are still in its infancy (Schuldiner et al. 2000; Boheler et al. 2002).

A major challenge in the design of (heart) microtissues is timely and well-balanced

induction of vascularization to ensure long-term supply of large artificial tissues with

oxygen/nutrients as well as their connection to the host vascular system following

implantation (Griffith et al. 2002). As a typical cell is connected to the capillary network

within an average perimeter of 60 m in vivo, oversized microtissues may show hypoxia-

/nutrient deprivation-induced cell death in their centers (Intaglietta et al. 1996). Although

localized delivery of heterologous VEGF to cardiac lesions has recently been shown to induce

vascularization and connection of transplanted myoblasts to the coronary artery, overdosing

of this growth factor was associated with the formation of hemangiomas and cell death

(Richardson et al. 2001).

In this study we have refined the hanging drop cultivation technology to design

functional heart microtissues from purified primary neonatal rat (NRC) and mouse (NMC)

cardiomyocytes and from cardiomyocyte-containing cell mixtures reflecting the natural cell

composition of neonatal rodent hearts. These myocardial microtissues could be adjusted in

size and were beating for over 3 weeks. In contrast to monolayer cultures and small-sized

spheroids, myocardial microtissues of 230 11 m in diameter secrete VEGF, which is

expected to facilitate vascularization during production of artificial tissues or following

implantation. In order to enable rational molecular interventions in myocardial microtissues to

refine cell phenotypes or to engineer production of desired protein therapeutics, we have

evaluated lentiviral transduction systems for straightforward transgene delivery into

cardiomyocytes cultivated in two or three dimensions.

We are convinced that the design as well as detailed characterization of myocardial

microtissues bundled with a powerful transduction technology will foster novel opportunities

in cardiac tissue engineering and the treatment of heart diseases.


Materials and Methods

Isolation and three-dimensional cultivation of neonatal rat and mouse

cardiomyocytes

Newborn rat (Wistar) and mouse (NMRI) hearts were dissected, digested with

collagenase (Worthington Biochemical Corp., Freehold, NJ) and pancreatin (Invitrogen,

Carlsbad, CA) and prepared as described by Auerbach et al. (1999) (Auerbach et al. 1999).

Rat cardiomyocytes were either cultivated as mixed populations reflecting the cell type

composition of rodent hearts (25% cardiomyocytes) or cultured as homogenous populations

following density gradient purification (over 95% cardiomyocytes). The mouse populations

contained up to 50% cardiomyocytes following preplating of isolated cells for 2 h in a tissue

culture dish.

Following isolation, cardiomyocyte-containing cell populations were seeded at

indicated concentrations into 60-well plates (HLA plate, Nunc Inc., Roskilde, Denmark). To

enable gravity-enforced microtissue formation in hanging drops, the 60-well plates were

incubated upside down at 37°C in a humidified atmosphere containing 5% CO2. Pure and

mixed cardiomyocyte cultures were maintained in plating medium (67% Dulbecco`s Modified

Eagle Medium (DMEM; Invitrogen), 17% M199 (Amimed AG, Basel, Switzerland), 10%

horse serum (cat. no. 16050-098, lot no. 3036354D, Invitrogen), 5% fetal bovine serum (FBS;

cat. no. A-15-022, lot no. A01129-242; PAA Laboratories, Linz, Austria), and 1%

penicillin/streptomycin solution (Invitrogen)). After cultivation for 3-4 days in hanging drops

the microtissues were harvested and kept for further analysis in non-adhesive culture dishes

containing maintenance medium (78% DMEM (Invitrogen), 20% M199 (Amimed AG), 1%

horse serum (Invitrogen), 1% penicillin/streptomycin solution (Invitrogen), and 10-4

mM

phenylephrine (Sigma Chemicals, St. Louis, MO). Cardiomyocyte monolayers were

cultivated in fibronectin-coated (10 g/ml human plasma-derived fibronectin (cat. no.

688851, lot no. 14814500, Roche Biochemicals, Basel, Switzerland)) culture dishes using

plating medium. After one day the medium was replaced by maintenance medium.

Cardiomyocyte-coated microtissues cultures were generated by producing a three-

dimensional feeder spheroid using 1’200 myocardial fibroblasts/drop in plating medium.

After 2 days 900 gradient-purified neonatal rat cardiomyocytes were added per feeder

spheroid. After an additional 4-day cultivation in hanging drops the cardiomyocyte-coated

spheroids were harvested and prepared for immunohistochemistry.


Video microscopy

Phase contrast images of beating myocardial microtissues were recorded using an

inverted microscope (Zeiss, Oberkochen, Germany) equipped with a CCD camera (Kappa

Opto-electronics GmbH, Gleichen, Germany) connected to a video recorder (Panasonic Inc.,

Hamburg, Germany).

Fluorescence-based characterization of cell morphologies

Myocardial microtissues were harvested following 7 days of cultivation in

maintenance medium. After a washing step in 2x phosphate-buffered saline (PBS; 150 mM

NaCl, 6.5 mM Na2HPO4 x 2 H2O, 2.7 mM KCL, 1.5 mM KH2PO4, pH 7.4; Sigma), the

spheroids were fixed for 1 hour in PBS containing 4% paraformaldehyde and subsequently

washed three times for 5 min in phosphate-buffered Triton X-100 (PBT, PBS containing

0.002% Triton X-100; Sigma). The spheroids were then permeated for 30 min in PBS

containing 0.5% Triton X-100. Primary antibodies specific for the indicated proteins as well

as fluorescence-labeled secondary antibodies were diluted in Tris-buffered saline (TBS,

20 mM Tris base, 155 mM NaCl, 2 mM EGTA, 2 mM MgCl2) containing 1% BSA and

incubated for 12 hours (primary antibody) at 4 C and 5 hours (secondary antibody) at room

temperature, respectively. Finally, the myocardial microtissues were washed in PBS and

embedded on glass slides using tris-buffered glycerol (a 3:7 mixture of 0.1 M Tris-HCl

(pH 9.5) and glycerol supplemented with 50 mg/ml n-propyl-gallat). In order to prevent

crunching of the myocardial microtissues between the slide and the cover slip 0.5 mm,

custom-made silicon spacers were used.

For immunofluorescence-based expression profiling the myocardial microtissues were

labeled with monoclonal antibodies specific for (i) sarcomeric- -actinin (Sigma; clone

EA53), (ii) polyclonal titin m8 (kindly provided by Mathias Gautel, King’s College London),

(iii) polyclonal myomesin (Agarkova et al. 2000), (iv) polyclonal -catenin (Sigma; cat no.

C2206), (v) collagen type I (Sigma; clone COL-1), (vi) monoclonal anti-collagen type IV

(Sigma; clone COL-94), (vii) polyclonal laminin (Sigma; cat no. L9393), (viii) polyclonal

fibronectin (Sigma; cat. no. F3648) or (ix) VEGF isoforms 110, 121 and 165 (Santa Cruz

Biotechnology Inc., Santa Cruz, CA; sc-152) and stained with Cy3-coupled secondary anti-

mouse (Jackson Immunochemicals, West Grove, PA; cat. no. 115-165-146) or FITC-coupled

anti-rabbit (ICN Pharmaceuticals, Hyland, CA) antibodies. F-actin was visualized using

A633-coupled phalloidin (Molecular Probes Inc., Eugene, OR). Nuclear staining of


cardiomyocytes was performed by using the fluorescent dye Draq5 (Biostatus Ltd., Shepshed,

Leicestershire, Great Britain): (i) NRC-derived microtissues were harvested by centrifugation

for 2 min at 50 x g and incubated for 10 min. in 100 �l PBS supplemented with 2 �l of a 5

mM Draq5 stock solution. Prior to immunostaining the microtissues were washed three times

with 1 ml PBS.

Phase contrast as well as fluorescence micrographs of cardiomyocyte monolayers

were recorded using an inverted microscope (HBO 50/AC, Zeiss, Oberkochen, Germany)

equipped with a digital camera (Axiocam HRm, Zeiss).

Confocal light microscopy

The imaging system consisted of an inverted fluorescence microscope (Leica

DMIRB/E, Heerbrugg, Switzerland) equipped with a Leica 20x/10x oil immersion objective,

a confocal scanner (Leica TCS SP1) featuring an argon-helium-neon laser and a Silicon

Graphics workstation (SGI, Schlieren, Switzerland) with Imaris 3D multi-channel image

processing software installed (Bitplane, AG, Zurich, Switzerland (Messerli et al. 1993)).

Lentivirus-based transduction technology

For production of replication-incompetent, self-inactivating lentiviruses a mixture

containing 94 l DMEM, 6 l Fugene 6 (Roche Diagnostics AG, Rotkreuz, Switzerland),

25 mM chloroquine, 1.5 g pLTR-G (encoding the pseudotyping envelope protein VSV-G of

the vesicular stomatitis virus (Reiser et al. 1996)), 1.5 g of the helper construct pCD/NL-

BH* (Mochizuki et al. 1998) and either 1.5 g of the YFP-encoding lentiviral expression

vector pMF351 (5’LTR-+-PhCMV-YFP-3’LTR U3) or the SAMY (secreted -amylase)-

encoding plasmid pMF364 (5’LTR-+-PEF1 -SAMY-3’LTR U3) was transfected into human

embryonic kidney cells (HEK293-T) (Mitta 2002; Schlatter et al. 2002). Following a medium

exchange after 24 h virus production was continued for another 48 h prior to filtration-based

harvesting of lentiviruses from the culture supernatant (0.45 m FP 030/2 filter; Schleicher &

Schuell GmbH, Dassel, Germany) which resulted in typical titers of 2 x 107 virus particles per

ml (Mitta 2002). In a standard transduction setting, 400’000 cardiomyocytes were infected in

a 6-well plate containing 2 ml medium and 200 l lentivirus suspension at a titer of

1x107 CFU/ml. Myocardial microtissues were directly transduced in hanging drops with 2 l

lentivirus suspension (1x107 CFU/ml).


Quantitative expression analysis of the secreted -amylase (SAMY)

100 l of SAMY-containing cell culture supernatant were centrifuged at 14’000x g for

2 min. 50 l were transferred into an Eppendorf cup also containing 1 ml substrate solution

(45 mg blue starch corresponding to 1 Phadebas®

tablet (cat. no. 10-5380-32, Pharmacia and

Upjohn, Uppsala, Sweden) dissolved in 4 ml H2O) and incubated for 15 min. at 70 C. The

reaction was stopped by addition of 250 l 0.5 M NaOH and the sample was centrifuged for 5

min at 14’000 x g prior to transfer into a 1 ml cuvette and colorimetric quantification at 620

nm (Schlatter et al. 2002).

Results

Production of myocardial microtissues

The cultivation of mammalian cells in hanging drops enables gravity-assisted

assembly of spheroids. Compared to shake-flask cultivation (Sperelakis 1978) or

centrifugation-enforced aggregation strategies (Armstrong et al. 2000), hanging drop-based

microtissues production may be considered as a rather natural three-dimensional cultivation

technology and most suitable for the functional design of complex myocardial spheroids. The

hanging drop cultivation technology was successfully adapted to produce myocardial

microtissues of neonatal rat (NRC) and mouse (NMC) cardiomyocytes. Under optimized

culture conditions 1’200 rat cardiomyocytes form in a single hanging drop a myocardial

spheroid of 130 ± 11 m in diameter with near 100% efficiency within four days (Figures 1

and 2). Microtissues derived from mouse populations of 1’200 cardiomyocytes reached 170 ±

12 µm in diameter at the same timepoint. As myocardium-derived cells are terminally

differentiated and proliferation-inert corresponding myocardial microtissues show an

invariant size/time profile when cultivated for a period of several days. Pure NRC cultures as

well as mixed NRC populations reflecting the cell type composition of rodent heart produce

myocardial microtissues of comparable size (130 ± 11 m; Figure 2A). Since the overall size

of myocardial microtissues is a direct function of the proliferation-incompetent

cardiomyocyte cell number it can be varied over a wide range (NRC: 130 11 µm –

230 11 m; NMC 170 12 µm – 320 19 m) by adjusting the concentration of the cell

suspension between 1’200 to 10’000 cells/hanging drop (Figure 2B). At and above 10’000


cells, myocardial microtissues reach a critical size associated with significant oxygen

limitation and attendant necrosis in the center (see below and Figure 7).

Figure 1: Time-dependent formation of myocardial microtissues in hanging drops. (A) Gravity-

enforced loose single-cell aggregation of density gradient-purified neonatal rat cardiomyocytes (NCR) at day 1

of a hanging drop culture. (B) After two days the cardiomyocytes rearrange to a lumpy pseudotissue until they

reach a compact microtissues state at day four (C). (scalebar = 100 m).

A

B

C

AA

BB

CC


Figure 2: Analysis of tissue growth and cell number/size profiles of neonatal rat (NRC) and

neonatal mouse cardiomyocyte (NMC)-derived myocardial microtissues produced in hanging drop cultures. (A)

Size profiles myocardial microtissues produced from 1’200 density gradient-purified NRCs, mixed NRC

populations (NRC Mix; 1’200 cells reflecting the typical cell type composition of rat hearts) as well as NMC

populations (1’200 cells total) at cultivation day 10. (B) Cell number-size correlations of NRC and NMC-derived

myocardial microtissues cultivated for four days in hanging drops. Since NRCs as well as NMCs are

proliferation-incompetent, the microtissue size is a function of the initial cell concentration inoculated in the

hanging drop.

48 h following cultivation of four-day-old myocardial microtissues in maintenance

medium supplemented with the 1-adrenergic agonist phenylephrine, NRC-derived

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microtissues started rhythmic contraction at beat frequencies of 60 beats per minute which

could be maintained for over three weeks. Figure 3 shows three-week-old beating NRC (Fig.

3A) and NMC (Fig. 3B)-derived myocardial microtissues, the video motion of which can be

accessed at http://www.biotech.biol.ethz.ch/martinf/staff/jens.html. Similar to the situation in

two-dimensional cardiomyocyte cultures, myocardial spheroids rapidly synchronize their

beating frequencies upon inter-microtissue contacts which exemplifies the capacity of

myocardial microtissues to form functional tissue interactions (Figure 4 and data not shown;

(Hertig et al. 1996)). In contrast to NRC-derived microtissues, NMC-based myocardial

spheroids initiate sustained contraction in the absence of any chemical stimuli following

aggregation.

Figure 3: Three-week-old NRC (A) / NMC (B)-derived beating heart microtissue which can be seen in

action at http://www.biotech.biol.ethz.ch/martinf/staff/jens.html. The sound of a human embryonic heart is added

to the motion picture to exemplify the regular contraction of myocardial microtissues at a frequency of 60 beats

per minute. (scalebar = 20 m).

A BA B


Figure 4: F-actin-based fluorescence micrograph of two interlinked NRC-derived myocardial

microtissues. Four-day-old myocardial microtissues have the potential to form microtissue-microtissue

superstructures and synchronize their contraction frequencies. (scalebar = 10 m).

Immunohistological characterization of myocardial microtissues

As the most physically energetic cell in the body contracting over three billion times

during an average human lifetime, cardiomyocytes are highly differentiated (Severs 2000).

Remodelling three-dimensional heart structures in vitro from isolated cardiomyocytes requires

maintenance of their shape and myofibrils while keeping the extraordinary mechanical

flexibility of this cell type. Also, electrical continuity between the cells has to be maintained

by intercalated disks, specialized cell-cell junctions, which are abundant in heart structures.

We have used a combination of immunofluorescence labelling and confocal microscopy to

characterize whether cardiomyocytes retained in vivo-like striation and cell-cell contacts

when grown as myocardial microtissues. In pure NRC- and NMC-derived myocardial

microtissues striation of myofibrils could be observed after a seven-day cultivation in

maintenance medium (Figures 5A and 5B). Also, -catenin-specific expression throughout

the entire myocardial microtissues indicates tissue-like cell-cell interactions within these

myocardial spheroids (Figures 5D and 5E) while individual cells retain their typical cell

morphologies associated with mature cardiomyocytes (Figure 5H).


Following production of myocardial microtissues from mixed cell populations

reflecting the natural cell type composition of rat hearts, muscle cells staining positive for

sarcomeric -actinin assembled at the periphery of myocardial spheroids following an as of

yet unknown self-organization or cell-positioning program within microtissues (Figure 5G).

Similarly, when coated directly onto a feeder spheroid assembled from myocardial fibroblasts,

the cardiomyocytes were evenly distributed on the surface (Figure 5G), showed intact cell-cell

contacts as evidenced by -catenin staining (Figure 5H) and demonstrated characteristic rod-

shaped morphology as well as intercalated discs (Figure 5I).

Figure 5: Immunohistological characterization of seven-days-old neonatal rat (NRC; A, D, G), mouse

(NMC; B, E, H) myocardial microtissues as well as NRC- coated fibroblast feeder spheroids (C, F, I). (A-C)

Sarcomeric -actinin was stained in red and F-actin in blue. (A, B, I) -actinin-based Z-disk-specific staining

visualizes intact myofibrils (see arrows). (D-F, I) Tissue-like cell-cell interactions throughout the entire

microtissues are exemplified by -catenin-specific staining in green and F-actin in blue. (H) In myocardial

mouse microtissues cardiomyocytes adopt a typical rod-shaped cell morphology reminiscent of primary

NMCNRC NRC Coat

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cardiomyocytes freshly isolated from rat hearts. (F) Following cultivation of myocardial microtissues produced

from mixed cell populations reflecting the natural cell type composition of rat hearts for seven days,

cardiomyocytes staining positive for -actinin are preferentially localized at the periphery of spheroids.

(overlapping staining for red ( -actinin) and green (F-actin) results in yellow). (scalebar = 10 m).

Characterization of the extracellular matrix of myocardial microtissues

The ability of artificial microtissues to produce a tissue-specific extracellular matrix

(ECM) typically associated with fundamental processes such as growth, differentiation, cell

migration and invasion will be essential for their use in clinical tissue engineering and gene

therapy (Weber et al. 1994; Giancotti et al. 1999). Development of a collagen I-containing

ECM is key to maintain cardiac mechanics during myocardial remodeling associated with

some heart diseases (Swynghedauw 1999). The natural myocardial ECM consists of collagen

I, fibronectin and proteoglycans forming the pericellular matrix and contains laminin as well

as type IV collagen as major constituents of the basal membrane (Farhadian et al. 1996). We

have performed immunofluorescence-based staining of extracellular matrix components in

order to characterize the ECM of neonatal rat and mouse cardiomyocyte-derived myocardial

microtissues as well as NRCs and NMCs cultivated in a two-dimensional anchorage-

dependent manner (Figure 6).

In cardiomyocyte monolayer cultures neither collagen I nor collagen IV were

produced whereas microtissues derived from pure and mixed NRC populations stained

positive for both collagen types (Figure 6A, C, D, F, H, I). By contrast, NMC-derived

microtissues showed only low-level collagen IV and no collagen I production (Figure 6E, J).

As collagen I is predominantly produced by fibroblasts, microtissues assembled from defined

mixed NRC populations accumulated higher levels of collagen I (Figure 6C). Myocardial

microtissues produced from purified cardiomyocyte cultures showed peripheral surface

expression of collagen I (Figure 6D). On the contrary, collagen IV is produced throughout

purified NRC-derived microtissues whereas myocardial spheroids generated from mixed NRC

populations stain positive for this collagen type predominantly at the spheroid’s surface

(Figure 6H, I). Fibronectin as well as laminin were expressed in two- as well as three-

dimensional NRC cultures (Figure 6K-T).


Figure 6: Comparative analysis of extracellular matrix (ECM) components of neonatal rat (NRC) and

mouse (NMC) cardiomyocytes cultivated in an adherence-dependent manner and as myocardial microtissue

produced from mixed whole-heart or density gradient-purified cardiomyocyte populations. To visualize

cardiomyocyte-specific structures in monolayer cultures titin m8 (shown in green) was used in combination with

collagen I and IV (shown in red; A, B, E, G) and myomesin (shown in green) was used together with laminin

and fibronectin (shown in red; K, L, P, Q). F-actin was stained with phalloidin (shown green) in combination

with collagen I (C-E), collagen IV (H-J), laminin (M-O) and fibronectin (R-T) (shown in red). (scalebar =

20 m).

Expression of vascular endothelial growth factor (VEGF) by myocardial

microtissues

Successful design of artificial tissues beyond a certain size as well as their connection

to the host capillary system following implantation requires vascularization (Griffith et al.

2002). The vascular endothelial growth factor (VEGF), a potent inducer of angiogenesis, is

induced under hypoxic conditions (Maulik et al. 2002). We have analyzed VEGF expression

in cardiomyocyte monolayers and myocardial microtissues of different sizes. Whereas NRCs

cultivated in monolayers and myocardial microtissues up to 130 11 m in diameter

K

A

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FN

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NRC

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NRC Mix NRCMonolayer

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NMC

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(corresponding to 1’200 NRCs) show no VEGF expression (Figure 7A, C), oversized

(230 11 m in diameter) myocardial microtissues assembled from 10’000 NRCs produce

this growth factor at high levels suggesting hypoxic conditions inside the spheroid (Figure

7E). Similar results were obtained with NMC’s cultivated as monolayers and microtissues

(Figure 7B, D, F).

Figure 7: VEGF (vascular endothelial growth factor)-specific immunostaining of neonatal rat and

mouse cardiomyocytes as monolayers (A, B) and myocardial microtissue cultures (C-F). (A, B) Sarcomeric

alpha-actinin-based striation of cardiomyocytes is shown in red and the presence of VEGF in green. (C, D)

Myocardial microtissue produced from 1’200 NRCs (130 m in diameter) and NMC’s (170 m in diameter),

which was stained with phalloidin (shown in red) and for VEGF expression (shown in green). (E, F) VEGF-

specific immunostaining (green) of oversized myocardial microtissue produced from 10’000 NRC’s and NMC’s

(NRC: 230 m in diameter; NMC: 320 m). (scalebar = 20 m).

Rat Mouse

D

A B

C

E F

Rat Mouse

D

A B

C

E F

DD

AA BB

CC

EE FF


Lentiviral infection of cardiomyocytes

Therapeutic interventions in mammalian tissues and artificial organs require an

efficient DNA transfer technology. Although adenovirus-based vectors enable remarkable

transduction efficiency in vitro, expression of delivered transgenes remains transient and

immune responses targeted against adenoviral proteins continue to be a major concern

(Schulick et al. 1995). Lentivirus-based transduction technology was shown to be highly

efficient for neonatal as well as adult cardiomyocytes (Rebolledo et al. 1998; Mitta 2002). We

have assessed the potential of third-generation self-inactivating lentiviral particles encoding

either the enhanced yellow fluorescent protein (YFP; pMF351, 5’LTR-+-PhCMV-EYFP-

3’LTR U3; (Mitta 2002)) or the secreted -amylase (SAMY; pMF364, 5’LTR-+-PEF1�-

SAMY-3’LTR U3; (Schlatter et al. 2002)) under control of the human PhCMV or PEF1�

promoters to transduce cardiomyocyte monolayers as well as NRC-derived myocardial

microtissues (Figure 8). pMF351-derived VSV-G (protein G of the vesicular stomatitis virus)-

pseudotyped lentiviral particles transduced NRC monolayer cultures with near 90% efficiency

and infected myocardial microtissues showed continuous bright yellowish fluorescence as a

consequence of viral EYFP delivery to peripheral cells (Figure 8). Interestingly, lentiviral

particles were unable to penetrate myocardial microtissues, as their centers remained

untransduced (Figure 8B).

In order to assess the specific heterologous protein production capacity of mammalian cells

grown in monolayers and microtissues, freshly isolated NRCs were transduced with SAMY-

encoding lentiviral particles (SAMY; pMF364, 5’LTR-+-PEF1 -SAMY-3’LTR U3; Schlatter

et al., 2002). For four days, half of the transgenic population was cultivated as a monolayer

and the other half in hanging drops to form myocardial microtissues. Isogenic control

infections of NRC monolayer cultures revealed no significant transduction-induced

morphological changes compared to mock-infected cultures (data not shown). Whereas the

monolayer culture produced respectable 5 U/cell -amylase NRCs assembled in myocardial

microtissues secreted over 6-fold more SAMY per cell (Figure 9). Owing to their high ectopic

protein production capacity microtissues may eventually become as valuable for in vivo

delivery of protein therapeutics in gene therapy scenarios as for tissue

engineering/replacement therapies.


Figure 8: Transduction of pMF351 (5’LTR- +-PhCMV-EYFP-3’LTR U3)-derived EYFP (enhanced

yellow fluorescent protein)-encoding lentiviral particles into NRC monolayer (A) and myocardial microtissue

cultures (B). (scalebar for monolayer (A) = 100 m; scalebar for myocardial microtissue (B) = 10 m).

Figure 9: Quantification of SAMY production of 1’200 NRCs transduced with pMF364 5’LTR- +-

PEF1 -SAMY-3’LTR U3)-derived SAMY (secreted -amylase)-encoding lentiviral particles followed by four-day

maintenance in monolayer cultures or hanging drop cultures to form myocardial microtissues.

A BA BS

AM

Y (

U/c

ell)

Microtissue Monolayer

0

10

20

30

40

SA

MY

(U

/cel

l)

Microtissue Monolayer

0

10

20

30

40

0

10

20

30

40


Discussion

Design and engineering of artificial heart tissues or entire hearts is one of the most

challenging scientific adventures of this millennium. Although advances in cardiac tissue

engineering are overdue since heart diseases are the leading non-infectious cause of death in

industrialized societies (i) production, (ii) rational reprogramming and (iii) in vitro assembly

of cells to form artificial heart structures have not yet become a clinical reality. As the most

energetically active cell in the heart, the cardiomyocyte is considered to be the key cell type

for cardiac tissue engineering (Severs 2000; Noble 2002). However, evolution of

cardiomyocytes which show unique mechanical flexibility, transformation resistance and

energetic activity has resulted in a highly specialized cell phenotype which is inert to current

cell expansion strategies, refractory to most gene transfer technologies and difficult to

assemble in three dimensions due to ongoing contraction (Datwyler et al. 1999; Magyar 1999;

Datwyler et al. 2001; Noble 2002; Pasumarthi et al. 2002).

So far, primary cardiomyocyte cultures from enzymatically dispersed fetal, neonatal or

adult vertebrate hearts are the only reliable source for pioneering cardiac tissue engineering.

Even though cardiomyocytes maintained in monolayer cultures have enabled detailed insight

into major structure- and gene-function correlations (Severs 2000), they are less responsive to

growth and differentiation factors and will therefore reveal fewer secrets relevant for cardiac

tissue engineering compared to cardiomyocytes assembled in three-dimensional cultures

(Armstrong et al. 2000). While three-dimensional cultivation of cardiomyocytes is generally

accepted to be key for cardiac tissue engineering there is ongoing controversy whether design

of artificial heart tissues requires scaffolds. Certainly, latest-generation biocompatible

scaffolds combine shape-supporting capacity with functional and bioactive properties (e.g.

differentiation, wound healing, cell/tissue adhesion, biodegradability) (Hubbell 1999;

Polonchuk et al. 2000). However, some scaffolds have been associated with post-implantation

side effects resulting from toxic degradation products, inflammatory reaction and poor

resorption (Yang et al. 2001). Scaffolds may still be the optimal solution for shaping rigid

tissue implants but they are intuitively less conceivable for the design of mechanically flexible

tissues exemplified by the myocardium (Hoerstrup et al. 2000; Ziegelaar et al. 2002).

The prevailing three-dimensional cultivation technology in cardiac tissue engineering

will have to unite several key characteristics including (i) long-term maintenance of the

beating capacity, (ii) multi-cell type cultivation, (iii) potential for self-organization,


polarization and microstructure formation between different cell types, (iv) production of an

extracellular matrix, (v) vascularization including induction of vascular vessel development

and connection to the host capillary system following implantation, (vi) development of inter-

tissue superstructures and (vi) compatibility with high-efficiency stable gene transfer

technologies to engineer complementary cell phenotypes or provide therapeutic interventions.

We have evaluated gravity-assisted myocardial microtissue formation following

cultivation of primary neonatal rat and mouse cardiomyocytes in hanging drops. Previously,

the hanging drop technology has generated generic impact on studying differentiation and

lineage control in embryoid bodies and on the production of hepatic microtissues(Magyar et

al. 2001; Boheler et al. 2002; Kelm 2002). Hanging drop-based cultivation of rodent

cardiomyocytes has resulted in myocardial microtissues which form characteristic z-band-

containing myofibrils enabling sustained and regular beating profiles for over three weeks.

Also, these functional myocardial microtissues retain their capacity to form microtissue-

microtissue superstructures which coordinate their beating frequency. Such interactions

between microtissues are of two-fold importance, first, they suggest functional integration

into the host organ following therapeutic implantation, and second, microtissue-microtissue

assembly may represent the building block towards larger-sized artificial tissues. A major

entity for higher order three-dimensional structures is the extracellular matrix, four major

constituents of which are expressed in rat myocardial microtissues: laminin, fibronectin as

well as collagen I and collagen IV. The only difference between rat and mouse cardiomyocyte

microtissues is the lack of collagen I production in mouse-derived spheroids. The

predominant production of collagen type I and IV (rat cardiomyocyte) and collagen type IV

(mouse cardiomyocytes) at the periphery of myocardial spheroids suggests an inside-out

polarization within these microtissues. Such a self-organization force is further exemplified

by the peripheral localization of cardiomyocytes in microtissues produced from mixed heart

cell populations reflecting the typical cell type composition of rodent hearts.

Although myocardial microtissues generated from heart-derived primary

cardiomyocytes cultivated in hanging drops show many of the aforementioned key

characteristics required for production of artificial tissue, their actual impact on cardiac tissue

engineering will depend on the design of artificial tissues beyond a few hundred micrometers

in diameter. As primary cardiomyocytes are terminally differentiated and proliferation-inert,

the size of myocardial microtissues is a direct function of cell number. However, due to

physico-chemical constraints, which limit nutrient and particular oxygen supply in the center

of artificial tissues cardiomyocyte-only-based microtissues cannot go beyond a certain size.


Interestingly, in contrast to cardiomyocyte monolayer cultures and small myocardial

microtissues (rat: 130 ± 11 m in diameter; mouse: 170 ± 12 m in diameter), oversized

cardiomyocyte-derived spheroids (rat: 230 ± 11 m in diameter; mouse: 320 ± 19 m in

diameter) produce VEGF. Such size-dependent VEGF expression may (i) mediate connection

to the host capillary system following implantation, (ii) enable in vitro vascularization in

multi-cell type-based microtissues, and (iii) eventually foster the design of large, fully

vascularized artificial (mini-) tissues.

Whatever artificial microtissue design will prevail in the future, successful cardiac

tissue engineering as well as heart-targeted gene therapy will require efficient gene transfer

technologies to express desired therapeutic or phenotype-modulating transgenes in

cardiomyocytes and/or myocardial microtissues. Successful transduction of cardiomyocyte

monolayers has been achieved with transgenic adenoviruses as well as Sindbis virus

(Datwyler et al. 1999; Zhou et al. 2000; Datwyler et al. 2001). However, both viruses mediate

only transient gene expression. We have recently designed a new series of lentiviral

expression vectors which transduce in their VSV-G-pseudotyped configuration even the

difficult-to-transfect adult rat cardiomyocytes (Mitta 2002). The same transgenic lentiviral

particles transduce the surface of myocardial microtissues at near 100% efficiency. A

comparative analysis of the production levels between cardiomyocytes cultivated as

monolayer or microtissue revealed that three-dimensional structures produced 6-fold more

reporter protein than two-dimensional cultures.

Complying with most of the aforementioned key tissue characteristics and bundled

with a powerful lentiviral transduction technology, myocardial microtissues are on their way

to establish cardiac tissue engineering as clinical alternatives to heart transplantation and

implantation of artificial non-biological hearts.

Acknowledgements

This work was supported by the Swiss National Science Foundation (grant no. 631-

065946), the Roche Research Foundation (grant no. 118-2001) and the Novartis Foundation

(grant no. 01C41). We thank Evelyne Perriard for isolation of neonatal mouse and rat

cardiomyocytes, Barbara Mitta for production/transduction of lentiviral particles, Stefan

Lange for video microscopy and Valeria Nicolini-Gonzalez as well as Beat P. Kramer for

critical comments on the manuscript.


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tissue engineering. Part I. Traditional factors." Tissue Eng 7, 679-689.

Zhou, Y. Y., S. Q. Wang, W. Z. Zhu, A. Chruscinski, B. K. Kobilka, B. Ziman, S. Wang, E.

G. Lakatta, H. Cheng and R. P. Xiao (2000). "Culture and adenoviral infection of adult

mouse cardiac myocytes: methods for cellular genetic physiology." Am J Physiol

Heart Circ Physiol 279, H429-436.

Ziegelaar, B. W., J. Aigner, R. Staudenmaier, K. Lempart, B. Mack, T. Happ, M. Sittinger, M.

Endres, A. Naumann, E. Kastenbauer and N. Rotter (2002). "The characterisation of

human respiratory epithelial cells cultured on resorbable scaffolds: first steps towards

a tissue engineered tracheal replacement." Biomaterials 23, 1425-1438.

Chapter 4

Self-Assembly of Sensory Neurons into Ganglia-Like

Microtissues

Kelm J.M., Ittner L.M., Born W., Djonov V. and Fussenegger M., (2005) J. Biotech., in press

Artificial Ganglia 56

Abstract

Unraveling intra- and inter-cellular signaling networks managing cell-fate control,

coordinating complex differentiation regulatory circuits and shaping tissues and organs in

living systems remain major challenges in the post-genomic era. Resting on the laurels of

past-century monolayer culture technologies, the cell culture community has only recently

begun to appreciate the potential of three-dimensional mammalian cell culture systems to

reveal the full scope of mechanisms orchestrating the tissue-like cell quorum in space and

time. Capitalizing on gravity-enforced self-assembly of monodispersed primary embryonic

mouse cells in hanging drops, we designed and characterized a three-dimensional cell culture

model for ganglion-like structures. Within 24 hours, a mixture of mouse embryonic

fibroblasts (MEF) and cells, derived from the dorsal root ganglion (DRG) (sensory neurons

and Schwann cells) grown in hanging drops, assembled to coherent spherical microtissues

characterized by a MEF feeder core and a peripheral layer of DRG-derived cells. In a time-

dependent manner, sensory neurons formed a polar ganglion-like cap structure, which

coordinated guided axonal outgrowth and innervation of the distal pole of the MEF feeder

spheroid. Schwann cells, present in embryonic DRG isolates, tended to align along axonal

structures and myelinate them in an in vivo-like manner. Whenever cultivation exceeded ten

days, DRG:MEF-based microtissues disintegrated due to an as yet unknown mechanism.

Using a transgenic MEF feeder spheroid, engineered for gaseous acetaldehyde-inducible

interferon- (ifn- ) production by cotransduction of retro-/lenti-viral particles, a short six-

hour ifn- induction was sufficient to rescue the integrity of DRG:MEF spheroids and enable

long-term cultivation of these microtissues. In hanging drops, such microtissues fused to

higher-order macrotissue-like structures, which may pave the way for sophisticated bottom-up

tissue engineering strategies. DRG:MEF-based artificial micro- and macrotissue design

demonstrated accurate key morphological aspects of ganglions and exemplified the potential

of self-assembled scaffold-free multicellular micro-/macrotissues to provide new insight into

organogenesis.

Introduction

Self-assembly is a basic phenomenon responsible for structural organization of living

systems. Although the differentiation of specific cell phenotypes and the layout map of cell


position in the early development of multicellular organisms are under strict genetic control

and imprinted in a localization-dependent manner, it is the process of cellular self-assembly,

mediated by cell-cell and cell-extracellular matrix (ECM) interactions, which fine-tunes the

organization of a particular cell quorum during histogenesis and organogenesis (Jakab et al.

2004; Wang et al. 2004). Steinberg’s pioneering hypothesis for the self-assembly of

embryonic cells was based on the concept of tissue fluidity and the assumption that

embryonic tissues can, in many respects, be regarded as liquids. In particular, on non-

adhesive surfaces or in suspension, multicellular aggregates reach their lowest free-energy

state by adopting a spherical shape, similar to liquid droplets (Steinberg 1963). The follow-up

differential adhesion hypothesis explains the liquid-like behavior as a function of tissue

surface and interfacial tensions resulting from cohesive (e.g., N-cadherin-mediated) and

adhesive (e.g., integrin-mediated) forces (Beysens et al. 2000; Lauffenburger and Griffith

2001; Steinberg 1963; Tepass et al. 2000). Akin to assembly and structure-shaping forces in a

multicellular embryonic developmental state, monodispersed cells derived from living tissues

retain their imprinted potential to self-assemble into defined in vivo-like structures

(Armstrong 1989; Duguay et al. 2003; Kelm and Fussenegger 2004; Layer et al. 2002;

Steinberg and Foty 1997).

In the vertebrate nervous system, the molecular crosstalk between neurons and glial

Schwann cells is key to maintaining and extending brain structures and innervated tissues

(Chen et al. 2003; Gordon-Weeks 2004). The neurons’ expression of trophic factors trigger

glial cell-fate commitment and glial cells then control survival, differentiation and

interconnection of associated neurons (Lemke, 2001). Although many neuronal differentiation

factors have been identified to date, the details of differentiation control and its coordination

with cell differentiation in neighboring tissues remain largely elusive (Chotard and Salecker

2004; Edlund and Jessell 1999; Griffiths and Hidalgo 2004; Lemke, 2001).

Since in-vivo experiments resolved several of the multifaceted phenomena underlying

neuro-, glia- and ganglia-genesis, future advances in this field will depend on information

gathered from complementary in vitro cell culture systems. The technology for sustained

cultivation of neurons as well as glia cells in a primary cell (co-)culture format will thus play

a pivotal role in deciphering the molecular events orchestrating cell-cell and cell-ECM

interaction in the developing nervous system (Zahir and Weaver 2004). Owing to the

aforementioned occurring in developing tissues, primary cells may lose their tissue-specific

phenotype following isolation and cultivation in a classical two-dimensional cell culture


format (Abbott 2003; Zhang 2004). Furthermore, the culture media, matrices and the absence

of neighboring cell communities may alter morphogen levels/gradients, which could bias the

readout of two-dimensional culture settings (Svenningsen et al. 2003). For these reasons, a

variety of sophisticated cell culture systems have been devised, which include multiple cell

types and consider the requirements for the development and organization of the peripheral

nervous system (Gingras et al. 2003; Suuronen et al. 2004b, Pittier et al. 2004). A priori,

sensory neurons and Schwann cells, derived from embryonic dorsal root ganglions (DRGs)

cocultivated with embryonic fibroblasts, may mimic the complex reciprocal interactions

between neurons and glia cells as well as either of these cell types with the connective tissue.

We have used gravity-enforced self-assembly of mouse DRG-derived cells, cocultured with

mouse embryonic fibroblasts (MEFs) in hanging drops, to create a robust three-dimensional

cell culture system to provide further insight into gangliogenesis.

Material and Methods

Isolation of mouse embryonic dorsal root ganglia (DRG) and fibroblasts

Eight to 12 DRGs, isolated from each E12 (embryonic day 12) embryo of time-mated

NMRI mice ((Taconic M&B A/S, Ry, Denmark) and from each E15 embryo of time-mated

Wistar rats (Janvier Elevage, Le Genest Saint Isle, France), were treated with Hank’s solution

(0.25% trypsin, Invitrogen, Carlsbad, CA) for 30 min. at 37°C, centrifuged for 3 min. at 130g

(800 rpm) and resuspended as monodipersed DRGs in neurobasalTM

medium (Invitrogen)

supplemented with 20% newborn calf serum (cat. no. P30-0400, lot no. P230704, PAN

biotech GmbH, Aidenbach, Germany) and 10 ng/ml nerve growth factor (2.5S-NGF;

Invitrogen). Cell-specific labeling of dissociated embryonic DRG populations was performed

using the PKH26 cell linker kit according to the manufacturer’s instructions (Sigma

Chemicals, Buchs, Switzerland). Mouse embryonic fibroblasts (MEF) were isolated from E14

embryos derived from time-mated mice (ICR-M-TKneo2, (Stewart et al. 1987)). Following

removal of the head and liver, the embryos were transferred to an ice-cold trypsin/EDTA

(0.05%/0.02%) solution (Pan Biotech GmbH) and incubated for 12h at 4°C. Subsequently, the

trypsin/EDTA solution was removed and the cells plated in Dulbecco’s modified Eagle

medium (DMEM; Invitrogen) supplemented with 10% fetal bovine serum (FBS, cat. no.

3302-P231902, lot no. P231902, Pan Biotech GmbH). Neonatal rat heart fibroblasts (RHF)

were isolated from newborn Wistar rats (Janvier Elevage). The hearts were dissected, digested


with collagenase (Worthington Biochemical Corp., Freehold, NJ) and pancreatin (Invitrogen)

and processed as described by Auerbach and coworkers (Auerbach et al. 1999).

Cell culture and microtissue production

MEFs, RHFs and NIH/3T3 (ATCC No. CRL-1658) were expanded in DMEM

supplemented with 10% FBS, 1% non-essential amino acids (Invitrogen) and 1%

penicillin/streptomycin solution (Sigma Chemicals). Monolayer fibroblast cultures were

trypsinized and mixed with monodispersed DRG populations at a cell ratio of 5x103/6x10

2

(MEF/DRG) prior to seeding into 60-well plates at indicated cell densities (HLA plate,

Greiner-Bio One, Frickenhausen, Germany). In order to enable gravity-enforced microtissue

formation in hanging drops, the 60-well plates were incubated upside down. Multicellular

microtissues were maintained in neurobasalTM

medium supplemented with 20% newborn calf

serum and 10 ng/ml NGF. For mixed MEF-DRG monolayer cultures, 4x105 MEFs were

seeded into 35 mm cell culture dishes and proliferation inactivated by adding 10 µg/ml

mitomycin C (Sigma Chemicals) for 2 h. After washing the monolayers three times in

phosphate-buffered saline (PBS; 150 mM NaCl, 6.5 mM Na2HPO4x2H2O, 2.7 mM KCL,

1.5 mM KH2PO4, pH 7.4), 7.5x104 DRG-derived cells were added.

Macrotissue assembly

Cylindrical agarose moulds (4% agarose in PBS [Sigma Chemicals]) containing a

3x10 mm casting chamber were produced from custom-designed Teflon models. In order to

produce a single macrotissue 900 microtissues were transferred to an agarose mould and

cultivated under static culture conditions in neurobasalTM

medium supplemented with 20%

newborn calf serum and 10 ng/ml NGF for two days at 37°C in a humidified 5% CO2-

containing atmosphere.

Immunofluorescence-based cell characterization

Microtissues were prepared for immunochemistry either as entire microtissues or as

frozen sections. Microtissues were harvested, washed twice in PBS, fixed for 1 h in 4%

paraformaldehyde-containing PBS, washed three times for 5 min. in phosphate-buffered

Triton X-100 (PBT; 0.002% Triton X-100 in PBS) and subsequently permeabilized for

60 min. in 0.5% Triton X-100-containing PBS. Primary antibodies, specific for desired


proteins as well as fluorochrome-conjugated secondary antibodies were diluted in 1% BSA-

containing Tris-buffered saline (TBS; 20 mM Tris base, 155 mM NaCl, 2 mM EGTA, 2 mM

MgCl2) and sequentially incubated with permeabilized microtissues for 12h at 4 C, with three

PBS washings between incubation periods. The microtissues were then washed in PBS and

embedded on glass slides using Tris-buffered glycerol (a 3:7 mixture of 0.1 M Tris-HCl

[pH 9.5] and glycerol supplemented with 50 mg/mL n-propyl-gallat). Tailored 0.5 mm

spacers were used to prevent squashing of the microtissues between the slide and cover slip.

The following primary antibodies were used: beta-catenin (rabbit, polyclonal, Sigma

Chemicals), BrdU (mouse, monoclonal, Sigma Chemicals), Brn-3A (mouse, monoclonal,

Chemicon, Hofheim, Germany), fibronectin (mouse, monoclonal, Sigma Chemicals), Gap-43

(mouse, monoclonal, Oncogene, Cambridge, MA, USA), laminin (rabbit, polyclonal, Sigma

Chemicals), myelin binding protein (MBP ; rat, monoclonal, Serotec, Düsseldorf, Germany),

N-cadherin (rabbit, polyclonal, BD Bioscience AG, Basel, Switzerland), neurofilament

(rabbit, polyclonal, Juro Supply GmbH, Luzern, Switzerland), neurofilament (mouse,

monoclonal, cat. Sigma Chemicals), Sox10 (mouse, monoclonal, kindly provided by Lukas

Sommer), and stained with Cy3-coupled secondary anti-mouse (Jackson Immunochemicals,

West Grove, PA, USA), Cy2-coupled anti-rat (Jackson Immunochemicals) or FITC-coupled

anti-rabbit (ICN Pharmaceuticals, Hyland, CA, USA) antibodies. F-actin was visualized using

A633-coupled phalloidin (Molecular Probes Inc., Eugene, OR, USA).

Histology

Macrotissues were washed once in PBS (Sigma Chemicals) and fixed for 4h at 4°C in

PBS containing 4% paraformaldehyde (Sigma Chemicals). Tissues were embedded in

TissueTec (Merck Eurolab AG, Dietikon, Switzerland) and frozen for 30 min at –25°C.

Sections (10 µm ) were generated using an Ultracut device (Zeiss, Feldbach, Switzerland).

Standard protocols were used for hematoxilin/eosin stainings on frozen sections as described

by Sheehan and Hrapchak (Sheehan and Hrapchak 1980).



DMIRB/E, Glattbrugg, Switzerland) equipped with Leica oil immersion objectives

(40x/20x/10x), a confocal scanner (Leica TCS SP1) featuring an argon and helium-neon laser


and a Silicon Graphics Workstation (SGI, Schlieren, Switzerland) with Imaris 3D multi-

channel image processing software installed (Bitplane, AG, Zurich, Switzerland (Messerli et

al. 1993)).

Transmission Electron Microscopy

For electron microscopic studies microtissues were fixed by total immersion in 0.1M

cacodylate buffer (pH 7.4, 350 mOsm) containing 2.5% glutaraldehyde. Tissue blocks were

postfixed in osmium tetroxide, block-stained using uranyl acetate, dehydrated by increasing

the ethanol concentrations and embedded in Epon 812 according to Djonov et al. (Djonov et

al., 2000) (all chemicals from Merck Eurolab AG, Dietikon, Switzerland). Semithin 1-µm

sections were stained with toluidine blue and visualized using a Leica DMIRB/E light

microscope (Leica Microsystems, Glattbrugg, Switzerland). Ultrathin sections, 80 to 90 nm

thick, were picked up on Formvar-coated (polyvinyl formal; Fluka Chemie AG) copper grids,

double-stained with lead citrate (Merck Eurolab AG) and uranyl acetate and monitored on a

Philips EM 400 electron microscope (FEI AG, Zurich, Switzerland).

Gas-inducible ifn- expression

Retroviral particles encoding 5’LTR-driven expression of Aspergillus nidulans’ AlcR

and the G418 resistance gene (neo), constitutively transcribed by the phosphoglycerate kinase

promoter (PPGK) (pWW506; 5’LTR-+-alcR-PPKG-neo-3’LTR) (Weber et al. 2004), were

produced using the packaging cell line GP-293 (Clontech, Basel, Switzerland; (Burns et al.

1993)) according to manufacturer’s protocol. Lentiviral particles encoding ifn- , controlled by

the AlcR-dependent acetaldehyde-inducible promoter PAIR (pWW430; 5’LTR-+-PAIR-ifn- -

3’LTR U3) (Weber et al. 2004), were produced as described before (Mitta et al. 2002). MEFs

were co-transduced with pWW506-derived retroviral (4 x 104

transducing units (TU/ml)) and

pWW430-derived lentiviral (1.4 x 105 TU/ml) particles (quantified as described by Mitta and

coworkers (Mitta et al. submitted)), mixed with DRG-derived monodispersed cells (see

above) and cultivated in hanging drops for five days. Thereafter, ifn- expression was induced

for 6h by gaseous acetaldehyde, evaporating from a 50 µl acetaldehyde-ethanol (1:4) solution

in the lid of a multi-well hanging-drop cultivation plate. IFN- production was quantified

with a Hu-ifn- ELISA kit according to the manufacturer’s protocol (R&D Systems,

Minneapolis, MN, USA).


Results

Cellular re-organization

Monodispersed cell populations, derived from dorsal root ganglions (DRG) of day 12

(E12) mouse embryos, were used as the source of Schwann cells and sensory neurons, the

major constituents of the peripheral nervous system. DRG-derived cell populations (900

cells / microtissue) were mixed with mouse embryonic fibroblasts (MEF; 5000

cells / microtissue) and grown for five to seven days in hanging drops to mediate gravity-

enforced reaggregation into multicellular spherical microtissues. Starting from a

homogeneous DRG:MEF mixed culture (Figures 1A and B) both cell populations self-

assembled into coherent microtissues within 24h of cultivation in hanging drops (Figure 1C).

Immunofluorescence microscopy revealed autonomous segregation of cell populations into a

central MEF spheroid harboring a growing pole of neurofilament structures (Figures 2A and

B; see Figure 4 below). On cultivation day 7, extensive polar ganglion-like caps formed on a

coherent MEF spheroid (Figures 2C and D; see Figure 4 below). Microtissues, derived from

mouse DRG-NIH/3T3 and embryonic (E15) rat DRG-rat heart fibroblasts co-cultures, also

developed polar neurofilament structures after cultivation in hanging drops for 7 days (Figure

2E and 2F). In contrast, DRG-derived cell populations cultivated on proliferation-inactivated

MEF feeder layers failed to show any type of ganglion-like assembly of sensory neurons

(Figures 2G and 2H).

Figure 1: Gravity-enforced microtissue self-assembly of mixed mouse dorsal root ganglion (DRG)-

derived cells and mouse embryonic fibroblast (MEF) populations. Monodispersed cell populations were

generated from twelve-day mouse embryos and cultivated in hanging drops. Following two-hour cultivation in

hanging drops DRG:MEF-derived cells form a homogeneous mixed population (A) exemplified by PKH26-

labeling of DRG-derived cells in red (B). Within the first 24 hours of hanging drop-based cultivation, DRG:MEF

populations form coherent microtissues by gravity-enforced self-assembly (C). (scalebar = 100 m)

A B CA B C


Figure 2: Auto-segregation of mouse-derived DRG:MEF microtissues into a polar DRG cells-derived

cap and a MEF-based feeder spheroid following prolonged cultivation in hanging drops. Concomitant with

microtissue formation 24h post seeding of mixed DRG:MEF populations into hanging drops, DRG-derived cells

segregate into a peripheral cell layer on a MEF-based core spheroid (A, B). On cultivation day 7, DRG-derived

cells form polar ganglion-like filament structures on the MEF feeder spheroid (C, D). Polar assembly of neurons

could also be observed after co-cultivation of mouse-derived DRGs with NIH/3T3 (E) and rat-derived DRGs

with rat heart fibroblasts (F) for 5 days in hanging drops. Microtissues are visualized by phase-contrast (A, C)

and immunofluorescence microscopy (B, D), showing neurofilament-specific staining of DRG-derived cells in

green and an F-actin-specific expression by the MEF spheroid in red. (G, H) Fluorescence micrographs of

monolayer cultures derived from 12 day-old mouse embryos grown on a two-dimensional proliferation-

inactivated MEF feeder layer. The MEF feeder layer is visualized by F-actin-specific staining (red) and the

neuronal network stains positive for neurofilament (green) (G). Brn-3A-positive staining specifies neurons as

sensory neurons (Brn-3A shown in red, neurofilament shown in green) (H). (scalebar = 30 m)

Development of 3D neuronal structures

To demonstrate that the ganglion-like cap structures originated from self-organization

of sensory neurons we co-visualized neuron-(Brn-3A) and neurofilament-specific cell staining

of microtissues on cultivation days 1, 3, 5 and 7. 24h post seeding of the DRG:MEF co-

culture in hanging drops, a peripheral assembly of sensory neurons was already prominent

(Figure 3A). In a three-day-old microtissue, neurofilament structures had developed and

continued to grow for four more days (Figures 3B-D). Images of bromodeoxyuridine (BrdU, a

mitotic indicator) did not reveal significant cell division, suggesting that ganglion-like

structures had assembled from sensory neurons or neuronal precursor cells present in the

DRG cell preparations (Figures 3E and 3F). Detailed analysis of ganglion-like structures by

G

H

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3D projection of scanning confocal micrographs revealed the full scope of polar ganglion

assembly along with an axon-like outgrowth of sensory neurons covered by Schwann cells

staining positive for Sox10, a Schwann cell-specific transcription factor (Lobsiger et al. 2002)

(Figure 4). Axonal structures covered with Schwann cells were myelinated in 10-day-old

microtissues as shown by myelin binding protein (MBP)–specific immunofluorescence

(Figures 5A and 5B). This observation was confirmed by ultrastructural analysis (Figure 5C).

Neuron-specific growth-associated protein 43 (Gap-43) is typically expressed in

outgrowing axons during development, regeneration, axonal sprouting and synaptic plasticity.

In all of these processes, Gap-43 is a major constituent of axonal growth cones managing

axon motility and pathfinding (Wiese et al. 1992). We have detected Gap-43 in axonal

structures developed from ganglia-like cap structures (Figure 6A). During development and

regeneration, axon guidance is mediated by different cellular factors as well as by

extracellular matrix (ECM) components including laminin and fibronectin. In DRG:MEF-

derived microtissues, ECM-based stimuli are provided by the MEF feeder spheroid

expressing high levels of laminin and fibronectin (Figures 6B and 6C). Furthermore, it has

recently been established that cell type-specific segregation of co-cultured cell populations is

mediated by cadherin-subtype-specific interactions (Duguay et al. 2003). In DRG:MEF-

derived 3D co-cultures, N-cadherin is exclusively expressed in sensory neurons, suggesting a

role of this intercellular adhesion protein in the self-assembly of sensory neurons into

ganglion-like structures (Figure 6D).

Figure 3: Assembly and proliferative activity of neurofilament cap structures in DRG:MEF

microtissues. DRG:MEF microtissues cultivated for 1 (A), 3 (B), 5 (C) and 7 (D) days in hanging drops were

visualized by Brn-3A (red; A-D) and neurofilament (green; A-D) expression as well as bromodeoxyuridine

E

FD

CA

B

E

FD

CA

B


(BrdU) incorporation (red; E, F) by immunofluorescence. On cultivation day 1, sensory neurons assemble at the

periphery of a MEF-based core spheroid and produce polar ganglion-like neurofilament structures, which exhibit

low proliferative activity (G, H). (A-D scalebar = 20 m; E, F scalebar = 5 m)

Figure 4: Three-dimensional confocal micrograph projection of neurofilament structures developed in a

5 day-old DRG:MEF spheroid. (A) Ganglia-like neurofilament structures (green). (B) Ganglia-like

neurofilament structures (green) showing Brn-3A expression specific for sensory neurons (red). (C) Schwann

cells, stained for specific expression of the transcription factor Sox10 (red), accumulate alongside neurites

(green). (scalebar = 30 m)

Figure 5: Immunofluorescence micrographs of axon-covering myelin sheaths visualized by

neurofilament- (red, (A)) and myelin-binding protein- (MBP) specific co-staining (green, (B)). Transmission

electron microscopy (TEM)-based ultrastructural analysis confirmed myelination (arrow) of axonal structures

(asterisk) (C). (A, B scalebar = 10µm; C scalebar = 1 µm)

AA BB CAAAA BBBB CC

CA

B

CAA

BB


Long-term cultivation of DRG:MEF microtissue cultures

DRG:MEF-derived microtissues, cultivated for more than ten days, lost their tissue

integrity and disintegrated. Interferon- (IFN- ), approved in the past decade for the

treatment of relapsing-remitting multiple sclerosis, is a well-known factor in preserving

neuronal structures (Kappos 2004; Njenga et al. 2000). Using the aforementioned DRG:MEF-

derived microtissue design we grew wild-type DRG-derived cells on a transgenic MEF

(MEFtransgenic) feeder spheroid, engineered for inducible IFN- production using the gas-

inducible AIR (acetaldehyde-inducible regulation) technology (Weber et al. 2004). MEFs

were cotransduced with retro- and lenti-viral particles, which mediate constitutive expression

of the Aspergillus nidulans AlR transactivator (pWW506; 5’LTR-+-alcR-PPKG-neo-3’LTR;

retrovector) and enable AlcR-dependent IFN- expression from the acetaldehyde-inducible

promoter PAIR (pWW430; 5’LTR-+-PAIR-ifn- -3’LTR U3; lentivector). IFN- production by

MEFtransgenic feeder spheroids was induced for 6h by placing acetaldehyde into the lid of multi-

well hanging drop cultivation plates containing 5-day-old DRG:MEFtransgenic microtissue

cultures. Acetaldehyde, evaporating from the lid, dissolves into the hanging-drop culture

media at regulation-effective concentrations and induces PAIR-driven IFN- production

(Weber et al. 2004). Such a 6-h induction profile resulted in IFN- levels of 235 9 pg/ml in

the supernatant of the microtissue culture (Figure 7A). Following ifn- induction, all the

spheroids were harvested, washed three times in phosphate-buffered saline (PBS) to remove

the remaining acetaldehyde and cultivated for another nine days in hanging drops. Toluidine

blue-stained sections of DRG:MEFtransgenic microtissue, cultivated in the presence and absence

of acetaldehyde, demonstrated that IFN- production prevented time-dependent microtissue

disintegration. Whereas spheroids cultivated for 14 days in the absence of IFN- production

disintegrated, microtissues subjected to pulsed ifn- expression maintained a homogeneous

tissue format and the aforementioned cellular tissue organization (Figures 7B-E).

Ultrastructural analysis confirmed exclusive disintegration of IFN- -repressed feeder

spheroids highlighted by dissolution of the nulcear membrane and emergence of digestive

vacuoles filled with cell debris (Figures 7C and 7E).


Figure 6: Immunofluorescence micrographs of 7 day-old DRG:MEF spheroids stained for Gap-43 (A;

shown in green), laminin (B; shown in green), fibronectin (C, shown in green) and N-cadherin (D, shown in

green). Neurofilament is shown in red throughout (yellow in overlay with green) (A-D). (A, B, C scalebar =

20 m; D scalebar = 10 m)

Figure 7: Gas-inducible interferon- (ifn- ) expression preserves the integrity of DRG:MEF spheroids.

DRG:MEF spheroids, containing MEFs engineered for gas-inducible ifn- expression, were analyzed for IFN-

production on cultivation day 5 (A), and the integrity of microtissues exposed to IFN- (B, C) or no IFN- (D,

E) was scored by toluidine blue-based staining semi-thin sections as well as transmission electron micrographs

(TEM) on cultivation day 14. (B, D scalebar = 20 m; C, E scalebar = 1 µm)

Assembly of innervated macrotissues

While microtissues represent a powerful model system for the study of multicellular

tissue affairs, tissue engineers will require larger tissue structures for clinical applications. We

have therefore designed an agarose-based casting mould for assembly of scaffold-free

ganglia-like microtissues to higher-order macrotissues. 900 microtissues, each composed of

5’000 MEFs with 600 DRGs, were seeded into the agarose mould. After two days in the

mould culture, spheroids had assembled into a single coherent 3 mm-thick macrotissue

cylinder (Figure 8A). Hematoxylin- and eosin- (H&E) staining confirmed that microtissues

A C DBA C DB

+ acetaldehyde - acetaldehyde

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completely fused to the macrotissue although spherical substructures were still be anticipated

at some places within the macrotissues (Figure 8B-D). Cavities resulting from inter-spheroid

fusion were filled by fibroblasts (Figures 8B-D). Neurons assembled preferentially at the

periphery of macrotissues but failed to form specific poles typical of MEF:DRG microtissues

(Figures 8E, F).

Figure 8. Macrotissues produced from 900 5 day-old DRG:MEF-derived microtissues assembled in an

agarose mould for 2 days (A). Hematoxylin and eosin-based staining of frozen sections (B-D) and

immunochemical staining of neurofilament (green) and F-actin (red; E, F). (A, scalebar = 500 µm; B-F scalebar

= 20 µm)

Discussion

Although many of the key cellular and extracellular regulatory components have been

identified over the past decade, including soluble growth factors, insoluble matrix factors and

receptors on the cells themselves, the fundamental principles by which cells organize into

structured tissues remains largely elusive. Yet, only the ability to understand and control

spatial distributions of multiple cell populations will enable rational approaches to tissue

engineering applications where precise multicellular organization in three-dimensional

structures is required. Particularly important tissue engineering objectives include

vascularization and innervation of tissues (McIntire 2002; Nomi et al. 2002; Suuronen et al.

2004a; Suuronen et al. 2004b).

B

A

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EC

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EC

D


A sensory nerve supply is crucial for optimal tissue function and to enhance wound

healing of its target tissue. We have designed and characterized the first tissue-engineered

three-dimensional culture system that morphologically and physiologically reproduces a

peripheral nerve regeneration process in vitro by co-culturing mouse embryonic fibroblasts

and DRG-derived cells in a scaffold-free format. Short-range cell-cell interactions are

important for controlling cellular organization and cell-fate decisions, as exemplified by

neural crest stem cells (Hagedorn et al. 2000). Since connective tissue is a key regulator of

axonal outgrowth it should be included in a three-dimensional in vitro model of the peripheral

nervous system development (Gingras et al. 2003). In order to alleviate potential interference

of scaffold material and scaffold-breakdown products with short-range cell-cell interactions,

we conceived a completely cell-based 3D co-culture model, embedding embryonic dorsal root

ganglia (DRG) cell populations in a 3D fibroblast feeder environment.

The aim of tissue engineering is not only to create desirable organs, but also to better

understand the fundamental mechanisms and principles of biological organization. Classical

tissue engineering has been based on seeding cells into biodegradable polymer scaffolds or

gels, culturing and expanding the composite cell-scaffold material in sophisticated bioreactors

and implanting the resulting tissue into the recipient organisms where maturation of the new

organ takes place (Griffith and Naughton 2002). During early embryogenesis, sequential well-

orchestrated forces rearrange cells into structures, which are imprinted by follow-up

molecular cell-fate decisions and an extensive extracellular matrix (ECM) coordinating the

interface between differentiating/differentiated cells. Gravity-enforced cell assembly of

primary embryonic cells in hanging drops is a straightforward technology for producing

organized, multi-cell type-based three-dimensional cell-culture models. Owing to their high-

throughput compatibility – both for production and analysis – microtissues derived from

mixtures of a few hundred monodispersed primary cells represent a flexible tissue culture

system for studying classical tissue engineering strategies, generating readout in drug

discovery and testing initiatives with unmatched precision and fostering unprecedented

insight into developmental principles (Lauffenburger and Griffith 2001; Zhang 2004).

As predicted by the differential adhesion hypothesis, which suggests cell type-specific

assembly based on net forces resulting from cadherin-mediated interactions between identical

cell types and cell-ECM interactions (Duguay et al. 2003; Lauffenburger and Griffith 2001;

Steinberg 1962a; Steinberg 1962b), embryonic DRG-derived sensory neurons mixed with

MEFs segregated from the fibroblasts to assemble a polar cap structure on a MEF-derived


feeder spheroid. Sensory neurons exclusively expressed N-cadherin, an intercellular adhesion

molecule shaping and maintaining the pole structure. Following polar cap assembly, the

sensory neurons formed an organized axonal outgrowth in a time-dependent manner.

Formation of axon-like structures likely resulted from re-assembly of existing sensory

neurons rather than from proliferation-based outgrowth, since BrdU-based assessment of cell-

cycle activities of sensory neurons were scored too low to account for the observed massive

organization of axonal fibers. Gap-43, the growth-associated protein known to coordinate and

guide axonal outgrowth, was strongly expressed in three-dimensional neuronal structures.

Furthermore, Schwann cells, specified by expression of the transcription factor Sox10,

showed a high in vivo-like tendency to align along nerve fibers. Thus, DRG:MEF-based

microtissues enabled key structures, which are relevant for detailed studies of peripheral nerve

system development and tissue innervation, including polar N-cadherin-based assembly of

ganglion-mimicking structures, guided Gap-43-mediated outgrowth of axon-like fibers,

sensory neuron-Schwann cell interactions and myelination of axons. Most of these

morphologic and cellular crosstalk characteristics fail to establish to a similar extent in an

isogenic two-dimensional cultivation system, suggesting that maintenance of cells in a tissue-

like format is essential for gaining further insight into aspects of cell-cell/cell-matrix

interactions relevant to tissue engineering (Cukierman et al. 2001).

Micro-scale tissue engineering – the production of microtissues – is a science at the

interface of engineering and organ replacements and exploits three-dimensional multi-cell

type cultures (i) for obtaining new insight into inter-cellular crosstalk (Hynds et al. 2004), (ii)

as building blocks for higher-order tissue design (Jakab et al. 2004) and (iii) for high-

throughput-compatible drug screening and testing (Bhadriraju and Chen 2002). Currently

available two-dimensional cell culture systems are limited and unreliable with respect to their

screening and testing readout as many drug (target) functions are sensitive to a multicellular

microenvironment (Bhadriraju and Chen 2002).

To date, only a few cell culture systems exist, all of which rely on scaffolds or

matrices to assemble sensory neurons, Schwann cells and connective tissue into a three-

dimensional format (Gingras et al. 2003; Suuronen et al. 2004b). Our strategy for producing

microtissues from monodispersed embryonic DRG populations by gravity-enforced self-

assembly represents the first scaffold-free approach to generating ganglia-like structures,

compatible with high-throughput assays and designed to discover/study factors triggering

cell-fate/morphologic control of sensory neurons and Schwann cells. These cell types are


currently in the limelight because of their association with different untreatable

neurodegenerative diseases (England and Asbury 2004).

As shown by a variety of recent examples, scaffold-free micro-scale tissue engineering

will expand our knowledge of developmental phenomena and will almost certainly impact

future drug testing and discovery initiatives.

Acknowledgements

We thank Maurice Kleber and Lukas Sommer for their generous advice and supply of

primary cells as well as Lucilla Nobbio and David Fluri for critical comments on the

manuscript. This work was supported by the Swiss National Science Foundation (grant no.

631-065946), the Swiss State Secretariat for Education and Research within EC Framework 6

and Cistronics Cell Technology GmbH, Einsteinstrasse 1-5, CH-8093 Zurich, Switzerland.

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Chapter 5

VEGF Profiling and Angiogenesis in Human

Microtissues

Kelm J.M., Diaz Sanchez-Bustamante C., Ehler E., Hoerstrup S.P., Djonov V., Ittner L. and

Fussenegger M., (2005) J. Biotech., in press

Microtissue Vascularization 76

Abstract

Owing to its dual impact on tissue engineering (neovascularization of tissue implants)

and cancer treatment (prevention of tumor-induced vascularization), management and

elucidation of vascularization phenomena remain clinical priorities. Using a variety of

primary human cells and (neoplastic) cell lines assembled in microtissues by gravity-enforced

self-aggregation in hanging drops we (i) studied size and age-dependent VEGF production of

microtissues in comparison to isogenic monolayer cultures, (ii) characterized the self-

organization and VEGF-production potential of mixed-cell spheroids, (iii) analyzed VEGF-

dependent capillary formation of HUVEC (human umbilical vein endothelial cells) cells

coated onto several human primary cell spheroids and (iv) profiled endostatin action on

vascularization in human microtissues. Precise understanding of vascularization in human

microtissues may foster advances in clinical tissue implant engineering, tumor treatment, as

well as drug discovery and drug-function analysis.

Introduction

Transplantation remains the preferred clinical intervention for the treatment of organ

failures. Owing to a global shortage in donor organs alternative strategies providing

bioengineered tissue for replacement of damaged, injured or missing tissues are of high

clinical priority (Griffith and Naughton 2002). An artificial tissue with more than a few cubic

millimeters cannot survive by simple diffusion and requires formation of new blood

capillaries to supply essential nutrients/oxygen and enable connection to the host vascular

system following implantation (Griffith and Naughton 2002; Mooney and Mikos 1999).

Likewise, tumors do not grow beyond a few millimeters unless they become vascularized by

directing an ingrowth of capillaries from adjacent blood vessels (Folkman and Kaipainen

2004). Therefore, precise control of pro- and anti-angiogenic activities is of prime clinical

interest in current cancer therapy and tissue engineering initiatives (Bergers and Benjamin

2003; Nomi et al. 2002). The maturation of nascent vasculature, formed by vasculogenesis or

angiogenesis, requires recruitment of mural cells, generation of an extracellular matrix and

specialization of the vessel wall for structural support and regulation of vessel function. In

addition, the vascular network must be organized to provide parenchymal cells with sufficient

nutrients. All of these processes are orchestrated by physical forces, as well as a panoply of


ligands and receptors whose spatio-temporal expression patterns are tightly regulated (Jain

2003; Yancopoulos et al. 2000).

Angiogenesis is a morphogenic process of new blood capillaries emerging from

preexisting vessels which consists of six major steps including (i) vasodilatation of the

parental vessel, reducing endothelial cell contact, (ii) degradation of the basement membrane

by a variety of proteolytic enzymes, (iii) migration and proliferation of endothelial cells at the

spearhead of new vessels, (iv) production of the capillary lumen and formation of tube-like

structures, (v) basement membrane synthesis and (vi) recruitment of vascular smooth muscle

cells. The sequence of molecular events resulting in angiogenesis requires precise fine-tuning

of multiple signaling pathways, cell-cell and cell-matrix interactions (Jain 2003).

Vascular endothelial growth factor (VEGF), a major regulator of neovascularization

under physiological and pathological conditions (Gerber et al. 1999), is produced in five

homodimeric isoforms (VEGF121, VEGF145, VEGF165, VEGF189, VEGF206), which differ in

their expression levels and their localization. VEGF family members are involved in (i) the

formation of immature vasculature (VEGF-VEGF receptor 2-mediated signaling in

angioblasts results in formation of the dorsal aorta and the cardinal vein) (Yancopoulos et al.

2000); (ii) induction of migration and proliferation of endothelial cells (Conway et al. 2001);

(iii) vessel dilatation and sprouting in the presence of angiopoietin-2 (Tsigkos et al. 2003),

(iv) stabilization of immature vasculature (VEGF-induced platelet-derived growth factor

secretion of endothelial cells facilitates recruitment of mural cells) (Blau and Banfi 2001); (v)

sequestration of angiopoietin 2 which destabilizes vessels (Tsigkos et al. 2003); (vi)

suppression of apoptosis (Folkman 2003), (vii) branching, remodeling and pruning of

vasculature (protease-mediated release of matrix-sequestrated VEGF) (Peirce and Skalak

2003) and (viii) vessel specialization (arterial growth promoted by VEGF-VEGF receptor 2-

neuropilin 1 signaling). Recent studies indicate that VEGF action goes beyond vascularization

and may be involved in neurogenesis (Jin et al. 2002), as well as growth- and survival-

modulation of chondrocytes (Schipani et al. 2001).

The biological effects of VEGF are extremely dose dependent. Loss of even a single

allele results in fatal vascular defects in the embryo (Ferrara and Alitalo 1999) and

insufficient levels of VEGF lead to post-natal angiogenesis and ischemic heart disease

(Carmeliet et al. 1999). Several control circuits are at work to balance VEGF action. Hypoxia-

mediated induction of hypoxia-inducible factor (HIF-)1 resulting in VEGF production, and

endostatin, a matrix-associated protease-mediated cleavage product of collagen XVIII,

inhibiting VEGF-induced mobilization of endothelial cells are some examples (Schuch et al.


2003; Semenza 2003). A variety of strategies for therapeutic angiogenesis have been designed

including (i) delivery of recombinant angiogenic molecules through controlled-release

devices (Jain and Carmeliet 2001) and (ii) functionalized matrices (Richardson et al. 2001;

Zisch et al. 2003) or (iii) transfection/transduction of (engineered) angiogenesis-modulating

cDNAs (Elson et al. 2001; Isner 2002).

Although most of today’s knowledge on vascularization regulatory networks has been

derived from in vitro monolayer cultures (Vailhe et al. 2001), three-dimensional (3D)

cultivation technologies may reveal further insight. Current 3D models include formation of

primitive vascular networks in vitro by coculturing endothelial cells with mural cells or their

precursors (Hirschi et al. 1999; Korff et al. 2001). However, several groups have suggested

that blood vessels of artificially pre-vascularized cells/tissues remain essentially self-

contained and do not become connected with the surrounding vasculature (Lee et al. 2000;

Springer et al. 1998).

Based on our previous observations that VEGF production in myocardial microtissue

is strictly correlated to cell number and microtissue size (Kelm et al. 2004a; Kelm et al.

2004b) we have established an entirely human cell-based microtissue format to provide new

insight into VEGF production, angiogenesis and blood vessel formation.


Isolation of primary human aortic fibroblasts

In order to obtain primary human aortic fibroblasts (HAF), de-endothelialized vessel

segments of the human aorta were minced and cultivated in a 37°C humidified 5% CO2-

containing atmosphere and Dulbecco's modified Eagle's medium (DMEM, Invitrogen,

Carlsbad, CA, USA) supplemented with 10% fetal calf serum (FCS; cat. no. A-15-022, lot no.

A01129-242; PAA Laboratories, Linz, Austria) and 1% penicillin/streptomycin solution

(Invitrogen). Pure HAFs which had migrated out of the tissue pieces after 10 to 14 days were

serially passaged and expanded for 4 to 6 weeks under aforementioned conditions to desired

cell numbers.

Cell culture

Human primary aortic fibroblasts (HAF), the hepatocellular carcinoma cell line

(HepG2, DSMZ: ACC 180), newborn human foreskin fibroblasts (Hs68, ATCC: CRL-1635)


and human fibrosarcoma cells (HT-1080, ATCC: CCL-121) were expanded as monolayer

cultures in DMEM supplemented with 10% FCS. Human umbilical vein endothelial cells

(HUVEC, No. C-12200; Lot 0111701), normal human dermal fibroblasts (NHDF, No. C-

12300; Lot 1070402) and human umbilical artery smooth muscle cells (HUASMC, No. 252-

05; Lot 1222) were obtained from PromoCell (Heidelberg, Germany). HUVECs (PromoCell,

No. C-22110), HUASMCs (PromoCell, No. C-22162) and NHDFs (PromoCell, No. C-23010)

were expanded in monolayer cultures using PromoCell’s endothelial cell media supplemented

with 10% FCS (HUVECs and HUASMCs only). Human articular chondrocytes (HAC, kindly

provided by Millenium Biologix) were cultured in DMEM/F12 (Invitrogen) supplemented

with 10% FCS. All cell types were cultivated at 37°C in a humidified 5% CO2-containing

atmosphere.

Microtissue production

Monolayer cultures of desired cell types were trypsinized and single-cell suspensions

seeded at indicated cell densities into 60-well plates (HLA plate, Greiner-Bio One,

Frickenhausen, Germany). In order to enable gravity-enforced microtissue formation in

hanging drops, the 60-well plates were incubated upside down. Pure, multicellular and coated

microtissues were produced/maintained in DMEM medium (Invitrogen) supplemented with

10% FCS and 1% penicillin/streptomycin solution. Following cultivation for 2 to 8 days in

hanging drops the microtissues were harvested for further analysis. Onion skin-like

multicellular microtissues were produced in two steps: (i) production of the core feeder

spheroid by 2-day cultivation in hanging drops followed by (ii) cocultivation of feeder

spheroids and monodispersed coating cells (for example HUVECs) in hanging drops.

Fluorescence-based characterization of cell morphologies

Microtissues were harvested, washed twice in phosphate-buffered saline (PBS;

150 mM NaCl, 6.5 mM Na2HPO4x2H2O, 2.7 mM KCL, 1.5 mM KH2PO4, pH 7.4; Sigma

Chemicals, St. Louis, MO), fixed for 1h in PBS containing 4% paraformaldehyde and

subsequently washed three times for 5 min in phosphate-buffered Triton X-100 (PBT, PBS

containing 0.002% Triton X-100; Sigma). The microtissues were then permeabilized for

60 min in PBS containing 0.5% Triton X-100. Primary antibodies specific for desired proteins

as well as fluorescence-labeled secondary antibodies were diluted in 1% BSA-containing

Tris-buffered saline (TBS, 20 mM Tris base, 155 mM NaCl, 2 mM EGTA, 2 mM MgCl2) and


sequentially incubated with microtissues for 12 hours at 4 C with three PBS washings in

between. Finally, the microtissues were washed in PBS and embedded on glass slides using

Tris-buffered glycerol (a 3:7 mixture of 0.1 M Tris-HCl (pH 9.5) and glycerol supplemented

with 50 mg/ml n-propyl-gallat). Tailored 0.5 mm silicon spacers were used to prevent

crunching of the microtissues between slide and cover slip. Immunofluorescence-based

analysis of microtissues required antibodies specific for human von Willebrand factor (F3520;

Sigma), platelet-endothelial cell adhesion molecule-1 (PECAM-1; P8590, Clone WM-59;

Sigma), alpha-smooth muscle actin (A2547, clone 1A4; SIGMA), V 3 integrin (ab7167,

Abcam, Camebridge, UK) and/or vascular endothelial growth factor (VEGF) isoforms, 121

and 165 and 189 (sc-152; Santa Cruz Biotechnology Inc., Santa Cruz, CA) all of which were

visualized using Cy3-coupled anti mouse (Jackson Immunochemicals, West Grove, PA; cat.

no. 115-165-146) and FITC-coupled anti-rabbit secondary antibodies (ICN Pharmaceuticals,

Hyland, CA). F-actin was stained using A633-coupled phalloidin (Molecular Probes Inc.,

Eugene, OR).

Toluidine blue staining and immunohistochemistry of paraffin-embedded

microtissue sections

Microtissues were harvested, washed once in PBS (Sigma) and fixed for 2h at 4°C in

PBS containing 4% paraformaldehyde (Sigma). Following stepwise dehydration in ethanol,

microtissues were embedded in paraffin (Fisher Scientific, Wohlen, Switzerland). Toluidine

blue (Fluka Chemie AG, Buchs, Switzerland) staininigs of 5µm microtissue sections was

performed as described before (Sheehan and Hrapchak 1980). Immunohistochemical staining

included incubation of rehydrated 5µm microtissue sections with von Willebrand factor-

specific antibodies (F3520; Sigma) for 30 minutes at room temperature. Microtissue sections

were subsequently visualized for von Willebrand factor production using the Vectastain ABC

method (Vector Laboratories, Burlingame, CA) and the Metal Enhanced DAB Substrate

(Pierce Biotechnology, Rockford, IL). For antigen unmasking, microtissue sections were

heated for 5 minutes in 10 mM sodium citrate buffer (pH 5.8). Endogenous peroxidase

activity was blocked with 3% hydrogen peroxide prior to incubation with primary antibodies.



DMIRB/E, Glattbrugg, Switzerland) equipped with a Leica 20x/10x oil immersion objective,


a confocal scanner (Leica TCS SP1) featuring an argon and helium-neon laser and a Silicon

Graphics Workstation (SGI, Schlieren, Switzerland) with Imaris 3D multi-channel image

processing software installed (Bitplane, AG, Zurich, Switzerland (Messerli et al. 1993)).

Transmission Electron Microscopy

For electron microscopic studies spheroids were fixed by total immersion in 0.1M

cacodylate buffer (pH 7.4, 350 mOsm) containing 2.5% glutaraldehyde. Tissue blocks were

postfixed in osmium tetroxide, block-stained using uranyl acetate, dehydrated by increasing

ethanol concentrations and embedded in Epon 812 according to Djonov et al. (Djonov et al.

2000) (all chemicals from Merck Eurolab AG, Dietikon, Switzerland). Semithin 1 µm

sections were stained with toluidine blue and visualized using an Olympus Vanox BHS light

microscope (Olympus AG, Volketswil, Switzerland). Ultrathin sections of 80-90 nm

thickness were cut, picked up on Formvar-coated (polyvinyl formal; Fluka Chemie AG)

copper grids, double-stained with lead citrate (Merck Eurolab AG) and uranyl acetate, and

monitored on a Philips EM 400 electron microscope (FEI AG, Zurich, Switzerland).

ELISA-based VEGF quantification

VEGF production was quantified in the culture supernatants of confluent monolayers

and/or microtissue cultures using the DuoSET®

enzyme-linked immunosorbent assay

(ELISA) by R&D systems (Minneapolis, MN) according to the manufacturer’s instructions.

Results

VEGF production profiling of human cell-derived monolayer and

microtissue cultures

ELISA-based technology was used to quantify vascular endothelial growth factor

(VEGF) production by human cell lines and primary cells grown as monolayers or assembled

as microtissues. For unbiased VEGF profiling monolayer cultures were grown to confluence

and growth factor production compared to microtissues over a period of 24h. Whereas the

adult-derived human aortic fibroblasts (HAF adult; 0.21 ng/h*cell ± 0.025), the neoplastic cell

line HT-1080 (0.43 ng/h*cell ± 0.032) and the primary human articular chondrocytes (HAC;

1.21 ng/h*cell ± 0.055) produced considerable VEGF amounts, growth factor levels of


primary human dermal fibroblasts (NHDF), primary children-derived human aortic fibroblasts

(HAF child) and the human newborn foreskin fibroblast cell line Hs68 were undetectable

(Figure 1A). Scaffold-free microtissues derived from the same cell types were generated

using gravity-enforced self-assembly of monodispersed cells. Based on previous observations

suggesting cell type-specific cell number – microtissue size correlations we have used tailored

cell concentrations to obtain microtissues of 350 µm in diameter following a 3-day cultivation

period (Kelm et al. 2003). Whereas Hs68 spheroids failed to produce detectable VEGF levels,

NHDF- (0.09 ng/h*cell ± 0.028), HAF child- (0.27 ng/h*cell ± 0.028), HAF adult-

(0.92 ng/h*cell ± 0.026) and HT-1080- (1.55 ng/h*cell ± 0.231) derived microtissues showed

increased growth factor production compared to isogenic monolayer cultures. Only HAC

spheroids’ specific VEGF production (0.09 ng/h*cell ± 0.011) was inferior to corresponding

monolayer cultures (Figure 1B). For detailed analysis of microtissue size - VEGF production

profiles, gravity-enforced cell assembly of selected cell types was initiated using 500, 2’500,

5’000 and 10’000 cells per spheroid. Plotting of microtissue size vs. VEGF production

revealed cell type-specific correlations (Figure 2).


Figure 1: Quantification of vascular endothelial growth factor (VEGF) secretion of different human

primary cells (human aortic fibroblasts [HAF], human dermal fibroblasts [NHDF, Hs68], human articular

chondrocytes [HAC]), and the human fibrosarcoma cell line HT-1080 grown to confluent monolayers (A) or

scaffold-free microtissues of 350 µm in diameter (B).

Cell Type

VE

GF

[n

g/h

*ce

ll]

0.0

0.2

0.4

0.6

0.8

1.0

1.2

1.4

1.6

1.8

2.0

NHDF HAF ChildHs68 HAF Adult HT-1080 HAC

Monolayer

A

Hs68 NHDF HACHAF child HAF adult HT-1080

Cell Type

VE

GF

[n

g/h

*ce

ll]

Microtissue

0.0

0.2

0.4

0.6

0.8

1.0

1.2

1.4

1.6

1.8

2.0

B

Cell Type

VE

GF

[n

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h]

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HAF Child

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Spheroid Diameter [µm]

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/ml*

h]

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HAF Adult

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HepG2

Figure 2: Microtissue size – VEGF production profiles of primary human aortic fibroblast- (HAF, adult

and child), human fibrosarcoma cell- (HT-1080) and human hepatocellular carcinoma cell- (HepG2) derived

microtissues produced by 3-day gravity-enforced assembly in hanging drops.

Self-organization potential of different cell phenotypes in a microtissue

format

Blood vessels exhibit a particular architecture, with endothelial cells lining the blood

vessels and forming a cellular interface between the bloodstream and the surrounding tissue.

In order to investigate self-organization forces underlying cell movement during angiogenesis

we aggregated HAF:HUVEC (10’000:1’000 cells) and HepG2:HUVEC (1’200:600)

suspension cocultures to mixed microtissues in hanging drops. After 7 days, gravity-enforced

assembly resulted in HAF:HUVEC- and HepG2:HUVEC-derived microtissues of 800 µm and

350 µm in diameter, respectively. Most importantly, intra-microtissue forces separated

HAF/HepG2:HUVEC populations which resulted in concentric HAF/HepG2-inside:HUVEC-

outside structures reminiscent of blood vessel cross-sections (Figure 3A and B). By contrast,

human umbilical artery smooth muscle cell (HUASMC):HUVEC-derived microtissues

(600:600) were characterized by an inner HUVEC core enveloped by a HUASMC layer

(Figure 3C).


Figure 3: Immunohistological characterization of the self-organization potential of microtissues

produced by 7-day gravity-enforced assembly of mixed human aortic fibroblast cells (HAF): human umbilical

endothelial cell (HUVEC) (A), human hepatocellular carcinoma (HepG2):HUVEC (B) and human umbilical

artery smooth muscle cell (HUASMC):HUVEC (C) suspension cultures in hanging drops. HAF (A) and HepG2

(B) were visualized by F-actin-specific staining (red) and HUASMCs (C) by smooth muscle alpha-actin staining

(red), while HUVECs (A-C) were stained for von Willebrand factor (green). (scalebar = 20 µm).

VEGF profiling of microtissues assembled from different cell types

Based on increased VEGF production capacity of HAFs compared to similar-sized

NHDF microtissues we chose human aortic fibroblasts to exemplify VEGF production in pure

and multicellular primary human cell-derived spheroids (Figures 1B and 4A). Although

VEGF production was insignificant in supernatants of mixed HAF:HUVEC (10’000:1’200

cells) microtissues cultivated for 5 days, qualitative VEGF-targeted immunostaining revealed

that this vascular endothelial growth factor co-localized to HUVECs which had arranged to

enclose the inner HAF-containing microtissue core (Figure 4B-D). By contrast, pure HAF-

derived microtissues (10’000 cells) produced VEGF throughout the entire spheroid which

suggested HAF-based VEGF production to be compromised or consumed in self-organizing

microtissues assembled from mixed HAF:HUVEC co-cultures (Figure 4A). Modulation of

VEGF production following molecular crosstalk between different cell types is a well-

established phenomenon (Peirce and Skalak 2003; see below). Despite moderate but localized

VEGF production HUVECs failed to invade the HAF core and form capillaries, which is

typically a key step during angiogenesis (Figure 4).

A B CAA BB CC


Figure 4: Qualitative VEGF expression analysis of 5-days old pure human aortic fibroblast (HAF)-

derived microtissues and microtissues assembled from mixed HAF: human umbilical vein endothelial cell

(HUVEC) cultures. (A) VEGF-specific immunohistological staining (green) of pure HAF microtissues. (B-D)

HAF:HUVEC microtissues were stained for F-actin (blue, entire spheroid) (B), HUVECs were visualized by

Willebrand factor expression (red) and VEGF was monitored in green (C). (scalebar = 30 µm).

Angiogenesis-based capillary formation in microtissues

Initial experiments designed to characterize angiogenesis in hepatic microtissues were

based on HUVEC-coated HepG2 spheroids (HepG2 core – HUVEC shell; HepG2-HUVEC)

produced by cocultivating HepG2 spheroids (initiated with 500 cells), preassembled for 2

days in hanging drops (170 23 µm), with monodispersed HUVECs (900 cells). Since

HepG2 remain proliferation-competent in a microtissue format (Kelm et al. 2003), HepG2-

HUVEC and HepG2-only microtissues reached similar diameters of 400 50 µm after 7 days

post coating (Figure 5). Following invasion of the HepG2 core spheroids, HUVECs formed

multicell-based microvessels characterized by increased V 3 integrin expression typical for

A B

DC

AA BB

DDCC


neovascular endothelial cells (Brooks et al. 1994; Koh et al. 2004). By contrast, microvessel

formation could not be observed in HepG2-only control spheroids (Figure 5).

Figure 5: Capillary formation in hepatic microtissues. Coated HepG2-HUVEC (A-D) and pure

HepG2 (E, F) spheroids grown for 7 days in hanging drops. Toluidine blue-stained sections of HepG2-HUVEC

(A) and HepG2-only (E) microtissues. Immunohistologic von Willebrand factor-specific staining of endothelial

cells (dark brown) in HepG2-HUVEC (B, C) microtissues reveals HUVEC-based vessel formation (C, arrow;

see also Figure 9). Immunocytologic staining of entire cells for A633-coupled phalloidin (red) and V 3 integrin

(green) shows capillaries with peripheral V 3 integrin expression (arrows) in HepG2-HUVEC (D) but not in

HepG2-only microtissues. (scalebar = 20µm).

Instead of generating HAF:HUVEC microtissues by gravity-enforced assembly of

mixed HAF and HUVEC cultures (see above and Figure 4), we coated pure HAF microtissues

of different size (125 µm – 335 µm; produced by 2-day gravity-enforced assembly) by

cocultivation with 600 – 1’200 monodispersed HUVECs in hanging drops (Figure 6). Coated

HAF-HUVEC microtissues (HAF core - HUVEC shell) of different size were cultivated for 6

days prior to analysis of VEGF-induced HUVEC migration into the inner HAF core (Figure

6A-E). Whereas HUVECs failed to penetrate HAF core microtissues of 125 µm in diameter

(500 fibroblasts) (Figure 6A), these endothelial cells infiltrated HAF microtissues assembled

from 2’500 (225 µm in diameter) and 5’000 (285 µm in diameter) cells likely following the

B

D F

A

E

CBB

DD FF

AA

EE

CC


HAF-generated hypoxia-induced VEGF gradient (Figure 6B, C). In oversized HAF-HUVEC

microtissues (10’000 cells, 335 µm in diameter) HUVECs assembled into extensive capillary

structures inside the HAF core (Figure 6D). Interestingly, HUVEC-coated NHDF

microtissues (233 µm 12 µm) were able to induce HUVEC invasion and capillary formation

despite NHDF’s low VEGF production (Figure 6F).

Figure 6: Size-dependent induction of angiogenesis-based capillary formation in multicellular

microtissues. Human aortic fibroblast (HAF)-derived spheroids of different size/cell number ([A], 125/500; [B],

225/2’500; [C], 285/5’000; [D], 335/10’000, diameter [µm]/cells) were assembled as microtissues for 2 days,

coated with 300 (A), 600 (B), 1’000 (C) or 1’200 (D) monodispersed human umbilical endothelial cells

(HUVEC) and cultivated for 7 days in hanging drops. (E) Inset of (D) showing a branching capillary at higher

magnification. (F) NHDF-derived microtissue (233µm/10’000 cells) coated with HUVEC (600 cells). (A-F)

HUVEC-specific markers PECAM-1 (platelet-endothelial cell adhesion molecule-1; red) and von Willebrand

factor (green) were immunostained. (scalebar = 20 µm).

Detailed HUVEC migration kinetics of HAF-HUVEC microtissues revealed that

HUVECs (i) assembled on the surface of preformed HAF spheroids in a shell-like manner 2

days post coating (Figure 7A), (ii) HUVECs started infiltrating the HAF core spheroid at day

4 (Figure 7B) and (iii) formed a well-structured capillary system on and beyond day 6 post

CA B

D FE

CCAA BB

DD FFEE


coating (Figure 7C). Interestingly, VEGF secretion of HAF-HUVEC microtissues was lower

compared to pure HAF spheroids suggesting that either crosstalk between HAFs and

HUVECs is modulating VEGF secretion or VEGF is bound/taken up by HUVECs (Figure 8).

Figure 7: Kinetics of angiogenesis-dependent capillary formation in human umbilical endothelial cell

(HUVEC, 600 cells)-coated human aortic fibroblast (HAF; 335µm/10’000 cells) spheroids. HUVEC-specific

staining of PECAM-1 (platelet-endothelial cell adhesion molecule-1, red) and von Willebrand factor (green) on

days 2 (A), 4 (B) and 6 (C) post coating. (scalebar = 30 µm).

A B CAA BB CC

A

Time [h]

VE

GF

[p

g/m

l]

100

300

500

700

900

48 96 144 192

HAF

HAF-HUVEC

A

Time [h]

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l]

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300

500

700

900

48 96 144 192

HAF

HAF-HUVEC

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HAF-HUVEC

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HAF-HUVEC


Figure 8: VEGF production kinetics of human umbilical vein endothelial cells (HUVEC)-coated on

human aortic fibroblast (HAF) spheroids and pure HAF-derived spheroids over 6 days. 125 µm (A, 500 cells) or

335 µm (B, 10’000 cells) HAF-derived spheroids optionally coated with 600 HUVECs.

Ultrastructural characterization of HAF-HUVEC microtissues 7 days post coating

demonstrated advanced development of vascular structures. In early developmental stages

many endothelial cells are characterized by intracellular lumen formation (Figure 9A-C).

Endothelial cells increase their lumen size, loose organelles and progressively attenuate

thereby adopting morphologies of the capillary endothelium (Figure 9C). At an advanced

stage of differentiation HUVECs exhibit a well-developed vesicular system which may

assemble to form typical transendothelial transport channels (Figure 9D). Similar to native

vessels HAFs and HUVECs develop a panoply of cell-cell interactions (Figure 9E).

BV

EG

F [

pg

/ml]

Time [h]

5000

15000

25000

35000

48 96 144 192

HAF

HAF-HUVEC

BV

EG

F [

pg

/ml]

Time [h]

5000

15000

25000

35000

48 96 144 192

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VE

GF

[p

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l]

Time [h]

5000

15000

25000

35000

48 96 144 192

HAF

HAF-HUVEC

HAF

HAF-HUVEC

HAF

HAF-HUVEC


Figure 9: Ultrastructure of cocultured HAF-HUVEC spheroids 7 days post coating. (A) Endothelial

cells are typically characterized by intracellular lumen formation (+). Endothelial cells are tightly covered by

human aortic fibroblasts (asterisk). The periendothelial space is indicated by arrowheads. (B) Capillary like-

structures result from fusion of endothelial cell (asterisks) protrusions. (C) Advanced differentiation stages are

characterized by enlarged lumina and organelle-free attenuated endothelial cells. Boxed regions are enlared in D

and E. (D) Vesicles typically open to the luminal (lv) and abluminal (av) sites or form transendothelial channels

(ch). Tight junctions (tj) and zonula adhaerens (za) demonstrate well-established endothelial-periendothelial cell

contacts. (E) The periendothelial space contain fibrils (asterisks) and cellular protrusions which often result from

HAF-HUVEC cell contacts (arrowheads).

A

E

DC

BA

E

DC

B


Inhibition of angiogenesis in HAF-HUVEC microtissues

The C-terminal cleavage product of collagen XVIII known as endostatin is a key anti-

angiogenic factor (Bloch et al. 2000; O'Reilly et al. 1997). We evaluated endostatin action on

HAF microtissues (10’000 cells) coated with 1’200 HUVECs. Endostatin (1 g/ml) was either

administered during or 2 days post coating. 6 days post coating the HAF-HUVEC

microtissues were immunoprofiled for PECAM-1 and von Willebrand factor expression.

Whereas endostatin was able to prevent angiogenesis when supplied during coating, later

addition of this antiangiogenic factor to mature HAF-HUVEC microtissues failed to inhibit

HUVEC-based capillary formation in the HAF core (Figure 10B and 10C). Similar to addition

of endostatin, HAF spheroids coated with a mixture of HUVECs and human umbilical artery

smooth muscle cells (HUASMC) (HAF-HUVEC/HUASMC; 10’000-600/600 cells) did not

develop any HUVEC-based capillary systems confirming the potential of HUASMCs to

inhibit VEGF-mediated vascularization (data not shown) (Peirce and Skalak 2003).

Figure 10: Endostatin-mediated angiogenesis suppression in human aortic fibroblast (HAF)

microtissues coated with human umbilical vein endothelial cells (HUVEC). Endostatin impact on HUVEC-

driven capillary formation in HAF microtissues was monitored following administration of this anti-angiogenic

factor at coating (A) as well as 2 days post coating (B). Untreated HAF-HUVEC microtissues were used as

control (C). HAF-HUVEC spheroids were cultivated for 6 days and vascularization was analyzed by PECAM-1-

(red) and von Willebrand factor- (green) specific staining. (scalebar = 20µm).

Discussion

Endothelial cells are the central organizational unit of (micro-)vascular structures.

Their lineage commitment, expansion, organization, and assembly into ordered and tissue-

specific interconnecting vascular structures are required for embryonic development,

CBA CCBBAA


organogenesis, wound healing, reproductive tissue cycles, tumorigenesis and a number of

other pathological conditions involving inflammation (Daniel and Abrahamson 2000). The

elucidation of the molecular basis of tissue-specific angiogenesis remains largely elusive,

owing to the panoply of complex interactions which have to be temporally and spatially

coordinated (Eliceiri and Cheresh 2001). In order to obtain new insight into

neovascularization of human adult tissues, bioengineered tissues or development of tumor

vessels 3D, angiogenesis models are the preferred systems to study complex cell-cell

interactions and signal integration circuits managing the balance of pro- and anti-angiogenic

cellular activities in a clinically relevant manner (Hirschi et al. 1999; Niklason et al. 1999;

Vailhe et al. 2001).

We have refined the hanging drop technology to generate fully size-controlled

multicellular tumor spheroids (MCTS)/microtissues of a wide variety of different cell types

which exhibited size/cell number-dependent expression of the vascular endothelial growth

factor (VEGF) [31, 46]. Microtissue size-dependency of VEGF expression is a result of

hypoxic conditions at the center of oversized spheroids which induce HIF-1 a transcription

factor controlling VEGF production (Semenza 2001). Although VEGF can induce mature

vessels at an appropriate dosage and appears to have a relatively wide therapeutic window,

the balance between clinical benefit and pathologic side effect is likely to differ with genetic

predisposition, age and disease status (Blau and Banfi 2001). The tumor cell used in this study

(HT-1080) produced increased VEGF levels in monolayers compared to isogenic cultures of

primary human mesodermal cells. Moreover, MCTS exclusively assembled from HT-1080

cells exhibited a near linear cell number-VEGF production profile while expression of this

growth factor reached a plateau in microtissues composed of 2’500 to 5’000 non-tumorigenic

cells (HAF, NHDF). VEGF expression under normoxic conditions in monolayer cultures and

the linear correlation between cell number-VEGF secretion indicates increased hypoxia-

independent VEGF production in tumor cells included in this study. Although absolute

VEGF/microtissue size profiles of different human primary cells revealed cell type- and

donor-specific differences their relative levels remained comparable. As an exception, VEGF

expression profiles of HAC microtissues were decreased compared to control monolayer

cultures which supported findings by Schipani and coworkers suggesting HIF-1 -independent

VEGF induction in chondrocytes (Schipani et al. 2001).

An ideal bioengineered tissue will have to be assembled from multiple organ-specific

cell types, while retaining hierarchical cellular architecture and supporting

endovascularization compatible with connection to the native vascular systems of organs.


Despite induction of angiogenesis by growth factors, 3D self-organization of different cell

types is a prerequisite for engineering of functional and fully vascularized tissue equivalents.

Mixed cultures of HepG2:HUVEC, HAF:HUVEC and HUASMC:HUVEC had shown a high

degree of self-organization mimicking in vivo cellular architecture: HUVECs moved to the

periphery of the microtissue building the barrier to the surroundings. Interestingly, mixed

HUASMC:HUVEC microtissues exhibited the opposite HUVEC-inside/HUASMC-outside

setup which contrasts previous HUVEC-outside/HUASMC-inside organotypic cultures

assembled using liquid methylcellulose. These findings exemplify modulation of self-

organization forces within microtissues by culture additives/matrices (Korff and Augustin

1999; Korff et al. 2001).

The interplay between environmental and genetic impact on tumor

angiogenesis/proliferation is complex and remains largely elusive (Carmeliet and Jain). A

major challenge is to establish a generic cell culture model mimicking tumor development

and angiogenesis of a wide variety of tumorigenic cell types (Kelm and Fussenegger 2004;

Kelm et al. 2003). Although all cancer cell lines used in this study produced VEGF, we have

chosen HepG2 as tumor model since hepatocellular carcinomas are among the most malignant

cancers affecting over 1 million patients per year (Kim et al. 2002). Although HepG2-derived

MCTS produced little VEGF compared to other 3D-assembled cancer cell lines (DU-145,

MCF-7, HT-1080), VEGF production was sufficient to induce vascularization in coated

HepG2-HUVEC microtissue and reduced core necrosis compared to pure HepG2-derived

MCTS.

The development of cell-based therapies to assist/replace diseased tissue is among the

most promising initiatives in regenerative medicine (Petit-Zeman 2001). Yet, a major

challenge on our way towards the design of artificial organs is vascularization management to

provide oxygen/nutrient supplies and removal of waste products (Griffith and Naughton 2002;

Zandonella 2003). Precise knowledge of the impact of tissue microenvironments on

metabolism, proliferation and growth-factor production will be essential for elucidation of

microvessel formation (McIntire 2002). Although often declared an engineering target, VEGF

production can be fine-tuned by multiple cell types arranged in 3D structures in a self-

sufficient manner (Elson et al. 2001; Isner 2002; Jain and Carmeliet 2001; Richardson et al.

2001; Zisch et al. 2003). HAF-HUVEC and NHDF-HUVEC microtissues have exemplified

that a high angiogenic potential can be induced by controlled cell assembly without

administration of any growth factor. 6 days post coating, cellular crosstalk combined with

hypoxia-induced VEGF production established a dense microvessel network, as a


consequence of which VEGF production decreased significantly. Interference of

HUASMC:HUVEC coated onto HAF spheroids abolished capillary formation (data not

shown) which was reminiscent of HUASMC-mediated repression of vascularization in vivo

(Sato and Rifkin 1989). Capillary formation in HAF-HUVEC cultures could also be

prevented by addition of the well-known antiangiogenic factor endostatin, but only when

supplied at the time of coating. Interestingly, 2 days post coating the angiogenic machinery of

HAF-HUVEC microtissues was set for capillary formation and could no longer be impaired

or reversed by endostatin.

Microtissue design by gravity-enforced assembly of monodispersed cells in hanging

drops enables (i) precise size control, (ii) flexible cell-type composition, (iii) self-organization

of multiple cell types, (iv) intra-/inter-cellular crosstalk, (v) in vivo-mimicking of cellular

architecture and (vi) self-sufficient and auto-controlled vascularization of oversized spheroids.

Owing to the low cell number required for production of microtissues, this 3D culture system

is expected to be compatible with high-throughput drug screening and multi-level gene

function analysis. Also, with their auto-angiogenesis potential, microtissues could represent

the optimal format for (autologous) cell therapies, a vision which will have to be substantiated

by further in vivo studies.

Acknowledgments

We thank, Cornelia C. Weber, as well as Beat P. Kramer for critical comments on the

manuscript, Jean-Claude Perriard for his ongoing support, Millenium Biologix for providing

human articular chondrocytes and Heike Hall for supply of anti-integrin antibodies. Work in

the laboratory of M.F. is supported by the Swiss National Science Foundation (grant no. 631-

065946) and a special research grant by the ETH Vice President for Research.

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survival." Genes Dev 15, 2865-2876.

Schuch G., J.V. Heymach, M. Nomi, M. Machluf, J. Force, A. Atala, J.P. Eder, J. Folkman, S.

Soker (2003). "Endostatin inhibits the vascular endothelial growth factor-induced

mobilization of endothelial progenitor cells." Cancer Res 63, 8345-8350.

Semenza G.L. (2001). "HIF-1, O(2), and the 3 PHDs: how animal cells signal hypoxia to the

nucleus." Cell 107, 1-3.

Semenza G.L. (2003). "Angiogenesis in ischemic and neoplastic disorders." Annu Rev Med

54, 17-28.

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Springer M.L., A.S. Chen,P.E. Kraft, M. Bednarski, H.M. Blau (1998). "VEGF gene delivery

to muscle: potential role for vasculogenesis in adults." Mol Cell 2, 549-558.

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beyond." Expert Opin Investig Drugs 12, 933-941.


Vailhe B., D. Vittet, J.J. Feige (2001). "In vitro models of vasculogenesis and angiogenesis."

Lab Invest 81, 439-452.

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"Vascular-specific growth factors and blood vessel formation." Nature 407, 242-248.

Zandonella C. (2003). "Tissue engineering: The beat goes on." Nature 421, 884-886.

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VEGF from synthetic, biointeractive cell ingrowth matrices for vascularized tissue

growth." Faseb J 17, 2260-2262.

Chapter 6

Improved Tissue-Transplant Fusion and

Vascularization of Myocardial Micro- and

Macrotissues Implanted into Chicken Embryos and

Rats

Kelm J.M., Djonov V., Hoerstrup S.P., Guenter C. I., Ittner L.M., Greve F., Hierlemann A.,

Perriard P.C., Ehler E. and Fussenegger M. (submitted)

Myocardial Microtissue Implants 101

Abstract

Cell-based therapies and tissue engineering initiatives are gathering clinical momentum

for next-generation treatment of tissue deficiencies. Capitalizing on gravity-enforced self-

assembly of monodispersed primay cells in hanging drops we have produced adult and neonatal

rat cardiomyocyte-based myocardial microtissues which could optionally be vascularized

following coating with human umbilical endothelial cells (HUVEC). Within myocardial

microtissues, individual cardiomyocytes showed native cell shape and morphologies and

established electrogenic coupling via intercalated disks. This resulted in the coordinated

contracting microtissues, which was recorded by means of a novel multi-electrode CMOS

microchip. Myocardial microtissues ( m3 scale), coated with HUVECs and cast in a custom-

shaped agarose mould, assembled to coherent macrotissue (mm3 scale), characterized by an

extensive capillary network with typical vessel ultrastructures. Following implantation into

chicken embryos, myocardial microtissues produced the vascular endothelial growth factor

(VEGF), which recruited the embryo’s capillaries to functionally vascularize the rat-derived

tissue implant. Similarly, transplantation of rat myocardial microtissues into the pericardium of

adult rat resulted in time-dependent resorption of myocardial microtissues and co-alignment of

implanted and host cardiomyocytes within seven days. Myocardial microtissues and custom-

shaped macrotissues, which are either vascularized in vitro or recruit the host vasculature

following in vivo implantation, exemplify the potential of artificial tissue implants for

regenerative medicine.

Introduction

The heart is the first organ to form in the embryo and all subsequent activities in life

depend on its function. The past decade has witnessed decisive advances in understanding cardiac

function and dysfunction, both genetically and molecularly levels. Although such insights into

the mechanisms of heart development and disease have stimulated new therapeutic opportunities

for the prevention and palliation of cardiac pathogenesis, mortality rates associated with heart-

related pathologies remain at the top of disease statistics in industrialized countries (Olson 2004).

Since cardiac myocytes lose their ability to divide after birth, the regenerative capacity of adult


heart tissue is limited, and substantial cell loss or dysfunction, such as occurs during myocardial

infarction, is largely irreversible and may lead to progressive heart failure (Pasumarthi and Field

2002).

Transplantation of excitable myogenic cells within the dysfunctional zone is a possible

therapy for restoring cardiac function. For example, initial studies with injected skeletal muscle

cells showed promise; this strategy is, however, unlikely to be suitable for long-term therapy

owing to the failure of the injected cells to become electrically coupled to the heart and to

different contractile properties between cardiac and skeletal muscle cells (Murry et al. 2004).

Although several cell phenotypes have been tested, cardiomyocytes emerged as the preferred cell

type because of their inherent structural, electrophysiological and contractile properties (Reinecke

et al. 1999). However, clinical applications are hampered by the paucity of cell sources for

human cardiomyocytes and by the limited evidence of direct functional integration of host and

donor cells (El Oakley et al. 2001; Rubart et al. 2003; Olson 2004). Nevertheless, injection of

bone marrow and stem cells derived from cloned mouse embryos into the border zone of the

infarcted myocardium have been reported to promote cardiac regeneration and improve cardiac

function in animal models (Kehat et al. 2004; Lanza et al. 2004). However, there is disagreement

as to the efficiency with which exogenous stem cells can colonize the heart and adopt a

cardiomyocyte cell fate (Kehat et al. 2004; Murry et al. 2004; Nygren et al. 2004). Recently,

human embryonic stem cell-derived cardiomyocytes successfully paced the ventricle in swine

with complete heart block, showing that transplanted cardiomyocytes can in principle survive,

function and integrate with host cells and operate as a biological alternative to the electronic

pacemaker (Kehat et al. 2004). However, embryonic stem cells can form teratomas in vivo which

presents another set of therapeutic challenges.

To date, cardiomyocyte-based prototype cell therapy initiatives capitalized on the

transplantation of fetal, neonatal (NRCs) and adult (ARCs) rat cardiomyocytes (Reinecke et al.

1999). While ARCs died 24h after transplantation, 50% of grafted NRCs survived the first day

but more than 90% perished within the first week (Zhang et al. 2001). Despite several attemps to

limit the death of grafted cell suspensions by antiapoptosis engineering (Datta et al. 1999),

localized high-dose application of single cells remains technically challenging. Therefore, pre-

transplantation assembly of monodispersed cells to tissue-like structures is considered to be an

alternative. A variety of scaffold-containing (for example, collagen and ceramics) and scaffold-


free (for example, stacking of monolayers and reaggregation) strategies have been designed to

produce three-dimensional cardiac tissue constructs, some of which have been transplanted

successfully (Shimizu et al. 2002; Zimmermann et al. 2002; Zandonella 2003; Zimmermann and

Eschenhagen 2003; Kelm et al. 2004).

The potential hurdles in cardiac tissue engineering are the requirements that

cardiomyocyte-derived artificial tissues become seamlessly integrated into the damaged

myocardium without becoming a substrate for arrhythmogenesis, and that they be able to induce

neovascularization of the regenerated myocardium. Capitalizing on gravity-enforced self

assembly of primary cardiomyocytes in hanging drops we recently reported the production of

highly adhesive and functionally beating myocardial microtissues. They produced the vascular

endothelial growth factor in a size-dependent manner, suggesting that they may stimulate

angiogenesis after transplantation (Kelm et al. 2004). In this study we (i) pioneered production of

myocardial microtissues from adult rat cardiomyocytes, (ii) designed a novel CMOS microchip

for assessing of electrongenic microtissue performance, (iii) assembled myocardial microtissues

to larger fully vascularized tissue implants using custom-shaped agarose moulds, which (iv)

managed successful vascularization crosstalk following the grafting of myocardial microtissues

into chicken embryos and (v) provided seamless integration after implantation into the

pericardium of adult rats. We are convinced that microtissue-based scale-up, resulting in fully

vascularized, custom-shaped, scaffold-free and transplantation-ready tissue units will improve

cell-based cardiac-related therapies in the not-too-distant future.


Preparation of primary cells

Neonatal rat cardiomyocytes (NRCs) were isolated from dissected newborn rat (Wistar;

Janvier Elevage, Le Genest Saint Isle, France) hearts by digestion with collagenase (Worthington

Biochemical Corp., Freehold, NJ) and pancreatin (Invitrogen, Carlsbad, CA, USA), according to

protocols by Auerbach and colleagues (Auerbach et al. 1999). Adult rat cardiomyocytes (ARCs)

were prepared as described elsewhere (Eppenberger and Zuppinger 1999). NRCs and ARCs were

kept in plating medium (67% Dulbecco’s Modified Eagle Medium [DMEM; Invitrogen], 17%


M199 [Amimed AG, Basel, Switzerland], 10% horse serum [cat. no. 16050-098, lot no.

3036354D, Invitrogen], 5% fetal bovine serum [FBS; cat. no. 3302-P231902, lot no. P231902;

PAN Biotech GmbH, Aidenbach, Germany], 1% penicillin/streptomycin solution [Invitrogen]).

Human umbilical vein endothelial cells (HUVECs, PromoCell, Heidelberg, Germany) were

expanded in endothelial cell growth medium (PromoCell, cat. no. C-22010) supplemented with

10% FBS (PAN Biotech GmbH, Aidenbach).

Microtissue production

After isolation, cardiomyocytes were seeded at indicated cell concentrations and cell-type

compositions into 60-well plates (HLA plate, Greiner-Bio One, Frickenhausen, Germany). To

enable gravity-enforced self-assembly of microtissues in hanging drops, the 60-well plates were

incubated upside down at 37°C in a humidified atmosphere containing 5% CO2.

Macrotissue assembly

In order to assemble microtissues to larger-sized artificial macrotissues we designed

cylindrical agarose casting moulds, 3 mm in diameter. Negative Teflon®

casting moulds were

filled with 4% agarose (Sigma Chemicals, Buchs, Switzerland) in phosphate-buffered saline

(PBS; 150 mM NaCl, 6.5 mM Na2HPO4 x 2 H2O, 2.7 mM KCL, 1.5 mM KH2PO4, pH 7.4) to

generate non-adhesive positive casting moulds to imprint the desired shape on the macrotissue.

Microtisses of indicated cell types and cell numbers were transferred to the desired agarose

moulds and cultivated in cell type-specific media under static conditions at 37°C in a humidified

atmosphere containing 5% CO2.

Immunofluorescence analysis

Microtissues were washed once in PBS (Sigma Chemicals) and fixed for 4h at 4°C in

PBS containing 4% paraformaldehyde (Sigma Chemicals). Immunofluorescence-based analysis

of microtissues was performed as described elsewhere (Kelm et al. 2004) using primary

antibodies specific for sarcomeric- -actinin (mouse monoclonal, Sigma Chemicals; clone EA53),

connexin-43 (rabbit polyclonal, Zymed, San Francisco, CA, USA), or myomesin (mouse

monoclonal (Grove et al. 1984)) was used and stained with FITC-coupled secondary anti-mouse


(Jackson Immunochemicals, West Grove, PA, USA) or Cy3-coupled anti-rabbit (ICN

Pharmaceuticals, Hyland, CA) antibodies. Cell nuclei were stained with 1 g/ml 4',6-diamidino-

2-phenylindole (DAPI, Molecular Probes Inc., Eugene, OR, USA).


The imaging system consisted of an inverted fluorescence microscope equipped with 20x

or 10x oil immersion objectives and a confocal scanner (Zeiss LSM510; Carl Zeiss AG,

Feldbach, Switzerland) with argon and helium neon lasers installed. Images were processed using

up-to-date Zeiss software (Carl Zeiss AG).

Transmission electron microscopy

Tissue samples were fixed by immersion in 0.1M cacodylate buffer (pH 7.4, 350 mOsm)

containing 2.5% glutaraldehyde. Tissue blocks were postfixed in osmium tetroxide, block-stained

using uranyl acetate, dehydrated by sequential incubation in increasing ethanol concentrations

and embedded in Epon 812 according to Djonov and coworkers (Djonov et al. 2000) (all

chemicals from Merck Eurolab AG, Dietikon, Switzerland). Semi-thin 1 µm sections were

stained with toluidine blue and visualized by an Olympus Vanox BHS light microscope

(Olympus AG, Volketswil, Switzerland). Ultra-thin sections of 80-90 nm were cut with a

diamond knife and picked up on Formvar-coated (polyvinyl formal; Fluka Chemie AG) copper

grids, double-stained with lead citrate (Merck Eurolab AG) and uranyl acetate and monitored on

a Philips EM 400 electron microscope (FEI AG, Zurich, Switzerland).

Microchip-based electrophysiology

The 5 5 mm2 bio-electronic chip has been designed as an array of 4 4 platinum

electrodes (20 20 µm2). An individual circuitry block manages signal conditioning of a single

four-electrode array, which enables simultaneous recording in the absence of multiplexer

switching. Electrode-recorded cell signals are high-pass filtered (100 Hz cut-off) to prevent input

saturation by signal differences between reference and measurement electrodes, intensified (gain

10x, 100x) by a programmable gain amplifier in the circuitry block, low-pass filtered (4 KHz

adjustable cut-off) and digitalized by external circuitries (Greve et al., unpublished). Microtissues


(see above for assembly) were prepared for electrogenic recording by cultivating them for six

days in non-adhesive cell culture dishes (Greiner-Bio One). Myocardial microtissues (10 days

old) were placed on the electrodes and incubated at 37°C and 5% CO2. Within 2 h, myocardial

microtissues attached to the platinum electrodes; contraction was initiated by adding of 10-4

mM

of the -adrenergic agent phenylephrine (Sigma Chemicals) before recording of microtissue

signals.

Chicken chorioallantoic membrane (CAM) assay

Chicken embryos were cultured using the shell-free method (Ribatti et al. 2001). Artificial

tissues were grafted atop the growing chicken chorioallantoic membrane (CAM) on embryonic

day 9 and cultured for two days.

Transplantation of myocardial microtissues into rat hearts

Prior to transplantation, myocardial microtissues were labeled for 30 min in Hanks’

balanced salt solution (HBSS, Sigma Chemicals) supplemented with 10 mM CellTrackerTM

(CMTMR, Molecular Probes) and then washed three times in HBSS. Adult rats were treated with

the analgesic Temgesic (300 µl prior to surgery; Essex Chemie AG, Luzern, Switzerland) by

intraperitoneal injection and then anesthetized in an isoflurane chamber. Anesthetized rats were

endotracheally intubated and ventilated with positive-pressure (0.6 l/min, 3 ml tidal volume,

oxygen-supplemented (2 ml) air) using a Harvard ventilator (Harvard Apparatus, Hollisten,

Massachusetts, USA). Under general anesthesia a 1 cm long incision was made directly posterior

to the xiphoid. The chestwall was lifted up in order to expose the diaphragm. Myocardial MTs

were injected into the pericard through the membraneous part of the diaphragm. The injection

hole was sealed with fibrin glue (Tissucolduo, Baxter, Unterschleissheim, Germany). The

abdominal incision was closed in layers with 4-0 silk running sutures. After surgery, the rats were

kept warm and in isolation until they were fully awake when they were returned to their filter

cages. Postoperatively, the rats were injected with decreasing intraperitoneal Temgesic every six

hours for two days. Rats were sacrificed after one (n= 4), four (n=4) and seven days (n=4). The

control group was injected with HBSS and was sacrificed after seven days (n=2).


Results

Microtissues assembled from adult cardiomyocytes

Although several cell types are currently considered for myocardial tissue repair (Hassink

et al. 2003), only cardiomyocytes form intercalated disks, which are essential for

electrophysiological coupling. In contrast to neonatal cells, which will probably raise ethical

questions, allogenic adult cardiomyocytes are routinely isolated from the auricular cordis and

represent an alternative cell source for future cardiac tissue engineering. However, cultivation is a

challenge, which has prevented the 3D assembly of adult rat cardiomyocytes (ARCs), and

monodispersed ARCs implanted into rat hearts failed to survive for more than 48 h (Reinecke et

al. 1999). Using gravity-enforced self-assembly we produced artificial myocardial microtissues

after a one-week cultivation of monodispersed ARCs in hanging drops (Figures 1A and B).

Within the myocardial microtissues, ARCs adopted a native rod-shaped cell phenotype,

established inter-cardiomyocyte contacts via gap junctions (Figure 1C) and produced the vascular

endothelial growth factor (Figure 1D), similar to NRC-derived microtissues (Kelm et al. 2004).

After 14 days in culture (seven days in hanging drops, followed by seven days in standard

culture) ARCs at the microtissues’ periphery had aligned and were functionally interconnected

via intercalated disks (Figure 1G), while VEGF was produced throughout the entire microtissue

(Figure 1H). These results exemplify that adult cardiomyocytes arranged in a scaffold-free

microtissue retain their native cellular reorganization capacity as well as their angiogenic

potential.


Figure 1. Immunohistologic characterization of myocardial microtissues produced by gravity-enforced self-

assembly of adult rat cardiomyocytes (ARCs) for 7 (A-D) and 14 (E-H) days in hanging drops. ARC morphology

was visualized by immunohistologic staining specific for sarcomeric -actinin (red; A-F, D, H), DAPI (blue; A, E),

7d 14d

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-catenin (green; B, F), myomesin (red; C, G), connexin 43 (green; C, G) and vascular endothelial growth factor

(VEGF; green; D, H).

Microchip-based electrophysiologic analysis of myocardial microtissues

To assess inter-cardiomyocyte crosstalk and coordination of electrogenic activity across

NRC-derived microtissues we used metal oxide semiconductor (CMOS) microchip technology

(Greve et al., unpublished). The CMOS chip was designed as a four-block, 20 20 µm2 array,

each containing four platinum electrodes, a circuitry block harboring a high-pass filter and a

programmable amplifier as well as a central external low-pass filter and digitalization units

(Figures 2A and B). Using this CMOS microchip technology we recorded the electrogenic

activities of myocardial microtissues (assembled from 10,000 NRCs for 10 days) for two hours

following phenylephrine-mediated contraction stimulation using the -adrenergic agent

phenylephrine. Although electrogenic signals, recorded by extracellular electrodes represent local

changes in the membrane potential of the individual cardiomycocyte the overall signal is

modulated by the electrically coupled cell quorum (Hescheler et al. 2004). Starting with 0.25 mV

the recorded current increased to 1.75 mV, probably reflecting the initial beating of few

cardiomyocytes which spreads throughout the microtissue and results in coordinated contractions

(Figure 2C). This observation confirms electrophysiologic coupling among cardiomyocytes

embedded in an artificial microtissue structure.


Figure 2. Microchip-based electrophysiology. The 5 5 mm2 CMOS bio-electronic microchip, mounted on a

ceramic dual-inline (DIL) package (A), was designed as an array of 4 4 platinum electrodes (20 20 µm2),

surrounded by a nitride passivation area and a reference electrode. External circuitries manage signal processing and

chip control. Each four-electrode block harbors a circuitry unit for signal conditioning, which enables the

simultaneous readout of the four electrodes (B). Electrogenic microchip readout of a myocardial microtissue

assembled from 10,000 neonatal rat cardiomyocytes, cultivated for 10 days, placed and maintained for 2 h on an

individual electrode and stimulated by addition of the -adrenergic agent phenylephrine (10-4 mM) (C).

Design and neo-vascularization of higher-order macrotissues assembled from

individual myocardial microtissues

Since inter-microtissue interactions are expected to be similar to graft-host connection,

underlying molecular forces can be exploited to assemble macrotissue supra-structures from

individual microtissues. In designing larger-sized artificial tissue structures for clinical

applications, vascularization, necessary to sustain metabolic activities of the tissue core, will be

A B

C

A B

C


the prime size-limiting parameter. Previous studies exemplified in-vitro vascularization of human

aortic fibroblast-based core microtissues, whose VEGF production recruited peripheral human

umbilical endothelial cells (HUVEC) to develop a capillary system (Kelm et al. 2005). In order to

produce vascularized myocardial macrotissues from individual microtissues, 300 myocardial

microtissues, assembled from 10,000 NRCs (four-day assembly of core spheroid) coated with

1,200 HUVECs (five days coating in hanging drops), were cast in a custom-designed cylindrical

4% agarose mould, 3 mm in diameter. The agarose mould, produced from a Teflon®

mould,

provided an adhesion-free shape constraint for inter-microtissue crosstalk. Microtissues were

allowed to aggregate in agarose moulds for four days and the resulting macrotissues were kept

for another four days in non-adhesive agarose-coated culture dishes. During maturation, NRC-

HUVEC microtissues assembled into coherent macrotissues characterized by a peripheral layer

of HUVECs exhibiting their typical longitudinal cell morphology (Figure 3A-C). NRC-only

control macrotissues were less coherent and exhibited ruffled tissue structures, probably the

result of apoptosis progression evidenced by nuclear chromatin condensation, apoptotic bodies

and cytoplasm vacuolization (Figures 3E-G). Ultrastructural analysis revealed two types of

developing capillaries but only in HUVEC-containing NRC macrotissues (Figure 3G, H): (i)

tubular structures shaped by two or more endothelial cells connected by intercellular junctions

(Figure 3G) and (ii) lumen formation by a single endothelial cell, which corresponds to a

seamless capillary (Figure 3H).


Figure 3. Ultrastructural analysis of macrotissues assembled from HUVEC (1,200 cells)-coated NRC

microtissues (A-C) and pure NRC-derived microtissues (300; 10,000 NRCs each) (D-H) by cultivation in a

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cylindrical agarose mould (3 mm in diameter) for four days followed by four-days in non-adhesive culture dishes.

Toluidine blue staining of macrotissue sections reveal that pure NRC macrotissues are less coherent and develop

vacuoles (arrows) and apoptotic bodies (arrowheads) (D-H). In contrast, macrotissues produced from HUVEC-

coated NRCs show assembly of HUVECs at the macrotissue’s periphery (A, B) as well as development of capillaries

(C, arrow). Transmission electron micrographs NRC-HUVEC macrotissues reveal developing capillaries

characterized by lumen structures initiated within individual HUVECs (asterisk; arrow indicates lumen) (G) as well

as tubular lumen structures shaped by two or more HUVECs (asterisk; arrows indicate junctions) (H).

Inter-species vascularization crosstalk enables connection of myocardial

microtissues to the chicken embryo vasculature

Connection of artificial tissues to the host vasculature is essential for successful tissue

engineering (Nomi et al. 2002). We have previously reported that rat-derived myocardial

microtissues, exceeding 230 m in diameter (corresponding to 10,000 cells), show hypoxia-

induced VEGF production, which is expected to be one of the key factors for host tissue-

mediated vascularization of artificial tissue implants (Kelm et al. 2004). To study microtissue-

host vascularization crosstalk we implanted NRC-derived myocardial microtissues atop the

chorioallantoic membrane (CAM) of chicken embryos. Morphological analysis at the

microtissue/CAM interface revealed complete integration of the NRC-derived microtissue into

the chicken CAM (Figure 4A). Following a well-evolved vascularization scheme chicken and rat

cells inter-operated seamlessly to establish a fully functional joint vasculature (Figure 4B-C).

Microtissue-based VEGF production was sufficient to recruit the chicken embryo’s vasculature

to invade and fully vascularized the microtissue graft. Ultrastructural analysis of microtissue

implants revealed mature pericyte-covered microvessels (Figure 4E, F). Efficient recruitment of

host vascularization by microtissue-mediated VEGF production suggests that prevascularization

of artificial tissue implants may be dispensable for some transplantation scenarios.


Figure 4. NRC-derived myocardial microtissues (10’000 NRCs/microtissue; 10 days old) were implanted

and cultivated for 48 h atop the chorioallantoic membrane (CAM) of embryonic chicken embryos (day 10).

Toluidine blue staining of microtissue-CAM sections reveals invasion of microtissues into the chicken embryo

(asterisks; A-D). Erythrocyte (Er)-containing microvessels (indicated by arrows) from the chicken embryo invade

and vascularize the microtissue implants (B-D). Transmission electron microscopy reveals that microvessels (E) and

capillaries (F) are covered with pericytes (Pe), which indicates development of mature vessels.

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Integration of implanted myocardial microtissues into rat hearts

To characterize in-vivo integration of myocardial microtissues into the heart of adult rats

and evaluate their potential as minimal therapeutic tissue engineering units we injected NRC-

derived microtissues into the pericardial cavity. Myocardial microtissue implants were produced

by gravity-enforced self-assembly of 2,500 monodispersed NRCs for four days in hanging drops

and stained with a fluorescent CellTrackerTM

to trace tissue implants in the animal (Zhang et al.

2001). Implantation required a surgery (15 min); a 10 mm incision was made behind the xiphoid

of anesthetized adult rats followed by chest wall lift-up, injection of 300 microtissues (in 200 l

HBSS) into the pericardial cavity, sealing of the puncture with fibrin glue and suturing of the

abdominal incision. The rats were sacrificed one, four or seven day(s) post surgery. Fluorescence

microscopy revealed that CellTrackerTM-labbeled microtissues had penetrated the myocard by

day 4 (Figure 5). Confocal microscopy analysis of sarcomeric -actinin-specific fluorescent

staining of frozen heart explant sections revealed that myocardial microtissues were already

being incorporated into the rat heart 24 h post implantation (Figure 6A, B). Four days after

infection, the myocardial microtissue was completely embedded in the rat heart, but its spherical

shape was retained (Figure 6C, D). However, by day 7, the transplanted myocardial microtissues

had completed seamless resorption in the absence of structural abnormalities and displayed a

longitudinal cell shape, characteristic of the neighboring host cardiomyocytes. (Figure 6E, F).


Figure 5. Microscopic analysis of myocardial explant sections of rats which had been given a pericardial

injection of CellTrackerTM-labeled myocardial microtissues (assembled from 2,500 NRCs for four days in hanging

drops) four days post transplantation (A-C). Control groups were given buffer injections (D-F). Phase contrast (A),

fluorescent (B), and overlay (C) micrographs show the implanted microtissue (arrowhead) inside the rat

myocardium. (D, E, F) show the respective sections of control animals.


Figure 6. Confocal microscopy analysis of CellTrackerTM-labeled myocardial microtissues, assembled from

2,500 NRCs after cultivation for four days in hanging drops, implanted into the pericardium of adult rats. Treated

rats were sacrificed and the microtissue-myocard interface analyzed on days 1 (A, B), 4 (C, D) and 7 (E, F) by

sarcomeric -actinin- (red; A-F) and CellTrackerTM-specific (B, D, F) (immuno-) fluorescence microscopy.

Microtissues had integrated into the myocardial wall 24 h after surgery (A, B). Four days after transplantation, the

microtissues had completely integrated in the rat’s myocardium, while the microtissues’ spherical shaped was

retained (C, D). Seven days post surgery, implanted and host cardiomyocytes co-aligned, covering all the implant

tracks (E, F).

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Discussion

Because of the limited regeneration capacity of the adult heart damage to the

muscle usually results in irreversible cardiac dysfunction. The design of myocardial regeneration

strategies is thus important to the patient because of the limitations of standard replacement

therapies. Recent reports suggested that endogenous cardiac stem cells may proliferate in the

myocardium under certain conditions and that stem cells infiltrate from the bone marrow to the

heart where they are expected to contribute to repair (Messina et al. 2004; Schuster et al. 2004).

However, discussions are ongoing as to whether bone marrow-derived cells, which penetrate

target tissues, are reprogrammed by the native tissue environment or fuse with target cells

requiring repair (Nygren et al. 2004; Pomerantz and Blau 2004). In the past decade, cardiac cell

transplantation has emerged as the leading strategy for replacing damaged or diseased

myocardium (Melo et al. 2004). Several transplantation initiatives have shown that direct

injections of cell suspensions of fetal neonatal cardiac myocytes into experimental myocardial

infarcts improved the remodeling and function of the heart (Reffelmann et al. 2003). Others have

replicated those encouraging results by using skeletal myoblasts (Menasche et al. 2001), bone

marrow-derived cells (Orlic et al. 2002) or embryonic stem cells (Gepstein 2002; Kehat et al.

2004). Although post transplantation survival was rather limited, only cardiomyocytes enabled

seamless electrogenic cell-host tissue contacts devoid of arrhythmias (Rubart et al. 2003).

To solve the problems associated with single-cell injections, tissue engineers advanced

scaffold design to (i) provide an extracellular matrix substitute for the infarct area, (ii) supply a

temporary support for self or implanted cells and (iii) control the size, shape, strength and

composition of the graft in vitro (Leor and Cohen 2004; Olson 2004). There are a variety of

techniques for constructing beating cardiac patches for transplantation: (i) Shimizu and

colleagues grew rat cardiomyocyte monolayers on polymer surfaces, which were then detached,

stacked and allowed to fuse before subcutaneous implantation into rats (Shimizu et al. 2002). Six

months post transplantation, the cardiac patch was beating and had been infiltrated by host blood

vessels. (ii) Eschenhagen and coworkers produced rings of engineered cardiac muscle by seeding

neonatal rat cardiomyocytes either into a scaffold or by mixing them with collagen and shaping

the tissue in a ring template (Zimmermann et al. 2002; Zandonella 2003; Zimmermann and

Eschenhagen 2003). Instead of seeding cells into scaffolds, cardiac tissues could also be


assembled by organ printing (Jakab et al. 2004). This prototype strategy uses a cell printer which

sequentially plots cells onto a thin thermo-reversible gel support following a software blueprint

of the desired organ shape.

No matter how sophisticated and functionalized scaffolds become, they will remain alien

to the native tissue, with the risk for complications. With the advent of myocardial microtissues,

produced by gravity-enforced self-assembly of monodispersed cardiomyocytes in hanging drops,

the question as to whether scaffolds may become dispensable must be posed again (Kelm et al.

2004). Microtissue production is straightforward and highly flexible, enabling the design of pure

or multi-cell type spheroids as well as cell layers coated onto a core feeder spheroid using a

variety of different cell types including embryonic stem cells, chondrocytes, hepatocytes, retinal

cells, tumor cells and cardiomyocytes (Itskovitz-Eldor et al. 2000; Anderer and Libera 2002;

Layer et al. 2002; Kelm and Fussenegger 2004; Timmins et al. 2004). Thus far, microtissues have

provided a welcome system to for studying tissue assembly, inter-cellular crosstalk and drug

function in a tissue-like in vitro context (Kelm et al. 2005). However, little is known about the

potential of microtissues for regenerative medicine. Recent advances in myocardial microtissue

design have (i) established a linear cell number-microtissue size correlation, (ii) suggested inter-

microtissue superstructures, (iii) shown key cardiomyocyte-specific cell qualities and have

resulted in the (iv) development of an extracellular matrix, (v) the coordinated contraction of the

microtissue’s cell quorum and (v) the size-dependent production of VEGF (Kelm et al., 2004).

Using a novel CMOS-based microchip design we assessed the electrogenic characteristics

of myocardial microtissues (Heer et al. 2004). Electric signal readout from NRC-derived

microtissues revealed that the contraction of cardiomyocytes was coordinated, suggesting

functional electrochemical/mechanical coupling by intercalated disks prominent in these tissue

structures. Refinement of the interface between the biological systems and the microchip is

expected to (i) enable quality control of cardiac tissue implants, (ii) allow critical evaluation of

tissue explants of patients suffering from cardiac-related pathologies and optimize related

diagnosis and (iii) provide a platform for sophisticated drug discovery and/or drug-function

analysis.

The future success of cardiac tissue engineering in regenerative medicine will depend on

the design of larger tissues but with increased tissue size control of vascularization to prevent


ischemia will become more important. Capitalizing on native inter-microtissue interactions we

assembled individual microtissues to functional macrotissues. Control of shape, which is often

thought to be an exclusive asset of scaffolds, was achieved by casting individual microtissues

into custom-designed agarose moulds. Micro- as well as larger cardiac tissues were vascularized

in-vitro by co-assembly of HUVEC and cardiac myocytes, which resulted in a time-dependent

development of a robust capillary network across the myocardical tissues ready to be connected

to native tissues. Indeed, 60 hours after transplantation of myocardial microtissues into chicken

embryos, vascular crosstalk, resulting in the functional connection of the graft tissue to the

embryo, had been established. Furthermore, artificial heart tissues, implanted into the

pericardium of adult rats, integrated seamlessly into the myocardium, confirming in-vivo

compatibility of artificial heart patches.

With several sophisticated scaffold-based/free technologies available and

prototypic organ printing on the rise, electrogenic coupling, under the control and management of

vascularization on track, cardiac tissue engineers are fortunate to have a complementary

technology portfolio at their disposal enabling them to design clinical therapies in the forseeable

future. By combining the aforementioned technologies with high-throughput-compatible tissue

assembly systems and the newest bioprocess engineering tailored for the mass-production of

desired cell phenotypes, tissue engineers may succeed in providing next-generation strategies for

regenerative medicine.

Acknowledgments

We thank Evelyne Perriard for providing neonatal rat cardiomyocytes and Semjidmaa

Dashnyam for isolating of adult rat cardiomyocytes as well as Krystyna Sala-Szymanska for

preparing of the histological samples. This work was supported by the Swiss National Science

Foundation (grant no. 631-065946) and the Swiss State Secretariat for Education and Research

within EC Framework 6.

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Chapter 7

Design of Custom-Shaped Vascularized Tissues

Using Microtissue Spheroids as

Minimal Units

Kelm J.M., Djonov V., Ittner L.M., Born W., Hoerstrup S.P. and Fussenegger M. (submitted)

Macrotissue Generation 125

Abstract

Tissue engineering strategies are gathering clinical momentum in regenerative medicine

and are expected to provide excellent opportunities for the therapy of difficult-to-treat human

pathologies. Being aware of the requirement to produce lager-sized artificial tissue implants for

clinical applications, we used microtissues, produced by gravity-enforced self-assembly of

monodispersed primary cells, as minimal tissue units to generate scaffold-free vascularized

artificial macrotissues in custom-shaped agarose moulds. Mouse myoblast, pig and human

articular chondrocyte (PAC, HAC) as well as human myofibroblast- (HMF) derived microtissues

(µm3 scale) were all amalgamated to coherent macrotissues (mm

3 scale) of the desired shape and

to native tissue ultra-structures. Macrotissues, assembled from the human umbilical vein

endothelial cell (HUVEC)-coated HMF microtissues, developed a HUVEC-managed vascular

system, which functionally connected to the chicken embryo’s vasculature following

implantation. The design of scaffold-free vascularized macrotissues is a first step towards the

scale-up and production of artificial tissue implants for future tissue engineering initiatives.

Introduction

Tissue engineering integrates engineering principles and biological systems in order to

create therapeutic replacement structures (Vacanti and Langer 1999). Tissue development and

maturation is a process of high biological complexity and requires precise orchestration of cell

fate control, metabolic activities, structure-function correlations and crosstalk between and

among different cell populations in space and time. Mastering the custom-shaped assembly of

multiple cell types in an innervated and neovascularized three-dimensional (3D) format remains a

major challenge for current tissue engineering initiatives (Peirce and Skalak 2003; Mol et al.

2004). Pioneering strategies of tissue engineering were based on seeding cells into biodegradable

polymer scaffolds, culturing them in bioreactors and implanting the resulting tissue into the

recipient organism, where maturation of the new organ takes place. Scaffolds were always

thought to be important for tissue engineering as they provide biological, physical and chemical

cues to guide cellular differentiation and assembly in the third dimension (Hubbell 2003; Shin et


al. 2003). Despite advances in biomaterial science, which led to scaffolds with fewer

transplantation side effects resulting from toxic degradation products, the induction of

inflammatory reactions and poor resorption (Yang et al. 2001), most scaffolds fail to promote

vascularization unless they have been specifically functionalized (Vogel and Baneyx 2003).

Since the discovery that monodispersed cells aggregate to spheroids by gravity-enforced self-

assembly in hanging drops and the observation that multicellular spheroids self-organize into

functional tissue-like units according to the differential adhesion hypothesis, scaffold-free

microtissues have been considered as an alternative to artificial scaffold-containing tissues

(Steinberg and Foty 1997; Layer et al. 2002; Kelm and Fussenegger 2004). Self-assembly of

monodispersed cells to microtissues has been successful with a variety of different cell types

including (i) reconstruction of retinal layers (Rothermel and Layer 2001), (ii) design of functional

myocardial microtissues (Kelm et al. 2004), (iii) production of artificial ganglia (Kelm et al.

2005), (iv) construction of cartilage (Anderer and Libera 2002), and tumor cell lines (Kunz-

Schughart et al. 2004). Microtissues assembled in hanging drops attain diameters of only a few

hundred micrometers (100-500 µm in diameter). Beyond this size, diffusion-based nutrient and

oxygen supplies become limiting. In living tissues, cells rely on a capillary network within a

perimeter of 100-150 micrometers for oxygen and nutrient supplies (Tsai et al. 2003).

As well as the innervation and neovascularization of artificial tissues, interconnection to

respective host systems after implantation remain major challenges of current tissue engineering

initiatives (Lauffenburger and Griffith 2001). Neovascularization may be supported by (i)

functionalizing scaffolds with pro-angiogenic factors (Richardson et al. 2001; Zisch et al. 2003;

Zisch et al. 2003; Hall and Hubbell 2004), (ii) ectopic expression of pro-angiogenic factors

(Ajioka et al. 2001) or (iii) by taking advantage of cell-based in-vitro vascularization (Timmins et

al. 2004; Kelm et al. 2005).

For microtissue technology to make an impact on regenerative medicine, microtissues

must be scaled up to larger fully vascularized macrotissues. We have therefore determined

whether microtissues meet those requirements, by assembling microtissues to custom-shaped

neovascularized macrotissues and implanted them into chicken embryos. The observation that

artificial tissues, produced by microtissue scale-up and integrated into host tissues, were

functionally connected to the host vascular system post implantation suggests future

opportunities for applying microtissue-based tissue engineering to regenerative medicine.



Preparation of primary cells

The preparation of primary human myofibroblasts (HMFs) included mincing arterial

mammary segments followed by cultivation at 37°C in a humidified 5% CO2-containing

atmosphere in Dulbecco's modified Eagle's medium (DMEM) (Invitrogen, Carlsbad, CA)

supplemented with 10% fetal bovine serum (FBS; cat. no. 3302-P231902, lot no. P231902; PAN

Biotech GmbH, Aidenbach) and 1% penicillin/streptomycin solution (Invitrogen). Pure HMFs,

which had migrated from the tissue pieces after 10 to 14 days, were serially passaged and

expanded to the desired cell numbers in advanced DMEM (Invitrogen) supplemented with 10%

FBS and GlutaMax (Invitrogen) for four to six weeks under the above aforementioned

conditions.

Cell Culture

Pig (PAC) and human articular chondrocytes (HAC) (kindly provided by Millenium

Biologix, Schlieren, Switzerland) were expanded in DMEM/F12 (Invitrogen) supplemented with

10% FBS, mouse myoblasts C2C12 (ATCC CRL-1772) in DMEM (Invitrogen) supplemented

with 20% FBS (PAN Biotech GmbH, Aidenbach) and human umbilical vein endothelial cells

(HUVECs, PromoCell, Heidelberg, Germany) in endothelial cell growth medium (PromoCell,

cat. no. C-22010) supplemented with 10% FBS. All cell types were cultivated at 37°C in a

humidified 5% CO2-containing atmosphere.

Microtissue Production

After isolation and expansion, the cells were seeded at the indicated cell concentrations

and cell-type compositions into 60-well plates (HLA plate, Greiner-Bio One, Frickenhausen,

Germany). In order to enable gravity-enforced self-assembly of the microtissues in hanging

drops, the 60-well plates were incubated upside down at 37°C in a humidified atmosphere

containing 5% CO2. The specific cultivation conditions for microtissue production are outlined in

Table 1.


Macrotissue Assembly

In order to assemble microtissues to larger artificial tissues we designed two different

agarose casting moulds, (i) a cylindrical mould, 3 mm in diameter, and (ii) a ring-shaped mould,

1 mm thick (Figure 1A). Negative Teflon®

casting moulds were filled with 4% agarose (Sigma

Chemicals, Buchs, Switzerland) in phosphate-buffered saline (PBS; 150 mM NaCl, 6.5 mM

Na2HPO4 x 2 H2O, 2.7 mM KCL, 1.5 mM KH2PO4, pH 7.4) to generate non-adhesive positive

casting moulds to imprint the desired shape on the macrotissue. Microtissues of the indicated

types and numbers of cells were transferred to the respective agarose moulds and cultivated in

cell type-specific media under static conditions at 37°C in a humidified atmosphere containing

5% CO2 (see Table 1 for specific culture conditions).

Immunohistochemistry

The tissues were fixed in 2% paraformaldehyde, rinsed in 15% sucrose solution and

stored at -20°C in 70% ethanol. All the samples were dehydrated by sequential incubation in

increasing ethanol concentrations and embedded in paraffin wax. 3 µm sections, produced using

an Ultracut device (Zeiss, Feldbach, Switzerland), were transferred to gelatinized micro-slides

and air-dried overnight at 37°C. The samples were de-waxed in xylene (three changes),

rehydrated in ethanol and rinsed in Tris-buffered saline (TBS, 20 mM Tris base, 155 mM NaCl, 2

Table 1. Culture media used for cell expansion, microtissue production and macrotissues assembly

Cell Type Cell Expansion

Medium

Microtissue Assembly

Medium

Microtissue Cell

Number

Microtissue

Assembly Time

Microtissues Per

Macrotissue

Tissue Assembly

Time

C2C12 DMEM

+ 20% FBS

DMEM

+ 10% HS 10’000 4 [d] 1’200 10 [d]

PAC/HAC DMEM/F12

+ 10% FBS

DMEM

+ 10% HuS 5’000 5 [d] 600 14 [d]

HMF Adv. DMEM

+ 10% FBS

DMEM

+ 10% HuS 10’000

4 [d] + 4 [d] (HUVEC coating)

300 7 [d]

HUVEC ECGM

+ 10% FBS

___ 1’200 ___ 1’200 ___

Abbreviations: advanced DMEM, cell culture medium (Invitrogen); C2C12, mouse myoblast cell line; DMEM, Dulbecco`s Modified Eagle Medium

(Invitrogen); DMEM/F12, culture medium (Invitrogen); ECGM, endothelial cell growth medium (PromoCell); HAC, human articular chondrocytes;

HMF, human myofibroblasts; HS, horse serum (cat. no. 16050-098, lot no. 3036354D, Invitrogen); HUVEC, human umbilical vein endothelial cells;FBS, fetal bovine serum (PAN Biotech GmbH); HuS: human serum (cat. No. P30-2501, lot no. P520221, PAN Biotech GmbH); PAC, pig articular

chondrocytes.


mM EGTA, 2 mM MgCl2) (two changes). Endogenous peroxidase activity was eliminated by

treatment with 0.3% hydrogen peroxide for 10 min. The sections were subsequently heated for 15

min in a microwave oven (180 W) and blocked by incubation for 10 min in TBS containing 1%

casein (Sigma Chemicals). All sections were then incubated for 15 h at 4°C in TBS containing

primary antibodies specific for VEGF (rabbit polyclonal, Santa Cruz Biotechnology, Santa Cruz,

CA, USA) and CD31 (mouse monoclonal, Dako, Glostrup, Denmark). Then, the sections were

exposed to an affinity-purified biotinylated secondary antibody ([anti-mouse EO 433, anti-rabbit

EO 353, Dako, Glostrup, Denmark] diluted 1:200 in TBS) for 45 min at room temperature,

washed three times in TBS and then treated with the straptavidin-biotin-complex/horseradish

peroxidase (Dako, Glostrup, Denmark) for another 45 min at room temperature. The reaction

product was visualized by exposing the sections to 3-amino-9-ethylcarbazole or 3.3-

diaminobenzidine (Sigma Chemicals), which were then mounted in Aquatex (Merck, Darmstadt,

Germany). The negative controls were stained with non-specific mouse and rabbit sera.

Transmission electron microscopy

Micro- and macro-tissues were fixed by immersion in 0.1 M cacodylate buffer (pH 7.4,

350 mOsm) containing 2.5% glutaraldehyde. The tissue blocks were postfixed in osmium

tetroxide, block-stained using uranyl acetate, dehydrated by sequential incubation in increasing

ethanol concentrations and embedded in Epon 812 according to Djonov and coworkers (all

chemicals from Merck Eurolab AG, Dietikon, Switzerland) (Djonov et al. 2000). Semi-thin 1 µm

sections were stained with toluidine blue and visualized using an Olympus Vanox BHS light

microscope (Olympus AG, Volketswil, Switzerland). Ultra-thin sections of 80-90 nm were cut

using a diamond knife and picked up on Formvar-coated (polyvinyl formal; Fluka Chemie AG)

copper grids, double-stained with lead citrate (Merck Eurolab AG) and uranyl acetate and

monitored on a Philips EM 400 electron microscope (FEI AG, Zurich, Switzerland).

Chicken chorioallantoic membrane (CAM) assay

Chicken embryos were cultured according to the shell-free method (Ribatti et al. 2001).

Artificial tissues were grafted onto the growing chicken chorioallantoic membrane (CAM) at

embryonic day 10 and cultured for 3.5 days.


Results

Microtissue assembly to larger-sized macrotissues

Although microtissues enable new insight into intercellular crosstalk, specific 3D cell

phenotypes and tissue assembly, they are considered too to be small for generating clinical

impact in tissue engineering. Capitalizing on the tissue-like cell morphologies adopted in

microtissues we evaluated their potential to serve as building blocks for larger artificial tissues.

Therefore, 1,200 microtissues, produced from 10,000 mouse myoblasts (C2C12) aggregated in

hanging drops for four days, were assembled to macrotissues in custom-designed ring-shaped

1 mm thick agarose moulds for three days (Figure 1A). The resulting macrotissues were cut once

to produce a tissue string, which was cultivated for another week in agarose-coated culture

dishes. Macroscopic examination revealed that individual microtissues had amalgamated to a

coherent tissue structure (Figure 1B). Toluidine blue staining of 1 µm sections showed highly

organized tissue morphology reminiscent of muscle fibers composed of individual multinucleated

cells (Figures 1C-E). Although ultra-structural analysis revealed a typical longitudinal shape of

muscle cells, myofibrils did not develop, most likely due absence of mechanical forces and/or

electrogenic stimulation (Figure 1F) (Fink et al. 2000; Benjamin and Hillen 2003).


Figure 1. Production of C2C12-derived macrotissues. (a) Agarose mould generated from a negative

Teflon® form to assemble C2C12-derived microtissues to 1 mm thick ring-shaped muscle macrotissues. The ring-

shaped casting chamber is visualized by a blue dye. (b) C2C12-derived macrotissue assembled from 1,200 C2C12

microtissues, each containing 10,000 myoblasts, for three days and cultivated for another seven days in non-adhesive

agarose-coated plates. (d, e) Toluidine blue-stained semi-thin sections of muscle macrotissues reveals a coherent

tissue structure containing regions with a high content of multi-nucleated cells (d, e, arrows). Ultrastructural analysis

shows that longitudinally oriented muscle cells are interconnected by cell-cell contacts (F, circle).

ba

c d

e f

2 mm 0.5 mm

50 m 10 m

10 m 5 m

bbaa

cc dd

ee ff

2 mm 0.5 mm0.5 mm

50 m50 m 10 m10 m10 m

10 m10 m10 m 5 m5 m


Six hundred cartilage microtissues, assembled from 5,000 pig articular chondrocytes

(PAC) or human articular chondrocytes (HAC) for five days in hanging drops, were shaped into a

cylindrical macrotissue (3 mm in diameter) by cultivation in a specific agarose mould for seven

days in the absence of collagen-inducing factors like TGF- or ascorbic acid. Artificial cartilage

tissues were subsequently cultivated for another week in agarose-coated culture dishes. PAC-

derived microtissues had amalgamated into coherent tissues (Figure 2A) characterized by the

ubiquitous production of collagen (Figures 2B-F). Akin to the PAC-composed tissues, HAC-

derived artificial cartilage showed areas of increased collagen production (Figures 3A-F).

Toluidine blue staining revealed bright islet-like structures surrounded by squamous

chondrocytes with a palisade arrangement (Figures 3A-D), which ultrastructural analysis,

revealed to be a robust fibrillary network (Figures 3E and F). Artificial cartilage chondrocytes

exhibited an active euchromatin nucleus with a prominent nucleolus, a cytoplasm with a high,

rough endoplasmic reticulum content and a plasma membrane characterized by numerous

filopodia extending out into the surrounding extracellular matrix (Figures 3E and F).

Figure 2. Pig articular chondrocyte (PAC) microtissues (600, each containing 5,000 PACs) assembled in a

cylindrical agarose mould (3 mm in diameter) to a coherent artificial cartilage (a). Toluidine blue staining of semi-

thin paraffin sections of PAC-derived macrotissues tissues shows a robust fibrillary network embedded in an

extensive extracellular matrix (b, c, asterisks), and van Gieson staining reveals collagen production in light red (d-f,

asterisks).

c 20 ma

c

b 0.5 mm

d 0.5 mm

e

e 50 m

f

f 20 m

0.5 mm c 20 mcc 20 m20 maa

c

b 0.5 mm

c

bb 0.5 mm0.5 mm

d 0.5 mm

e

dd 0.5 mm0.5 mm0.5 mm

e

e 50 m

f

e 50 m50 m

f

f 20 mff 20 m20 m

0.5 mm0.5 mm


Figure 3. Human articular chondrocyte (HAC) microtissues (600, each containing 5,000 HACs) assembled

in a cylindrical agarose mould (3 mm in diameter) to a coherent artificial cartilage. Toluidine blue staining shows

bright islet-like structures (asterisk) surrounded by squamous chondrocytes in palisade arrangement (a-d).

Transmission electron micrographs (e, f) revealed that the islets are composed of robust fibrillary network embedded

in an amorphous matrix (e, f, asterisks). Artificial cartilage chondrocytes exhibited an active euchromatin nucleus

with a prominent nucleolus, a cytoplasm with a high, rough endoplasmic reticulum content (arrow head) and a

plasma membrane characterized by numerous filopodia (arrows) extending out into the surrounding extracellular

matrix (e, f).

a b

c d

e f

0.5 mm 25 m

10 m 10 m

5 m 1 m

aa b

cc dd

ee ff

0.5 mm0.5 mm 25 m25 m

10 m10 m 10 m10 m

5 m5 m 1 m1 m


Neo-vascularization of scaffold-free macrotissues

The aforementioned experiments have established microtissues as functional building

blocks, which can be assembled into scaffold-free tissue structures following self-controlled

assembly in adhesion-free agarose moulds. Along the way to designing larger artificial tissue

structures for clinical applications, will be the prime size-limiting parameter vascularization,

which sustains metabolic activities of the tissue core. Previous studies exemplified the in-vitro

vascularization of human aortic fibroblast (HAF)-based core microtissues, whose VEGF

production recruited peripheral HUVECs to develop a vascular structure (Kelm et al. 2005). In

order to pioneer in-vitro vascularized macrotissues we produced core human myofibroblast

(HMF) spheroids by cultivating 10,000 cells for four days in hanging drops, coating them with

1,200 HUVECs for four days in hanging drops and assembling 300 HMF-HUVEC spheroids in

the aforementioned cylindrical agarose mould. The HMF-HUVEC macrotissue aggregated for

three days and was cultivated for further 4 days in an agarose-coated culture dish. HMF-HUVEC

microtissues assembled into a coherent macrotissue (Figure 4A), characterized by a peripheral

layer of HUVECs (Figure 4C) reminiscent of inter-microtissue forces, which separated mixed

HMF-HUVEC populations into a HUVEC-coated HMF core spheroid during microtissue

assembly in hanging drops. In contrast, pure HMF macrotissues were less coherent and had a

ruffled surface (Figures 4B and D). Moreover, immunohistologic analysis of the endothelial cell-

specific surface marker CD31 revealed a dense network of endothelial cells throughout the HMF-

HUVEC macrotissue (Figures 4E, G, H), a cell network which was not be observed in pure HMF

control tissues (Figure 4F). The equivalence of cell type-specific reorganization (HMF-core,

HUVEC-shell) within multicellular micro- and macrotissues suggests that microtissues can

indeed be considered to be minimal tissue units, which can be up-scaled to larger tissue implants.


Figure 4. Artificial connective tissue assembled from 300 human myofibroblast (HMF)-composed

microtissues (10’000 HMFs) coated with 1’200 HUVECs (a, c, e, g, h) (HMF-HUVEC) or assembled from pure

10 m

a b

c d

e f

g h

0.3 mm 0.3 mm

10 m

0.3 mm0.3 mm

20 m 20 m

10 m10 m10 m10 m

aa bb

cc dd

ee ff

gg hh

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10 m10 m

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20 m20 m 20 m20 m

10 m10 m


HMF microtissues (b, d, f). While HMF-HUVEC macrotissues were coherent and showed a smooth tissue surface

(a) tissues composed of pure HMF microtissues displayed a ruffled surface (b), which was confirmed by toluidine

blue stained semi-thin sections (d). Sections of HMF-HUVEC showed exclusive assembly of HUVECs at the

macrotissue’s periphery (c; double-head arrow). Immunohistologic analysis of the endothelial cell-specific surface

marker CD31 (brown staining) revealed a HUVEC-shaped capillary network exclusive to HMF-HUVEC

macrotissues (e, g, h) (f, control macrotissue composed of HMFs only).

Implantation of HMF-HUVEC macrotissues into chicken embryos

In-vivo vascularization crosstalk between artificial and living tissues was assessed by

implanting in-vitro prevascularized HMF-HUVEC macrotissues (see above) atop the

chorioallantoic membrane (CAM) of chicken embryos. Neovascularized HMF-HUVEC

macrotissues integrated into the CAM 84 hours post implantation and were fully connected to the

chicken embryos’s vasculature (Figure 5A). At the macrotissue-CAM interface, a dense network

of erythrocyte-containing microvessels was detected, suggesting inter-species compatibility of

vascular systems (Figures 5B and C). Furthermore, ultra-structural analysis revealed that

individual HUVECs had developed intracellular lumen and that inter-HUVEC crosstalk had

established small microvessels with lumen filled with erythrocytes (Figure 5G). Regular

HUVEC-formed capillaries were characterized by an activated endothelium with intra-luminal

protrusions, a multitude of transport vesicles and an extracellular matrix formed by deposition of

collagen fibers (Figure 5H). In contrast, control macrotissues assembled from HUVEC-free HMF

populations were rejected, scar tissue with no signs of vascular crosstalk developed between the

control tissue and the chicken embryo (Figures 5D-F).

Immunohistological analysis of the human endothelial cell-specific surface marker CD31

confirmed the development of a vascular system across the macrotissue-embryo interface

(Figures 6A and B). Interestingly, HUVECs were found to accumulate preferentially at the

macrotissue-CAM interface managing inter-tissue vascularization. Some HUVECs even migrated

to the chicken embryo’s mesenchym (Figures 6C and D). VEGF-specific staining of macrotissue

implants showed high-level expression of this growth factor in non-vascularized implanted

control macrotissues known to suffer from hypoxia in their core (Kelm et al. 2004). However,

pre-vascularized macrotissue implants, fully connected to the embryo’s vasculature, exhibited


low levels of VEGF expression in the macrotissue, thus confirming a sufficient oxygen supply

inside the implant (Figures 6E and F).

Figure 5. Vascularization crosstalk of human myofibroblast (HMF)-derived macrotissues transplanted onto

chicken embryos. Artificial connective tissue assembled from human umbilical vein endothelial cell (HUVEC)-

Er

Er

a d

b e

c f

g h

1 mm 1 mm

100 m 100 m

50 m 50 m

5 m 5 m

b

c

e

Er

Er

aa dd

bb ee

cc ff

gg hh

1 mm1 mm 1 mm1 mm

100 m100 m 100 m100 m

50 m50 m 50 m50 m

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c

e


coated HMF (HMF-HUVEC) (a-c, g, h) or pure HMF microtissues (d-f) were grafted and cultivated atop the chicken

embryo’s chorioallantoic membrane (CAM) for 3.5 days. Toluidine blue staining of semi-thin sections across the

HMF(-HUVEC)/CAM interface revealed a significant angiogenic response of the chicken embryo exemplified by

large vessels invading the human tissue implant (b, arrows) as well as multitude of capillaries within the artificial

connective tissue (c, arrows). In contrast, pure HMF control macrotissues are not connected to the chicken embryo’s

mesenchyme and vasculature and remain non-vascularized. The graft is rejected as the chicken embryo develops an

enlarged CAM epithelium at the CAM-HMF tissue interface (e, f, asterisks). Transmission electron micrographs

show that (i) microvessels are filled with erythrocytes (Er) (g, h), (ii) sprouts of seamless non-perfused capillaries

formed by single (g, arrow head) or (iii) by adjacent endothelial cells (g, arrow). Most capillaries are activated, and

show a thick endothelium characterized by intra-luminal protrusions of trans-cellular vesicles (h, arrowhead). The

extracellular matrix shows deposition of collagen (h, arrows).

Figure 6. Immunohistologic analysis of human umbilical vein endothelial cell (HUVEC)-coated human

myofibroblast (HMF) (a, c, d, e) and pure HMF (b, f) microtissues assembled to macrotissues and grafted onto

chicken embryos’ chorioallantoic membrane (CAM) for 3.5 days. HUVEC-formed tubular capillary structures,

d

+ HUVEC - HUVEC

+ HUVEC - HUVEC

CD31 CD31

VEGFVEGF

ca

c

e

b

d

f

1 mm 1 mm

100 m 50 m

1 mm1 mm

d

+ HUVEC - HUVEC

+ HUVEC - HUVEC

CD31 CD31

VEGFVEGF

caa

cc

ee

bb

dd

ff

1 mm1 mm 1 mm1 mm

100 m100 m 50 m50 m

1 mm1 mm1 mm1 mm


staining positive for CD31, connected to CAM vessels and invaded the chicken embryo’s mesenchyme (c, d,

arrows). In contrast, HMF-only macrotissues show no vascular connection and remain separated from the host tissue

by a thick epithelium (see also Figure 5). Vascularization of the HMF-HUVEC tissues result in lower production of

the human vascular endothelial growth factor (VEGF) (e), compared to the HMF-pure control tissue (f).

Discussion

So far, classical tissue engineering has been based on seeding expanded cell populations

into synthetic scaffolds or hydrogels to provide a desired shape and implanting the resulting

artificial tissues into the recipient organism, where maturation of the new organ or integration

into host tissues occurred (Hoerstrup et al. 2000; Hench and Polak 2002; Shin et al. 2003; Langer

and Tirrell 2004). Despite decisive advances in the design and functionalization of the scaffold

during the past decade two major challenges remain: (i) Scaffolds fail to mimic essential natural

extracellular matrix (ECM) functions with temporal and spatial intricacy, and (ii) cell-scaffold

interactions lack the signal integration complexity observed for inter-cellular crosstalk in three

dimensions (Richardson et al. 2001; Vogel and Baneyx 2003; Lutolf and Hubbell 2005).

Alternative strategies for the design of scaffold-free tissues took advantage of stacking cell

monolayers (Shimizu et al. 2002) or capitalized on the self-assembly of monodispersed cells to

microtissues (Steinberg and Foty 1997; Kelm and Fussenegger 2004). Unencumbered by scaffold

materials, cells that assemble into microtissues develop their natural microenvironments, control

tissue dynamics and adopt in vivo-like structures. Non-limiting examples include self-assembly

of myocardial microtissues, Kelm et al. 2004), microganglia-like structures (Kelm et al. 2005),

hepatic microtissues (Kelm et al. 2003), artificial cartilage (Anderer and Libera 2002), embryoid

bodies (Itskovitz-Eldor et al. 2000), and retinal structures (Layer et al. 2002). Microtissues have

since been appreciated as 3D tissue culture models and have been used successfully in this study

to assemble larger tissues in vitro.

Mammalian cell aggregates have been shown to display a high cell type-specific affinity,

which may result in the assembly of larger tissues. For example, transgenic Chinese hamster

ovary cells, embedded in hydrogel droplets, fused to ring-like structures (Jakab et al. 2004).

Using cell-specific microtissues to assemble (i) skeletal muscle (C2C12, derived from mouse

myoblasts), (ii) cartilage (derived from pig and human articular chondrocytes) and (iii)

connective (derived from human myofibroblasts) macrotissues, we have substantiated the use of


microtissues as minimal building blocks to generate larger scaffold-free artificial tissue at a mm3

scale. Macroscopic and ultra-structural analyses revealed that all macrotissues displayed tissue-

specific morphologies based on. C2C12 displayed skeletal muscle-specific, longitudinal,

multinucleated cell morphology, and chondrocytes were embedded in a collagen matrix even

without supplements of collagen-inducing substances like ascorbic acid or TGF- .

Increasing the size of artificial tissue beyond a certain threshold diameter will elicit

nutrient and oxygen limitations in the tissue’s core. Nature has evolved vascularization to ensure

metabolic supplies to individual cells in a tissue within a perimeter of 100 µm (Tsai et al. 2003)

Tissue engineers have elaborated two fundamental strategies to mimic vascularization in artificial

tissues: (i) using cells transgenic for the production of pro-angiogenic factors or scaffolds

functionalized with pro-angiogenic factors to recruit tissue-resident endothelial cells for implant

vascularization (Richardson et al. 2001; Hall and Hubbell 2004), (ii) in-vitro pre-vascularization

of implants using purified pro-angiogenic factors or specifically designed scaffold compositions

to enable rapid connection to the host vascular system (Borges et al. 2003; Wu, et al. 2004). The

use and release of pro-angiogenic factors remains problematic due to the inherent instability of

these proteins in vivo (Yancopoulos et al. 2000) and the risk of uncontrolled side effects

including angiomagenesis (Carmeliet 2000). In-vitro implant pre-vascularization using

endothelial cells was difficult as these cells became apoptotic after being embedded in gelatin,

fibrin, collagen or matrigel matrices (Nomi et al. 2002)

We found that coating the desired tissue spheroids with HUVECs results in

perfectly vascularized microtissues as HUVECs migrate to the microtissue’s core following the

established VEGF gradient. Unlike in matrices, HUVECs remain viable in a microtissue

environment (Kelm et al. 2005). As minimal tissue building blocks, HUVEC-vascularized

myofibroblast microtissues perfectly assembled to vascularized macrotissues, which connected to

chicken embryo’s vasculature following implantation. However, non-vascularized HUVEC-free

macrotissue implants were rejected. This observation substantiates the need for the pre-

vascularization of tissue implants beyond a certain size.

An ideal artificial tissue transplant should show three major characteristics: (i) adopt

tissue-typical cell morphologies and tissue integrity, (ii) exhibit seamless and functional

integration into the target tissue without production of significant scar tissue and (iii) connect to


the host vascular and nervous systems. Microtissue-based production of prevascularized tissues

meets all these criteria. With the tissue engineering details in place, bioengineers will have to

take over in order to design mass-production strategies for microtissues and provide clinics with

transplantation-ready artificial tissues.

Acknowledgments

We thank Sirpa Price for isolating of primary human myofibroblasts and Krystyna Sala-

Szymanska as well as Bettina de Breuyn for preparing of the histological specimens. This work

was supported by the Swiss National Science Foundation (grant no. 631-065946) and the Swiss

State Secretariat for Education and Research within EC Framework 6.

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Chapter 8

Synergies of Microtissue Design, Viral

Transduction And Adjustable Transgene

Expression For Regenerative Medicine

Kelm J.M., Kramer B.P., Gonzalez-Nicolini V., Ley B. and Fussenegger M., (2004)

Biotechnology and Applied Biochemistry 39, 3-16

Microtissue

Design

Transduction

Gene Control

Gene Therapy

Gene-Function

Analysis

Drug

Discovery

Tissue engineering

Animal-Free

Drug Testing

Cell-Phenotype Engineering

Biopharmaceutical

Manufacturing

Synergies of Microtissue Design, Transduction and Regulated Gene Expression 146

Abstract

In the past decade, regenerative medicine has evolved as an interdisciplinary field

integrating expertise from the medical, life and material science communities. Recent

advances in tissue engineering, gene therapy, gene-function analysis, animal-free drug testing,

drug discovery, biopharmaceutical manufacturing and cell-phenotype engineering have

capitalized on a core technology portfolio including artificial microtissue design, viral

transduction and precise transcription dosing of therapeutic or phenotype-modulating

transgenes. We provide a detailed overview on recent progress in these core technologies and

comment on their synergistic impact on current and future human therapies.

Introduction

Tissue engineering and gene therapy may be defined as rational therapeutic

interventions to restore diseased gene or tissue functions. In the past two decades advances in

gene therapy and tissue engineering have resulted in a variety of therapeutic advances to

generate and/or reinstate tissue function of skin (Beele 2002), cartilage (Hardingham et al.

2002), bone (Jadlowiec et al. 2003), intestine (Perez et al. 2002), myocardium (Eschenhagen

et al. 2002), pancreas (Prokop 2001) or liver (Samuel 2003). As tissue engineering and gene

therapy are becoming increasingly mature fields they start to merge on their way to foster

novel opportunities in regenerative medicine. Next-generation advances in regenerative

medicine require a specific technology portfolio including (1) systems for cultivation of

(primary) cells in three dimensions (3D) to form artificial (micro-) tissues, (2) efficient

transduction technologies to place preferred transgene configurations onto any target

chromosome (Pfeifer et al. 2002) and (3) human-compatible gene regulation technology for

adjustable molecular interventions as well as rational reprogramming of mammalian cells to

achieve desired cell phenotypes (Carmeliet 2000; Lee et al. 2000). Besides for regenerative

medicine (tissue engineering and gene therapy) these three core technologies also form the

integral basis for other therapeutic areas including (i) gene-function analysis (Greenfield

2000), (ii) drug discovery (Aubel et al. 2001), (iii) animal-free drug testing (Bhadriraju et al.

2002) and biopharmaceutical manufacturing (Fussenegger et al. 1998; Meents et al. 2002).


Figure 1: Integration of three core technologies is the basis for advanced regenerative medicine (tissue

engineering and gene therapy): (i) Microtissue design, (ii) transduction and (iii) gene control. These three key

technologies merge to provide impact on (i) animal-free drug testing, (ii) biopharmaceutical manufacturing, (iii)

cell phenotype engineering (targeted differentiation), (iv) drug discovery, and (v) gene-function analysis.


It has been shown for a wide variety of different cell types such as bone (Kale et al.

2000), liver (Khalil et al. 2001), cartilage (Girotto et al. 2003), and heart muscle cells

(Eschenhagen et al. 1997) that 3D cultivation is an absolute requirement to maintain cell-

specific functionality in vitro (Berthiaume et al. 1996), (Chen et al. 1997). In many cases,

cells cultivated in 2D and 3D differ in their expression profiles for 5 1 and 5 3 integrins,

paxillin and other cytoskeletal components, their phosphorylation status of focal adhesion

kinases and an extracellular matrix (ECM) is exclusively produced by 3D cultures

(Cukierman et al. 2001; Kelm et al. in press).

Beyond providing a physical support for cells the ECM modulates cell differentiation

(Edwards et al. 1998) as well as tissue remodelling (Badylak 2002) and manages inter-tissue

contacts (Giancotti et al. 1999). Also, cell surface receptors interacting with the ECM

integrate growth-modulating external signals and convey them to the cells’ signal

Microtissue

Design

Transduction

Gene Control

Gene Therapy

Gene-Function

Analysis

Drug

Discovery

Tissue engineering

Animal-Free

Drug Testing

Cell-Phenotype Engineering

Biopharmaceutical

Manufacturing

Microtissue

Design

Transduction

Gene Control

Gene TherapyGene Therapy

Gene-Function

Analysis

Gene-Function

Analysis

Drug

Discovery

Drug

Discovery

Tissue engineeringTissue engineering

Animal-Free

Drug Testing

Animal-Free

Drug Testing

Cell-Phenotype EngineeringCell-Phenotype Engineering

Biopharmaceutical

Manufacturing


transduction cascades (Streuli 1999). For example, neonatal mouse and rat cardiomyocytes

cultivated in 2D lack collagen IV whereas this ECM component is highly expressed in

myocardial microtissues (Kelm et al. in press).

The importance of the ECM, cell morphology and differentiation status as well as cell-

cell contacts for the integrity of any tissue has been well established (Martin et al. 2002).

However, recent advances in polymer science were required to design artificial support for 3D

cultivation of desired cell types. To date, two different strategies for 3D cultivation of

mammalian cells are available: Cultivation in/on artificial scaffolds and reaggregation.

Although collagens (Eschenhagen et al. 1997), foams (Fukuda et al. 2003), and hydrogels

(Khalil et al. 2001) have been shown to enable 3D cell culture, novel biocompatible and

biodegradable polymer scaffolds in particular those functionalized with wound-healing and

growth-promoting factors have become a standard in the production of artificial tissue

constructs (Richardson et al. 2001). Despite their decisive advantages for reshaping 3D

structures, scaffolds have been associated with undesired post-translational effects,

inflammation originating from toxic degradation products, poor resorption and high

production costs (Yang et al. 2001).

The reaggregation approach represents an alternative to the use of artificial scaffolds

for 3D cultivation (Layer et al. 2002). Reaggregation is based on the principle that cells in

suspension self-aggregate under appropriate culture conditions to form spheroids. The use of

spheroids has a long tradition in anticancer research since neoplastic cells arranged in a

multicellular 3D configuration show increased resistance to cytotoxic drugs (multicellular

resistance; MCR) compared to isogenic cells grown as monolayer which enables a more

precise assessment of in vivo drug performance (Desoize et al. 2000). Various cell types

have been cultivated as cellular reaggregates including undifferentiated embryonic bodies

(Itskovitz-Eldor et al. 2000) and neurospheres (Nunes et al. 2003) or terminally differentiated

cardiomyocytes (Kelm et al. in press) and hepatocytes (Khaoustov et al. 1999). Spheroids

produced from undifferentiated progenitor or stem cells display increased proliferation in 3D

compared to 2D monolayer cultures (Chen et al. 1997). By contrast, many cell aggregates

assembled from terminally differentiated cells typically proliferate as monolayers but fail to

do so in a 3D configuration in which they exclusively produce an extracellular matrix (see

above).

Classical cultivation systems for induction of reaggregation include non-adhesive

culture dishes, spinner flasks and roller bottles. Unfortunately, these culture technologies do

not enable control of spheroid size. Only the hanging drop technology which consists of


cultivating cell suspensions of defined cell number in specific multiwell plates incubated

upside down, provide efficient size management. Gravity-enforced reaggregation of the cell

suspension in the hanging drop produces spheroids of specific size which correlates with the

cell number of the inoculated suspension. Size control was recently established for

cardiomyocytes (Kelm et al. in press) and several neoplastic cell lines including hepatocytes

(Kelm et al. 2003). Control of microtissue size is key to avoid oxygen and nutrient limitations

in the spheroid center. For example, we have recently observed hypoxia-induced expression

of vascular endothelial growth factor (VEGF) in rat myocardial microtissues of 230 µm in

diameter whereas smaller-sized microtissues of 120 µm failed to secrete VEGF (Kelm et al. in

press). Management of size-dependent hypoxia-controlled VEGF expression in artificial

microtissues may prevent the risk for tumor formation resulting from sustained angiogenesis

(Lee et al. 2000).

Directed Cell Differentiation

Cell-based therapies require substantial supplies of specific cell phenotypes. Although

autologous cell material is preferred, the availability of desired cell phenotypes is often

limited. Therefore, different strategies have been designed to produce sufficient supplies of

desired cell phenotypes: (i) precise reprogramming of cell-cycle regulatory networks to

enable cell-cycle reentry of terminally differentiated cells for expansion, followed by

induction of a G1-specific growth arrest post implantation to prevent development of

neoplastic cell phenotypes (Blau et al. 1997; Fux et al. 2001), (ii) expansion of stem cells and

progenitor cells by addition of purified growth factors (Schuldiner et al. 2000), cocultivation

of different cell types (Condorelli et al. 2001) or transduction of growth and transcription

factor-encoding genes (Horb et al. 2003), (iii) rational reprogramming of desired mammalian

cells by conditional overexpression of specific differentiation and dedifferentiation factors

(Fux in press; Fux in press).

Future success of cell and tissue engineering for ex vivo expansion will be

based on technologies managing temporal proliferation control of mammalian cells through

well-balanced expression of growth-promoting and growth-suppressing genetic elements

(Mulligan 1993). Independent control of growth-promoting and -inhibiting genes requires

combination of two compatible regulation systems, such as the PipOFF and TetOFF systems

(see below). A recent example includes dual-regulated expression of p27Kip1 sense and

antisense (Fux et al. 2001).


Directed differentiation of human embryonic stem cells for tissue engineering is still

in its infancy but holds great promises for clinical applications, in particular for proliferation-

inert cell phenotypes including cardiomyocytes. To date, directed differentiation of a single

cell phenotype is still a scientific dream. Schuldiner and coworkers scored the differentiation

impact of eight different growth factors which resulted in cell populations displaying different

cell phenotypes and exemplified that most growth factors induce various differentiation

programs (Schuldiner et al. 2000). An alternative strategy to obtain single-phenotype

populations from differentiated stem cells is by sorting, although the cell material produced

may fail to reach clinical quantities at reasonable cost (Kolossov et al. 1998).

Understanding interactions of key (trans-) differentiation factors requires sophisticated

in vitro model systems. A well-investigated setting is the transdifferentiation of pancreatic

cells to hepatocytes based on the pancreatic cell line AR42J-B13 (Shen et al. 2000).

Furthermore, Fux and coworkers implemented dual-regulated expression of (i) MyoD and

Msx-1 and (ii) C/EBP- and BMP-2 for precise differentiation control of C2C12 cells into

osteoblasts, adipocytes and myotubes (Fux in press; Fux in press).

Gene-function analysis

A major determinant of cellular function and phenotype is the tissue transcriptome.

Transcriptome analysis enables elucidation of human diseases (Golub et al. 1999), reveals

targets of drug candidates and helps to determine cDNAs for therapeutic interventions

(Greenfield 2000). Also, global transcriptome changes resulting from cultivation of cells in

3D instead of 2D may provide critical information required for regenerative medicine. We

have recently produced hepatic microtissues by cultivating the human hepatocarcinoma cell

line HepG2 in hanging drops. HepG2-derived microtissues display several of the phenotypic

characteristics of normal liver cells when cultivated in 3D (Khalil et al. 2001; Kelm et al.

2003). The transcriptome of HepG2 grown in monolayers and in 3D was analyzed using a

macroarray containing 3528 human genes (Atlas™ Human 3.6 Array; Clontech, Palo Alto,

CA). Several liver-specific, metabolism-associated and detoxifying proteins were upregulated

in microtissues probably as a consequence of induction of HNF-3 , a transcription factor

modulating a variety of liver-specific target genes (Figure 2). Hepatocyte-specific gene

expression is known to be controlled by several transcription factors including HNF-1,

CCAAT/enhancer binding protein (C/EBP), HNF-3, HNF-4 and HNF-6 (Duncan et al. 1998).


Based on our transcriptome studies HNF-3 may be a key target to reprogram liver-specific

cell phenotypes.

Figure 2: Expression analysis of the human hepatocarcinoma cell line HepG2 cultivated as 3D

microtissues compared to 2D monolayer cultures. In order to cover the functionality of liver cells, expression

profiles of metabolic and detoxifying genes were analyzed using a macroarray containing 3528 genes (Atlas™

Human 3.6 Array; Clontech, Palo Alto, CA). Most of the liver-specific genes were exclusively upregulated in

micortissues suggesting that 3D cultivated is privileged in providing liver-specific functions.

Animal-free drug testing and drug discovery

Screening for novel drugs and design of new drug tests require reproducible cell-based

models which mimic in vivo-like tissue structures. The fact that many established cell-based


systems designed for drug testing show little correlation between in vitro and in vivo results

exemplifies the need for novel culture systems assessing drug action in a tissue-like

environment (Bhadriraju et al. 2002). 3D multicellular tumor spheroids have long been

suggested as an alternative to animal tests for scoring efficacy of drugs against solid tumors.

Owing to decisive advances in 3D cell culture technology in recent years, protocols for 3D

cultivation of a wide variety of primary cells and cell lines are now available and complete the

cell-based drug screening portfolio (Padron et al. 2000; Braeckmans et al. 2002). For

example, progress in reshaping fully functional myocardial microtissues beating over 3 weeks

may provide a high throughput-compatible assay platform for the detection of myocardium-

stimulating or beating frequency-modulating drugs (Kelm et al. in press). We believe that

synergies resulting from combined use of molecular therapeutic interventions and tissue-

mimicking 3D cultivation systems will foster novel opportunities in regenerative medicine.

Viral Transduction

The ability to manipulate the genetic constitution of any living system has come along

with various technologies. The strategies being pursued in this context cover a wide variety of

different areas from biopharmaceutical manufacturing and gene-function analysis to screening

systems and clinical applications. Clinical applications include some aspects of tissue

engineering, tissue and organ transplantation as well as gene therapy which has already

generated promising impact in clinical trials (Grossman et al. 1994; Blaese et al. 1995;

Cavazzana-Calvo et al. 2000). The science of gene transduction is based on administration of

genetic information in order to equip target cells with new protein-production capacities or

eliminate expression of disease-inducing determinants (see Amor et al. for a review (Amor

2001)). Whereas transduction of functional genes is used to complement single-gene defects

or mendelian diseases gene transfer-based repression of specific gene function may provide

therapeutic opportunities for acquired illnesses including heart disease and cancer where gene

therapy may have its greatest public-health impact (see Scollay et al. for a review (Scollay

2001)). Efficient transfer of vectors encoding therapeutic transgenes is key to the success of

any aforementioned gene-based therapy. Major challenges include precise integration of the

therapeutic transgene on the target chromosome as well as its timely and adjustable

expression. Currently available gene transfer technologies can be divided into two categories:

viral and non-viral vectors. Non-viral vectors also known as synthetic gene delivery systems

include liposomes, DNA-ligand conjugates, naked DNA, ballistic gene delivery and CaPO4


transfection (Curiel et al. 1992; Felgner et al. 1994; Geissler et al. 1994; Gao et al. 1995;

Dzau et al. 1996). The use of viral vectors for gene delivery holds great promise for basic

research and therapeutic applications. Viral vectors enable researches to monitor functions,

replace, correct, express or block expression of target genes, tag cells for fate determination or

change the physiological state of specific cell populations. At present, there are five major

classes of viral vectors used in clinical trials: oncoretroviruses, lentiviruses, adenoviruses,

adeno-associated viruses (AAVs) and herpes simplex-1 viruses (HSV-1s) (for review see Kay

et al. (Kay et al. 2001) and Koostra et al. (Kootstra et al. 2003)). Emerging viral transduction

systems include those derived from baculoviruses (Sarkis et al. 2000), alpha-viruses (Huang

1996) (for review see Schlesinger et al. (Schlesinger et al. 1999)), and vaccinia viruses which

are currently in clinical trials for cancer gene therapy (Peplinski et al. 1998).

Viral vectors for gene therapy

Most of the oncoretroviral vectors currently designed for gene therapy applications are

based on the moloney murine leukemia virus (MMLV) (Miller et al. 1993), which was used

for the pioneering human gene therapy trials tailored for the treatment of severe combined

immunodeficiency (SCID) (Blaese et al. 1995). Retroviral vectors are typically pseudotyped

with glycoproteins derived from the vesicular stomatitis virus (VSV-G) to expand the tropism

of chimeric viral particles. Following infection of the target cell, the genomic RNA of

oncoretroviruses is reverse transcribed into linear double-stranded DNA by the virion’s

transcriptase (Telenitsky 1997). Since the oncoretroviral virion is devoid of any nucleus-

targeting sequences or protein complexes access to the target chromosome for stable

integration requires cell division. Although oncoretroviral particles can only transduce

mitotically active cells their efficient transgene delivery and random integration of therapeutic

genetic information into target chromosomes which ensures sustained gene expression

rendered them the preferred gene transfer tool for gene therapy trials in the past decade

(Miller et al. 1990; Roe et al. 1993). However, results from recent clinical trial raised

concerns about oncoretrovirus-mediated induction of protooncogenes and dedifferentiation

factors following integration on the patient’s chromosome (Li et al. 2002; Schroder et al.

2002). Indeed, 2 out of 11 patients successfully treated for SCID later developed a leukemia-

like disorder due to integration of the oncoretroviral vector adjacent to the oncogene LMO2

(Hacein-Bey-Abina et al. 2003). Therefore, technologies for site-specific integration of

therapeutic gene cargos at preselected inert chromosomal locations is a current research focus


of the gene therapy community (Groth et al. 2000; Olivares et al. 2002). The most

encouraging clinical results associated with retroviral gene delivery were two young children

both of which suffered from a type of inherited immune disorder known as SCID. Both

children continue to live a normal life although long-term reconstitution from transduced

progenitor cells remained rather low. Prior to gene therapy the children were treated with a

novel drug to complement missing adenosine deaminase (ADA) function which may have

prevented the selective outgrowth of the transduced progenitor cells (Bordignon et al. 1995).

Lentiviruses which belong to the retroviral family recently came into the limelight of

current gene therapy initiatives since they are able to transduce non-dividing cells (Sadaie et

al. 1998). Most available lentiviral vectors adapted for secure gene transfer have been derived

from human HIV-1. Their inherent tropism for CD4+ T cells and macrophages has prompted

therapeutic approaches to treat HIV infection and AIDS. Pseudotyped with VSV-G

glycoproteins lentiviral particles are able to transduce muscle cells including cardiomyocytes

(Kelm et al. in press), hepatocytes (Naldini et al. 1996), hematopoietic stem cells (Naldini et

al. 1996; Kafri et al. 1997; Miyoshi et al. 1999), lung cells (for the treatment of cystic fibrosis;

(Goldman et al. 1997)), neurons (for gene therapy of Parkinson’s disease (Blomer et al.

1997)) and even chicken and insect cells (Mitta et al. 2002). Because HIV-1 is a human

pathogen, there is ongoing concern about the use of HIV-1-based lentiviral vectors for gene

therapy although third-generation lentivectors are devoid of recombination-competent

nucleotide stretches and fail to deliver any virus protein-encoding cistrons to the target cells.

Nevertheless, chimeric vectors based on non-human lentiviruses continue to be a current

research focus of the gene therapy community although the risk associated with transduction

of such viruses remains to be established (Browning et al. 2001). HIV-1-based vectors are

already in clinical trials for the treatment of HIV infections, yet their use in other gene therapy

settings will have to await clearance of remaining safety issues (Browning et al. 2001; Ikeda

et al. 2003). A DNA-based vaccine containing human immunodeficiency virus type 1 (HIV-1)

was tested for safety and host immune response in 15 asymptomatic HIV-infected patients

raising hopes since vaccine administration failed to induce local or systemic reactions

(MacGregor et al. 2000). Also, a phase I open clinical trial on the safety and tolerability of

single escalating doses of autologous CD4+ T cells transduced with HIV-antisense envelope

(env; VRX496) is currently conducted by the same group.

Adenoviral vectors have become a standard for gene therapy since they efficiently

transduce dividing as well as non-dividing cells, can be concentrated to high titers and

mediate high-level transgene expression from their episomal dsDNA genomes. A distinct


advantage of adenoviral vectors is their capacity to transduce target cells at low temperatures

which enables gene transfer prior to tissue or organ transplantation (Csete et al. 1994). First-

generation adenoviruses which only harbored deletions in one or two early genes (E1 and E3)

elicited a strong immune response against viral proteins which resulted in rapid elimination of

transgene expression (Kafri et al. 1998). In 1999, a patient treated with transgenic second-

generation adenoviruses against ornithine transcarbamylase deficiency even passed away

during a clinical trial (Raper et al. 2002). The development of the gutless adenoviral vector

which is devoid of most viral sequences prevented elimination of transduced cells by the host

immune system, failed to elicit inflammation in the target organ and showed reduced cellular

infiltration (Morsy et al. 1998). Yet, immune responses specific for the antigenic viral capsid

proteins cannot be alleviated to date. Gutless adenoviral vectors enable prolonged transgene

expression from their episomal non-replicating genomes. Thus, in dividing cells, transgene

expression may be lost over time due to dilution of episomal virions while transgenic

adenoviral episomes are prone to degradation in non-dividing cells. Adenoviral gene delivery

is currently the number-two transduction system in clinical trials. Adenoviral vectors seem to

be best for the treatment of (i) vascular and coronary artery diseases in which transient

transgene expression is preferred (Laitinen et al. 2000; Zhu et al. 2000), (ii) in therapies which

require short-term expression (Baltzer et al. 2000; Lai et al. 2001) and (iii) for cancer gene

therapy in which cellular toxicity and immunogenicity may increase antitumor effects

(Crystal et al. 1997; Brenner et al. 2000). The most prominent example of adenovirus-based

cancer gene therapy (already in phase II clinical trials) is the mutant adenovirus ONYX-015,

which is devoid of E1B gene expression. ONYX-015 can replicate in and lyse p53-deficient

target cells but remains replication-deficient in host cells expressing functional p53.

Mutations in p53 continue to be the major cause of cancer in humans (Bischoff et al. 1996;

Heise et al. 1997; Khuri et al. 2000).

Like adenoviruses, adeno-associated viruses have a broad tissue tropism. However,

transgenic particles derived from adeno-associated viruses integrate stably at specific sites on

the target chromosomes of dividing and non-dividing cells while showing little cytopathy

(Linden et al. 1996). Patients who received intramuscular injections of AAV vectors

transgenic for human factor IX expression in a clinical trial for hemophilia B showed

significant clinical benefits associated with reduced requirements for factor IX infusions. In

addition, these patients tolerated high AAV doses without eliciting vector-associated toxicity

or showing signs indicative for germ line transmission (Kay et al. 2000). Other clinical trials

for liver-based treatment of hemophilia are currently underway as are efforts for AAV-based


complementation of cystic fibrosis (protocols and abstracts available at

www.wiley.co.uk/genmed/clinical/ & www.dhhs.gov/). Recent advances in AAV vector

design included split units which concatamerize in a head to tail orientation upon transfer into

the nucleus. However, controversial results regarding efficiency of the system suggested that

further refinement of this concept will be required to solve packaging size limitation

associated with AAV vectors (Sun et al. 2000; Nakai et al. 2002). More recently, hybrid viral

vectors have been designed in an effort to combine transduction efficiency of adenoviral

vectors with stable long-term transgene expression associated with AAV and retroviruses

(Linden et al. 1996; Zheng et al. 2000).

Recombinant herpes simplex viral (HSV) vectors display a large transgene cloning

capacity. Currently, the major disadvantage of HSV-based vectors is their cellular and

immunological toxicities. Development of amplicon vectors, which are containing the HSV

origin of replication plus packaging signal and helper-free packaging systems which alleviate

contamination with replicating helper virus have further reduced cytopathic effects and

induction of immune responses of HSV-based transduction systems (Spaete et al. 1982;

Cunningham et al. 1993). The high neurotropism of HSV-derived vectors has initiated

therapies for malignant glioma (Markert et al. 2000). Additional clinical trials for HSV-

derived treatment of malignant melanoma and colon cancer are currently underway (protocols

and abstracts available at www.wiley.co.uk/genmed/clinical/ & www.dhhs.gov/).

Although oncoretroviral, lentiviral, adenoviral, AAV- and HSV-based vectors are all

currently used in clinical trials every application requires careful consideration of particular

characteristics associated with each transduction system (Table 1). Future advances in viral

transduction technologies will likely witness decisive refinements to increase gene-transfer

efficiency and safety. This may be achieved by generating chimeric viruses which harbor the

best genetic traits of several currently available systems (Lieber et al. 1999; Zheng et al.

2000). Also, ideal viral gene-transfer vectors will enable precise dosing of therapeutic

transgene in a gene therapy setting (see below).


Table 1: Comparison of viral vectors for gene delivery in clinical trials.

Gene regulation

Control of expression levels and duration of transgenes encoded on viral vectors

continues to be a challenge. First-generation retroviral vectors only supported constitutive

proviral transgene expression driven by the 5’ long terminal repeat (5’ LTR) (Gale et al.

2000). Then, heterologous promoters were introduced to achieve and/or restrict transgene

expression. Therefore, transcriptional targeting makes use of gene regulatory elements which

enable exclusive expression of therapeutic transgenes in specific cell types, for example the

promoters of (i) albumin in hepatocytes (Balague et al. 2000), (ii) synapsin and neuron-

Broad range, preferably

neurons

Broad range,

except for

hematopoietic

cells

Broad rangeBroad rangeOnly dividing

cells Target

Efficient transduction

of neural cells, largest

packaging

capacity

Integration into the target

genome, absence of

immune

response

Transduction of

a broad range of

cells, growth to

very high titers, efficient gene

expression

Integration

into the target

genome, Transduction

non-dividing

cells

Integration

into the target

genome,

absence

immune

response

Advantages

Disadvantages

Safety Issues

Pre-primed

patients

Inflamatory

Response

Transgene

Expression

Transgene Size

Genome

Cytotoxicity, transient

transgene

expression

Insertional

mutagenesis, Small

packaging

capacity,

safety

concerns, low

transduction

efficiencies

Episomal

transgene

expression,

Transgene

expression lost

over time,

immunogenic

Insertional

mutagenesis,

safety

concerns

associated

with HIV-derived

lentiviruses

Random

integration may

induce

oncogenesis,

low viral titers,

transduction limited to

dividing cells

Cytotoxicity Insertional

mutagenesis Inflamation

Insertional

mutagenesis

Insertinal

mutagenesis

Yes Yes Yes Only HIV-1

patients No

Strong (small,

amplicon) None

Strong

(small, gutless) NoneNone

transient stabletransientstablestable

40 Kb (150 Kb,

amplicon) 4.5 Kb

5-8 Kb (30-35 Kb,

gutless) 10 Kb8.5 Kb

dsDNA ssDNA dsDNA RNARNA

Herpes simplex

virus

Adeno-associated

virus AdenovirusLentivirusOncoretrovirus


specific enolase in neurons (Klein et al. 1998; Glover et al. 2003) and (iii) myosin light chain

1 in muscle cells (Shi et al. 1997). Furthermore, some regulatory elements have been reported

to promote transgene expression exclusively in tumors (e.g., carcinoembryonic antigen

(CEA)) (Lan et al. 1997) and hepatocellular carcinomas (e.g., alpha-fetoprotein, AFP) (Ido et

al. 1995; Cao et al. 1999).

More recently, the use of chimeric transcription control systems responsive to

antibiotics and other small molecules have enabled design of viral vectors for adjustable

transgene expression (see below for an overview on gene control systems). For example,

lentiviral vectors harboring tetracycline-, rapamycin- and progesteron analogue-responsive

gene expression components have provided regulated transgene expression following

transduction into a variety of different cell lines and tissues (Oligino et al. 1998; Pollock et al.

2000; Vigna et al. 2002). However, none of the transduction systems engineered for regulated

therapeutic transgene expression have reached clinical trials yet.

Perspectives

To date, viral transduction is the most efficient and flexible technology for transfer of

therapeutic transgenes into most cell types and tissues. Over 500 clinical trials have been or

are being conducted 60% of which focus on cancer treatment. Most of the vectors currently

enrolled in clinical trials are derived from oncoretroviruses and adenoviruses. However, a

great percentage of these vectors are still assessed for their safety following a particular gene-

delivery protocol (phase I). Detailed information on past and ongoing clinical trials is

available on the homepages of the National Institutes of Health (www.dhhs.gov/) or the

journal of gene medicine (www.wiley.co.uk/ genmed/clinical/).

Adjustable Transgene Expression

Control of gene expression in space and time is crucial for many applications in gene

therapy and tissue engineering. Several studies have demonstrated, that different levels of

important transcription factors govern distinct fates in differentiation and metabolism

(Duncan et al. 1998). Niwa and coworkers have expressed the transcription factor Oct-3/4 at

three precise levels, which resulted in three distinct fates of embryonic stem cells (endoderm,

retention of pluripotent phenotype, and trophectoderm) (Niwa et al. 2000). Another example

of therapeutic intervention which requires precise dosing is expression of the vascular

endothelial growth factor (VEGF), which has a high therapeutic potential for the induction of


angiogenesis/arteriogenesis in ischemic tissues (see above). Yet, sustained VEGF expression

results in the development hemangiomas which may even be fatal in mice (Carmeliet 2000).

Key characteristics of an ideal gene regulation system

Gene therapy and tissue engineering require effective gene regulation systems which

comply with the following criteria at a high standard: (i) high-level expression under induced

conditions, (ii) low leaky expression under repressed conditions, (iii) bioavailability and (iv)

pharmacokinetic profiles in favouring rapid switching of the transgene expression status, (v)

low immunogenicity of the gene control configuration, (vi) lack of interference with host

regulatory networks and (vii) compact genetic design to limit pleiotropic effects associated

with repeated molecular interventions on the host chromosome.

Antibiotic-controlled gene regulation systems

The antibiotic-controlled gene regulation systems are prevalent in basic research and

clinical applications. These systems consist of antibiotic-responsive promoters, transcription

modulators (activators and repressors) and antibiotics, which adjust binding of the modulators

to cognate operator sequences contained in the target promoters (Figure 3). The transcription

modulators consist of prokaryotic DNA-binding proteins fused to transcription activator (for

example, VP16 of Herpes simplex virus, (Triezenberg et al. 1988) (OFF-systems) or the

silencer domains (for example, KRAB of the human kox-1 gene (Moosmann et al. 1997)

(ON systems). Transgene expression controlled by OFF-type systems are repressed by

regulating antibiotics, while induction of ON systems requires administration of regulating

substances. To date, three antibiotic-responsive gene regulation systems are available. The

pioneering Tet system is responsive to tetracycline antibiotics (Gossen et al. 1992). This

system has seen many refinements in the past decade and now consists of an entire family of

different tetracycline-dependent transactivators and promoters (Baron et al. 1997; Kamper et

al. 2002; Krueger et al. 2003). The Tet systems and their derivatives have been extensively

used for basic in vitro and in vivo research (Fux et al. 2001; Malleret et al. 2001; Mallo et

al. 2003) as well as in prototype gene therapy settings tailored for bone regeneration

(Moutsatsos et al. 2001), reversion of -thalassemia (Samakoglu et al. 2002), and anemia in

mice (Sommer et al. 2002).


In the past years, two novel antibiotic-responsive gene regulation systems have been

developed following the generic design principle of the Tet systems. The PIP and the E.REX

systems adjust desired transgene expression in response to clinically licensed antibiotics of

the streptogramin and macrolide classes (Fussenegger et al. 2000; Weber et al. 2002). These

systems are compatible with each other and enable simultaneous regulation of up to three

independent transgenes within a single cell thus providing unmatched regulation complexity

for future clinical interventions.

Figure 3: Molecular setup of antibiotic-, quorum-sensing- and temperature-responsive gene regulation

systems following the same design concept. A DNA-binding protein (DBP) consisting of a bacterial response

regulator fused to a mammalian transcription-activation domain (TA; typically Herpes simplex-derived VP16)

binds to a specific operator module and induces polymerase (poly)-mediated gene of interest (goi) transcription

from minimal promoters (Pmin) in a molecule-responsive manner. With the exception of rTetR- and TraR-derived

proteins which bind chimeric target promoters in the presence of regulating molecules, transactivator-promoter

interactions of all other systems only form in the absence of modulating substances.

Transgene control by chemically induced dimerization

While antibiotic-responsive gene control systems rely on antibiotic-induced allosteric

changes of the modulator’s DNA-binding affinity chemically induced dimerization (CID)

reconstitutes a chimeric transactivator by heterodimerizing a DNA-binding and a

transactivating domain in an inducer-dependent manner. Latest-generation CID systems use

Operator Pmin goi pA

DBP

TA

DBP

TA

DNA-binding Protein (DBP’s)

•TetR

•rTetR

•PIP

•E

•TraR

•scbR

•RheA (at 37 °C)

Poly

Removed from operator by

•Tetracycline

•Absence of Tetracycline

•Pristinamycin

•Erythromycin

•Absence of 3-oxo-C8-HSL

•SCB1

•41 °C

Operator Pmin goi pAOperator Pmin goi pA

DBPDBP

TATA

DBP

TATA

DNA-binding Protein (DBP’s)

•TetR

•rTetR

•PIP

•E

•TraR

•scbR

•RheA (at 37 °C)

PolyPoly

Removed from operator by

•Tetracycline

•Absence of Tetracycline

•Pristinamycin

•Erythromycin

•Absence of 3-oxo-C8-HSL

•SCB1

•41 °C


rapamycin and non-immunosuppressive derivatives to heterodimerize an artificial ZFHD1-

binding domain fused to three tandem copies of FKBP with FRAP fused to the p65

transactivation domain of human NF- B (Rivera et al. 1996). In the presence of rapamycin

ZFHD-FKBP3-(FRAP-p65)3 binds and activates chimeric promoters containing twelve

ZFHD1 binding sites (Rivera et al. 1996; Pollock et al. 2002). The rapamycin-induced CID

system has been reported to mediate low leaky expression under non-induced conditions.

However, only sophisticated versions of the system reach the maximum expression levels

typically associated with the Tet and other antibiotic-responsive transcription control systems

(Go et al. 2002). The rapamycin-based CID system has already been successfully used for

gene therapy approaches in animal models (Auricchio et al. 2002; Auricchio et al. 2002).

Hormone-inducible gene expression

Prior to the advent of antibiotic-adjustable and dimerizer-inducible transcription

control systems technology for hormone-responsive transgene modulation had been

developed. Steroid receptors belong to a family of ligand-inducible transcription factors

which contain different DNA- and hormone-binding domains (Beato 1989). Fusion of the

hormone-binding domain to the DNA-binding domain of the yeast Gal4 protein as well as the

VP16 transactivation domain created an artificial transactivator whose binding to promoters

harboring specific Gal4 operator sites was hormone-adjustable. Three hormone-dependent

transcription control systems have been designed which are adjustable by either estradiol

(Braselmann et al. 1993), the progesterone analogue RU486 (Wang et al. 1994) or the insect

molting hormone ecdysone (No et al. 1996). However, due to pleiotropic impacts of these

hormones/hormone analogues on human physiology their use in the clinics may be limited.

Quorum sensing-based transgene modulation

Prokaryotes manage inter- and intrapopulation communication by quorum-sensing

molecules including acylated homoserine lactones which bind to receptors in target cells and

initiate specific regulon switches by modulating the receptor’s affinity to cognate promoters

(Bassler 2002). This bacterial cross-talk system has recently been adapted for controlling

transcription of desired transgenes in mammalian cells (Neddermann et al. 2003; Weber et al.

2003). Homoserine lactone-based mammalian gene regulation systems follow the generic

design principle of aforementioned antibiotic-adjustable control modalities. Despite clinical


compatibility ongoing use of antibiotics for controlling therapeutic transgene expression

remains controversial. Non-physiologically active regulating agents devoid of any pleiotropic

side effects would be preferred inducers. First in vivo validation of butyrolactones for

regulation of transgenes in mice did not indicate any side effects (Weber et al. 2003). Yet,

studies including long-term administration of this substance remain to be done. The existence

of near infinite bacterial quorum-sensing molecules/receptor pairs provides an inexhaustive

pool for the design of novel mammalian gene control systems in the not-too-distant future.

Temperature-dependent gene regulation

Fine-tuning of heterologous transgene expression in the absence regulating molecules

would represent a decisive advantage for manufacturing of difficult-to-produce protein

therapeutics. Two low-temperature-inducible gene regulation systems have been designed

(Boorsma et al. 2000; Weber et al. 2003): (i) Following a temperature shift from 37°C to

29°C, a mutated viral RNA-dependent RNA replicase initiates an RNA amplification loop

resulting in high expression levels of the transgene (Boorsma et al. 2000). (ii) The second

cold-inducible gene control system uses a Streptomyces albus-derived thermosensor whose

binding and activation of chimeric promoters could be modulated by 1°C steps between 37°C

and 41°C in DT-40 cells (Weber et al. 2003). Application of temperature-controlled gene

regulation systems will likely remain restricted to in vitro use as environmental temperature

changes are incompatible with precise titration of transgene expression in clinical settings.

Gene regulation systems in drug discovery

Applications of gene regulation systems go far beyond the mere regulation of desired

transgenes. Instead of using known antibiotics to regulate a specific transgene, gene

regulation systems configured for expression of a reporter gene can be used to find novel

inducing substances. Since prokaryotic antibiotic resistance regulators used for the desgin of

tTA, PIT or ET recognize almost any commercially available tetracycline, streptogramin or

macrolide antibiotic those proteins are likely able to detect antibiotic core structures. Aubel

and coworkers established a mammalian screening system for the detection of streptogramin

antibiotics. Chinese hamster ovary (CHO) cells transgenic for pristinamycin-responsive SEAP

(human secreted alkaline phosphatase) expression were used to screen for the presence of

streptogramin antibiotics in culture supernatants of Streptomyces isolates. This screening


system was sensitive compared to traditional antibiogram-based assays and provided a three-

in-one readout: (i) streptogramin core structure, (ii) bioavailability and (iii) non-cytotoxicity

of the detected substance (Aubel et al. 2001). Weber and coworkers refined this approach by

capitalizing on the antibiotic-responsive binding of bacterial resistance-response proteins, the

antibiotic “biosensors”, to their cognate operator sequences. The ELISA-type molecular

biosensor enables the detection of tetracycline, streptogramin and macrolide antibiotics in

spiked liquids including milk and serum with nanogramm precision. Binding of the

biosensors to their cognate DNA chemically linked to a solid surface is converted into an

immuno-based colorimetric readout correlating with specific antibiotic concentrations. Such

technologies will likely have far-reaching implications in the development of novel anti-

infective drugs and the enforcement of antibiotic bans in stock farming.

Use of gene-control systems in biopharmaceutical manufacturing

Regulated gene expression systems are not only of key interest for gene therapy

interventions, but also for reprogramming of mammalian cells lines to increase production of

protein therapeutics. Chinese hamster ovary (CHO-K1)-derived production cell lines

engineered for conditional expression of the cycline-dependent kinase inhibitors p21 or p27

arrested in the G1-phase of the cell cycle and exhibited up to 30-fold higher specific

productivities compared to proliferation-competent cells (Fussenegger et al. 1998; Meents et

al. 2002). Similar studies with hybridoma cells confirmed increased production of

proliferation-controlled cell lines (Watanabe et al. 2002).

Outlook

Although gene therapy originally set out to cure genetic diseases, most of today’s

clinical research and trials focus on the treatment of acquired multigenic disorders including

cancer and heart illnesses. These diseases have a more complex cause and require the

regulated expression of more than one therapeutic transgene. Next-generation gene therapy

scenarios will have to cope with the replacement of entire regulatory networks and signaling

cascades. Major parts of the toolbox required for multiregulated multigene therapeutic

interventions have recently been completed. Multicistronic and bidirectional expression

configurations enable simultaneous regulation of several transgenes (Baron et al. 1995;


Fussenegger et al. 1998; Fux et al. 2003; Fux et al. 2003). Furthermore, recent studies have

shown that all antibiotic-responsive gene control system are compatible and could be used for

independent control of up to three different transgenes (Weber et al. 2002).

Current approaches have linked different compatible gene regulation systems such that

the output of a first-level adjustable promoter controlled expression of a transactivator or

transrepressor which in turn activates expression of a second-level promoter which then

drives desired transgene transcription (Aubrecht et al. 1996). In some configurations target

promoters were designed to be responsive to different transactivators and could integrate

several internal signal into a one-gene expression readout (Hoshikawa et al. 1998; Freundlieb

et al. 1999). For example, Imhof and coworkers conceived a regulatory network consisting of

a hybrid promoter which was downregulatable by a constitutively expressed TetR-derived

transrepressor and inducible by a Gal4-VP16 transactivator (Imhof et al. 2000). Since Gal4-

VP16 was under control of its own target promoter this configuration enabled autocatalytic

coexpression of the transactivator and the gene of interest.

Recent advances in artificial gene network design capitalized on cascade-like

interconnection of three independent gene control systems. In this configuration the

tetracycline-dependent promoter drives tTA expression in an autoregulatory manner along

with cocistronic expression of the macrolide-dependent transactivator (ET). ET targets its

macrolide-responsive promoter controlling expression of the pristinamycin-dependent

transactivator (PIT) which in turn fine-tunes transcription of the gene of interest. When

transcription modulation is exerted by addition of either tetracycline, erythromycin or

pristinamycin the three-step regulatory network exhibited unprecedented regulation

characteristics: (i) absence of regulating antibiotics resulted in maximum transgene

expression, (ii) addition of tetracycline yielded precise 70% expression, (iii) administration of

erythromycin reduced expression of the gene of interest to 40% of maximal levels and (iv)

was completely repressed following addition of pristinamycin (Figure 3) (Kramer et al. 2003).

The three-step regulatory network enabled precise expression levels following administration

of clinical doses of regulating antibiotics. Adjusting desired transgenes using a single

antibiotic-responsive gene regulation system requires dosing of the specific antibiotic with

nanogram precision which remains challenging in the human body.

Advanced gene network including toggle switches (Figure 4) (Gardner et al. 2000),

oscillators (Elowitz et al. 2000), and logical transcription control (Hasty et al. 2002; Buchler

et al. 2003) which enable new dimensions in artificial transgene control have only been put

into practice in Escherichia coli. Until functional implementation of these latest-generation


control networks in mammalian cells will become a scientific reality molecular progress will

have to ensure that such systems are controllable, reliable and devoid of any genomic cross-

talk prior to their implementation in the clinics (Weiss et al. 2002). Gene circuit engineering

in mammalian cells is expected to enable complex multigene-based therapeutic interventions

which will foster decisive advances in regenerative medicine (Kitano 2002).

Figure 4: Examples of complex artificial gene regulatory networks. (A) Eukaryotic regulatory cascade

which enables precise expression fine-tuning to four distinct levels. This regulatory cascade consists of three

antibiotic-responsive gene regulation systems arranged in series, such that the output of one promoter directly

influences expression of the subsequent one: (i) The tetracycline-responsive promoter (TetO7-Pmin) is activated

in an autocatalytic way by the tetracycline-dependent transactivator (tTA) cocistronically expressed with the

macrolide-dependent transactivator (ET). (ii) ET activates transcription of the streptogramin-dependent

transactivator (PIT) driven by the macrolide-responsive promoter (ETR8-Pmin). (iii) Expression of the gene of

interest (goi) is eventually driven by the streptogramin-responsive promoter PIR3-Pmin following PIT-mediated

transactivation. Expression from this three-step regulatory cascade can be modulated by either tetracycline (Tet),

erythromycin (EM), or pristinamycin (PI). Addition of a specific antibiotic (2 g/ml) correlates with fixed goi

expression profiles: (i) 100%, no antibiotics, (ii) 70%, tetracycline, (iii) 40%, erythromycin, (iv) no expression,

pristinamycin. (B) The prokaryotic toggle switch consists of two repressible promoters arranged in a mutually

inhibitory manner. Each promoter controls expression of the repressor for the other one. Such a setup exhibits

bistable expression characteristics. If the promoter PLac is induced by transient addition of IPTG, expression of

the green fluorescent protein (GFP) is switched on. Following removal of the inducer IPTG, GFP expression

levels remain high owing to concomitant expression of the PL-specific repressor cI which prevents PL-mediated

Pmin tTA pAIRES ETTetO7

Pmin PITETR8 pA Pmin goiPIR3 pA

Tet

PIEM

Inactivation

Activation

A

B

Plac cI pAIRES gfp lacIPL pA

IPTG 42 °C

Pmin tTA pAIRES ETTetO7

Pmin PITETR8 pA Pmin goiPIR3 pA

Tet

PIEM

Inactivation

Activation

Inactivation

Activation

A

B

Plac cI pAIRES gfp lacIPL pA

IPTG 42 °C


expression of the Plac repressor LacI. GFP expression will be repressed following a temperature shift to 42 °C

which induces PL and promotes LacI-mediated shut down of Plac.

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181

Acknowledgements

I would like to take the chance to thank all those people who worked with me, helping to

improve the final outcome of this thesis.

Prof. Dr. M. Fussenegger: for giving me the opportunity to carry out my research in your

research group and to take advantage of your scientific experience. I appreciated very much

the scientific freedom and your scientific advice.

Prof. Dr. S. Werner: for being my coexaminer.

Dr. Elisabeth Ehler: for introducing me into the world of fluorescent antibodies and confocal

microscopy. I welcomed very much that I could ask every question to understand the cell

biology of heart muscle cells.

PD Dr. Valentin Djonov: for getting me into contact with the CAM analysis and interpretation

of histological specimens.

Prof. Dr. Simon Hoerstrup: for your advice.

Prof. Dr. Jean-Claude and Evelyne Perriard: for supplying me with primary cardiomyocytes

and suggestions whenever it was necessary.

Dr. Lars M. Ittner: for your motivating discussions and excellent help during my thesis.

Nadja Guiliani: for your continuous help and contribution to this work during your

apprenticeship.

Carlota Diaz Sanchez-Bustamante and Bettina Ley: for your ambitious support during your

diploma thesis.

David Fluri: for standing by my side in many battles and for creating a wonderful working

atmosphere in the lab.

182

Stefan Schlatter: for introducing me into the secrets of molecular biology and the conventions

of Swiss people. Unfortunately you left already after 1.5 years the lab to go to Australia.

The rest of the Fussi group and Peter Steiner: for their help and beer drinking assistance.

JENS KELM DEPARTMENT OF SURGICAL RASEARCH UNIVERSITY HOSPITAL

CH-8091 Zurich [email protected]

183

CURRICULUM VITAE

PERSONAL DATA

Name: Jens Kelm, Dipl. Biotech.

Date of Birth: 24 June 1971, Muelheim / Ruhr, Germany

Nationality: German

Current Position: Post-doctoral fellow at the University Hospital

Work Address: Jens Kelm

University Hospital Zurich

Tissue Engineering and CellTransplantation

LAB F-41

Raemistrasse 100

CH-8091 Zurich, Switzerland

Private Address: Jens Kelm

Baerenbohlstrasse 35

CH-8046 Zuerich, Switzerland

Tel: +41 1 3718328



184

EDUCATION

April 2005 – present Scientific assistant at the division of Tissue Engineering

and cell transplantation at the department of surgical

research and Clinic for Cardiovascular Surgery,

University Hospital, Zurich, Switzerland.

January 2001 – March 2005 Doctoral fellow at the Institute of Biotechnology, ETH

Zuerich, Switzerland; Ph.D. thesis under supervision of

Prof. Dr. Martin Fussenegger, and Prof. Dr. Sabine

Werner.

Title: “Design of Artificial Microtissues”.

September 2000 Graduation in Biotechnology, at the Technical University

of Braunschweig, Germany

September 1999 – April 2000 Diploma thesis at the Institute of Chemical Engineering,

University of Queensland, Australia, under supervision

of Prof. Dr. Ralf-Rainer Mendel (TU-Braunschweig) and

Prof. Dr. Lars Keld Nielsen.

Title: “Developing an Experimental Cell Cluster Model

to Evaluate Strategies for Tissue Engineering

Applications”.

September 1998 – March 1999 Semester work at the German Research Centre for

Biotechnology (GBF), Braunschweig, Germany, under

supervision of PD Dr. Roland Wagner. Title: Adaptation

of Immortalized Human Hepatocytes to Protein-Free

Culture Conditions”.



185

October 1994 Started as a student in the biotechnology program at the

Technical University of Braunschweig, Germany.

April 1988 – July 1993 High school, Muelheim, Germany, receiving the high-

school diploma.

April 1980 – March 1986 Primary school, Muelheim, Germany.

APPRENTICESHIP

August 1992 – June 1994 Vocational education as Biological Technical Assistant

(BTA) at the full-time vocational school for technical

assistants, Olsberg, Germany.

PUBLICATIONS

Schlatter S., M. Rimann, J. Kelm, and M. Fussenegger (2002). “SAMY, a novel mammalian

reporter gene derived from Bacillus stearothermophilus alpha-amylase”. Gene 282, 19-31.

Mitta B., M. Rimann, M.U. Ehrengruber, M. Ehrbar, V. Djonov, J. Kelm, and M. Fussenegger

(2002). “Advanced modular self-inactivating lentiviral expression vectors for multigene

interventions in mammalian cells and in vivo transduction” Nucleic Acids Res. 1;30(21):e113.

Kelm, J. M., N. E. Timmins, C. J. Brown, M. Fussenegger and L. K. Nielsen (2003). “Method

for generation of homogeneous multicellular tumor spheroids applicable to a wide variety of

cell types.” Biotechnol Bioeng 83, 173-80.


"Design of Artificial Myocardial Microtissues." Tissue Eng. 10, 201-214



186


“Synergies of Microtissue Design, Viral Transduction and Adjustable Transgene Expression

for Regenerative Medicine.” Biotechnol Appl Biochem. 39, 3-16

Kelm J.M. and M. Fussenegger (2004). “Microscale tissue engineering using gravity-enforced

cell assembly.” Trends Biotechnol. 22, 195-202.

Fux C., D. Langer, J.M. Kelm, W. Weber, and M. Fussenegger (2004). “New-generation

multicistronic expression platform: pTRIDENT vectors containing size-optimized IRES

elements enable homing endonuclease-based cistron swapping into lentiviral expression

vectors.” Biotechnol Bioeng. 86, 174-87.

Kelm J.M., C. Diaz Sanchez-Bustamante, E. Ehler, S.P. Hoerstrup, D. Djonov, L.M. Ittner and

M. Fussenegger (2005). “VEGF profiling and angiogenesis in human microtissues.” J.

Biotechnol. (in press).

Diaz Sanchez-Bustamante C., J. M. Kelm, B. Mitta and M. Fussenegger (2005). “Heterologous

Protein Production Capacity of Mammalian Cells in 2D and 3D cultures”. (in press).

Kelm J.M., V. Djonov, L.M. Ittner, W. Born, S.P. Hoerstrup, and M. Fussenegger

(submitted). “Design of Custom-Shaped Vascularized Tissues Using Microtissue Spheroids

as Minimal Building Units ”.

Kelm J.M., V. Djonov, S. P. Hoerstrup, C. I. Guenter, L. M. Ittner, F. Greve, A. Hierlemann,

J. C. Perriard, E. Ehler and M. Fussenegger (submitted). “Tissue-Transplant Fusion and

Vascularization of Myocardial Micro-and Macrotissues Implanted into Chicken Embryos and

Rats.”



187

POSTER PRESENTATIONS

Kelm J.M., P.D. Munro, S. Henning, S.C. Warren, C.J. Brown, and L.K. Nielsen (2001).

Homogenous Cell Clusters for Tissue Engineering Applications; The 17th

meeting of the

European Society of Animal Cell Technology Meeting, Tylösand, Sweden

Kelm J.M., E. Ehler., N. Timmis, L.K. Nielsen, M. Fussenegger (2001). “Exploring the Third

Dimension: Cell Culture Technology to Generate Microtissue” 1st Biennial Meeting of the

European Tissue Engineering Society, Freiburg, Germany

Kelm J.M., E. Ehler, and M. Fussenegger (2002). “Design of Artificial Microtissues”. 4. D-

Biol Symposium, Davos, Switzerland

Kelm J.M., Diaz-Sanchez-Bustamante C., and M. Fussenegger (2005). “VEGF Profiling and

Angiogenesis in Human Microtissues“.The 19th

meeting of the European Society of Animal

Cell Technology Meeting, Harrogate, United Kingdom

ORAL PRESENTATIONS

Design of Artificial Myocardial Microtissues, (2003) The 18th

meeting of the European

Society of Animal Cell Technology Meeting, Granada, Spain

Design of Artificial Microtissues: A Novel Concept for Cell-based Therapies and Tissue

Engineering, (2005) Biochemical Engineering XIV, Harrison Hot Springs, Canada

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