Development of a myocardial infarction

model by the design of a miniaturized

3D culture platform

T.M.P.M. Boonen S1204718

18th of October 2019

Faculty of Science and Technology Department of Applied Stem cell technologies

EXAMINATION COMMITTEE Chairman: Prof. Dr. P.C.J.J. Passier Daily supervisor: Dr. M. Catarino Ribeiro External member: M. Odijk

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Abstract

Currently the leading cause of death in the western world is cardiovascular disease (CVD) and in the

future the amount of people suffering from CVD is only projected to rise. However high-throughput

disease models are lacking. Since the generation of the first induced pluripotent stem cells (iPSCs),

these iPSCs have become of great interest in the field of drug testing and disease modelling. The big

problem with cardiomyocytes derived from iPSCs (iPSC-CM) is that after differentiation these cells

show an immature phenotype which manifests itself in an immature metabolism. Because of this

immature metabolism they cannot yet be used in modeling CVD. During this project an attempt has

been taken at maturing iPSC-CMs in two dimensional (2D) cell culture, by alteration of the carbon

source available in the medium. Unfortunately no mature metabolism was achieved and that is why

the focus shifted towards three dimensional (3D) culture possibilities. A 3D culture method of

suspending a strip of cardiac cells between two flexible pillars called an engineered heart tissue (EHT)

has been incorporated into a microfluidic platform. This platform, the microEHT array, allows for easy

parallel culture of miniaturized EHTs for over two months.

Keywords:

Myocardial infarction, cardiomyocytes, induced pluripotent stem cells, oxidative phosphorylation,

glycolysis, engineered heart tissue.

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Table of Contents Abstract ....................................................................................................................................................ii

I. Introduction .......................................................................................................................................... 1

1.1 Cardiovascular disease ................................................................................................................. 1

1.2 Human Induced Pluripotent Stem Cell derived CMs ................................................................... 1

1.3 Energy Metabolism ....................................................................................................................... 1

1.4 Myocardial infarction ................................................................................................................... 3

1.5 Cardiomyocyte maturation .......................................................................................................... 3

1.6 2D vs 3D models............................................................................................................................ 4

1.6.1 Engineered Heart Tissue ........................................................................................................ 4

1.6.2 Organ on a Chip ..................................................................................................................... 5

Scope of the internship ........................................................................................................................... 6

II. Materials and methods ....................................................................................................................... 7

2.1 Cell culture .................................................................................................................................... 7

2.1.1 hESC differentiation towards cardiomyocytes ..................................................................... 7

2.1.2 Dissociation of hESC derived cardiomyocytes ...................................................................... 7

2.1.3 Thawing hESC derived cardiomyocytes ................................................................................ 7

2.2 2D metabolism maturation .......................................................................................................... 7

2.2.1 Lactate Assay ......................................................................................................................... 8

2.2.2 ATP assay ............................................................................................................................... 8

2.3 3D disease model .......................................................................................................................... 8

2.3.1 Computer aided design (CAD) ............................................................................................... 8

2.3.2 Micro-milling .......................................................................................................................... 8

2.3.3 Cleanroom wafer production ................................................................................................ 9

2.3.4 Fabrication microfluidic chips ............................................................................................... 9

2.3.5 Preparation of a fibrin-based hydrogel ................................................................................. 9

2.3.6 Seeding microfluidic chips ..................................................................................................... 9

2.3.7 Crosslinking tailoring ........................................................................................................... 10

2.3.8 Microfluidic surface treatment ........................................................................................... 10

III. Results .............................................................................................................................................. 11

3.1 2D Maturation ............................................................................................................................ 11

3.2 3D disease modelling .................................................................................................................. 15

3.2.1 MicroEHT .............................................................................................................................. 15

3.2.2 MicroEHT array .................................................................................................................... 17

3.2.3 MicroEHT array cell culture ................................................................................................. 18

3.2.4 Upscaling microEHT size ...................................................................................................... 20

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3.2.5 Fibrinogen crosslinking time ............................................................................................... 21

3.2.6 Fouling of the microEHT array ............................................................................................. 22

IV. Discussion ......................................................................................................................................... 23

4.1 2D maturation ............................................................................................................................. 23

4.2 3D disease modelling .................................................................................................................. 23

4.3 Fibrinogen crosslinking time ...................................................................................................... 24

4.4 Fouling of the microEHT array .................................................................................................... 24

V. Conclusion ......................................................................................................................................... 25

VI. Future perspective ........................................................................................................................... 25

Acknowledgements ............................................................................................................................... 26

Bibliography ........................................................................................................................................... 27

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I. Introduction

1.1 Cardiovascular disease Currently the leading cause of death in the western world is cardiovascular disease (CVD) [1]. In the

future the amount of people suffering from CVD is only projected to rise [2]. Global cardiovascular

deaths are expected to rise from 16.7 million in 2002 to 23.3 million in 2030. In part, this is due to there

not being any treatments for damage to the heart muscle because of a myocardial infarction (MI).

Another reason for the expected rise is that about 50% of patients is nonresponsive to the current drug

therapies [3]. A consequence of this is that a lot of people are suffering from debilitating and ultimately

fatal effects of CVD. Therefore novel patient specific drug testing and individualized therapies are

needed.

Apart from CVD being a major burden on the worldwide health, another reason novel drug testing

strategies are needed is that cardiotoxicity is one of the principle forms of drug toxicity. Cardiotoxicity

accounts for most of the major drug recalls even in non-cardiac specific drugs [4]. Current drug testing

strategies heavily rely on animal testing, however fundamental differences exist between human and

animal cardiomyocytes (CMs) e.g. electrophysiological and contractile properties and

pharmacokinetics [5], [6].

1.2 Human Induced Pluripotent Stem Cell derived CMs Since the generation of the first induced pluripotent stem cells (iPSCs) by Yamanaka [7], these iPSCs

have become of great interest in the field of drug testing and personalized medicine. Most labs around

the world can now generate CMs from these iPSCs and therefore create patient specific disease models

[8], [9]. These human iPSC-CMs are generally differentiated by adding growth factors and small

molecules (e.g. Activin A, BPM4 and CHIR) at set time points to the iPSCs.

The big problem with iPSC-CMs is that after differentiation these cells show an immature phenotype.

This manifests itself in an immature metabolism, morphology, gap junction expression, contractility,

electrophysiology as well as immature calcium handling properties [10]. Overall these cells show a

fetal-like phenotype.

1.3 Energy Metabolism The adult heart has a high demand of energy, due to its continuous contracting motions to supply the

entire body with blood. Despite having this high-energy demand, the heart muscle has virtually no

energy reserves and therefore it must continually produce energy. The adult heart normally produces

90% of its energy through mitochondrial oxidative metabolism. For this, mainly fatty acids are used.

However, the heart is an “omnivore” and can use a plethora of fuels, including lactate, glucose,

amino acids and ketones [11]. On the contrary, fetal CMs seem to produce most of their energy

through glycolysis, with the mitochondrial oxidative metabolism being poorly developed. The

dependency of the fetal heart on glycolysis as an ATP-generating pathway stems from it residing in

an environment with low oxygen [12], [13]. The immature iPSC-CMs, similarly to the fetal CMs,

mainly produce energy through glycolysis. Immediately after birth, close to half of the total produced

ATP is obtained through glycolysis. 7 days into the newborn period, glycolysis decreases and

generates less than 10% of total ATP production, which is similar to values seen in the adult heart.

[14] (Fig. 1.1)

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Figure 1.1 A| Overview of the various pathways of energy substrate metabolism that

contribute to the production of ATP. B| Percent contributions of glycolysis and

oxidative phosphorylation (OXPHOS) (glucose oxidation, lactate oxidation, and fatty

acid oxidation) to ATP production during the immediate newborn/fetal period and

the neonatal period (from 7 to 21 days) in an isolated working rabbit heart (B). Taken

from [14]

Figure 1.2 The electron transport chain. Electrons pass through the ETC releasing energy,

which is used to create a proton gradient that fuels ATP synthase. Hereby

generating energy in the form of ATP. Taken from [16]

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Concurrently with the switch in preferred metabolite, maturation of mitochondria occurs, enabling

this switch. In the fetal CM the mitochondria are characterized by being small and immature with

underdeveloped cristae. They are located principally in the nuclear periphery and form a poor

network. Adult CMs are defined by mitochondria that have adopted a more tubular structure,

contain elongated cristae, and occur all throughout the cytoplasm. The development of the

mitochondria in fully matured cells is characterized by the initiation of oxidative phosphorylation

(OXPHOS) as evidenced by increased mitochondrial membrane potential, elevated oxygen

consumption, and increased adenosine triphosphate (ATP) production. [13], [15]

Oxidative phosphorylation is the result of a series of energy transformations, where firstly a carbon

source is oxidized in the Krebs cycle to yield electrons. These electrons are transported towards the

electron transport chain (ETC) by NADH and FADH2. As the electrons pass through the ETC they

release energy, part of which is used to pump protons out of the mitochondria. At the end of the ETC

the electrons are used to split O2 to form water. The previously pumped out protons are taken up

again by ATP-synthase to generate ATP. [16], [17] (Fig. 1.2)

1.4 Myocardial infarction Myocardial infarction mostly occurs upon blockage of a coronary artery. Usually this is caused by the

formation of a blood clot originating from a ruptured fatty deposit (plaque) [18]. During the time of

blockage (ischemia), cardiomyocytes are restricted in both their nutrient and oxygen supplies,

inhibiting the cells usage of OXPHOS, causing a significant depletion of the little ATP reserves.

Consequently this will lead to disruptions in the cardiac conduction system and deterioration of cardiac

contractility, which may result in arrhythmias and heart failure, respectively [19].

Especially when modelling myocardial infarction (MI) the fetal-like metabolism of the iPSC-CM is a big

problem. The principles of MI, the inhibition of OXPHOS, rest on the fact that cells are dependent on

oxygen to generate energy [20]. So the fact that the iPSC-CMs do not use oxygen to produce their

energy, means they are not sensitive to cell damage as a result of ischemia. To produce a model for MI

CMs are needed with a mature metabolism. So that when oxygen and nutrient availability are

restricted the CMs have the above-mentioned physiologically correct response.

1.5 Cardiomyocyte maturation Different strategies are being explored to shift the fetal-like glycolytic metabolism of iPSC-CMs towards

the more mature OXPHOS. Examples of this include: electrical stimulation [21], [22], mechanical

stretching [21], [23], [24], modelling of the carbon source [25], [26], prolonged culture time [27], [28],

or culturing the cells in a three dimensional (3D) environment [29]–[32].

Electrical stimulation has been shown to induce maturation in CM, but the parameter of maturation

that is quantified is usually only the electrophysiological maturation [21], [22]. However, this is just

one of the hallmarks of maturation. Other hallmarks such as the desired mature metabolism were not

tested. Similar results were obtained with regards to mechanical stretching. Cyclic stretch and strain

at 1–2 Hz promoted structural maturation, cell elongation and a higher degree of sarcomeric

organization [24]. Again no metabolic maturation was shown.

When hiPSC-CMs are cultured for over 1 year, maturation was observed with gradual changes in

sarcomere organization, cell elongation and cell size [28] However, there was considerable variability

among the cells analyzed. Therefore a recent review by Veerman et al. concluded that an adult

cardiomyocyte phenotype is never acquired by prolonged culture [27].

The most intuitive of the above-mentioned options is simply changing the availability of the carbon

source. Normal culture medium is comprised of high amounts of glucose and insulin forcing a cell

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towards glycolysis. It has been shown that, when cultured in medium in which the glucose is replaced

with galactose, iPSC-CM exhibit a shift in energy production from glycolysis towards OXPHOS [25], [33].

This happens because the production of pyruvate via glycolytic metabolism of galactose yields no net

ATP, making it energetically unfavorable. This forces these cells to have an increased reliance on

OXPHOS for energy [34].

1.6 2D vs 3D models Another line of research that is being pursued around the world is, going from a two dimensional (2D)

culture towards a 3D cardiac tissue. Just over twenty years ago, Eschenhagen et al were successful in

engineering a cardiac tissue from embryonic chicken CMs [29]. This started the field of 3D cardiac

tissue engineering. This field branched off into the development of in vitro substitutes of cardiac tissue

for regenerative medicine and preclinical drug development, and towards further advancing in

vitro models for heart function and disease [35]. Different ways of bioengineering 3D cardiac tissue

exist, including cell sheet technology [30], decellularized hearts [36], micro-patterns/devices [37], [38],

3D printing [31], [39] and hydrogel embedding [32], [40], [41].

All the mentioned methods of creating a cardiac tissue show great progress and promise. However,

they also come with their own difficulties. Stacking cell sheets allows for inclusion of different cell types

in a tissue, it also gets rid of the need of any artificial scaffold, because the cells generate their own

extra cellular matrix (ECM) [42]. However, some of the downsides of cell sheet technology are the

limited functional read outs that are possible on the tissue and the limited thickness that has been

achieved so far. De- and recellularizing heart tissue gained a lot of attention after Guyette et al. tried

to recellularize a complete human heart [43]. In spite of adding 500 million cells to the left ventricle of

the heart only 50% of the decellularized ECM contained cells after seeding. This highlights the

difficulties with this technique. Recellularization protocols still have to be optimized to make it viable.

3D printing of cardiac tissue is also a field which has seen a lot of development in recent years [44]. A

major topic that is being worked on is matching cell densities and mechanical properties of the printed

construct, to native tissue [45]. Another topic is the tradeoff of printing speed versus resolution. To

achieve a high resolution, currently the prints take a very long time. This resolution is needed to be

able to seed cells in close proximity to each other to from a tissue.

At the moment the most used strategy for making cardiac tissues is hydrogel embedding of CMs. It is

a simple and multiplexable way of generating tissues. By using a native ECM such as collagen, matrigel

and or fibrin, the cells are able to produce their own matrix interlocking with the native ECM, secrete

matrix metalloproteinases to re-arrange the ECM, and bind the matrix for migration, tension

generation, and tissue compaction. [46]

1.6.1 Engineered Heart Tissue Some of the most recent publications about 3D cardiac tissues use cells embedded in hydrogel calling

them by different names, e.g. human engineered cardiac tissues (hECT) [21], engineered human

myocardium (EHM) [41] or engineered heart tissue (EHT) [40], [47]. From hereon we will call a 3D

cardiac tissue an EHT.

Generally an EHT is comprised of several components including a casting mold, mechanical support,

cells (CM) and a hydrogel (Fig. 1.3). The first EHT was made of embryonic chick CM mixed with collagen

solution and allowed to gel between two Velcro-coated glass tubes [29]. Over the years all these

components have undergone improvements [48]. The embryonic chick CMs are replaced with human

cells to make the platform representable for humans. The collagen solution has been replaced with a

hydrogel based on the extracellular protein fibrinogen [49]. The benefit of this hydrogel is that the

thrombin-catalyzed polymerization is faster than collagen. Furthermore, the polymerization time can

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be tailored by varying the concentration of thrombin or through manipulation of the pH and ionic

strength of the reaction [50]. Two major benefits of the faster polymerization time are that a

homogeneous dispersion of cells is retained and the EHT can be supplemented with medium quicker.

Apart from this, also the casting mold and mechanical support underwent development in the past

years. Breckwoldt et al. demonstrate the possibility of creating a casting mold out of agarose [51]. The

benefit of this is that agarose can take any shape, which is only dependent on the mold placed into it

during cooling. This allows the generation of very specific tissue shapes. To this casted shape the cells

in gel can be added as well as the mechanical support. Instead of rigid glass tubes [29], flexible pillars

can be used. This is favorable because as soon as the fibrinogen is polymerized, by thrombin, the cells

start compacting and bending the flexible pillars. A thin muscle strip is formed under auxotonic loading.

The second advantage is that the bending of these pillars can be tracked to allow for an easy

quantification of contractility of the tissue. [51]

In a recent review of the progress of EHT development Mills et al stated that the capabilities of the

EHT can be further improved by downscaling current designs to generate miniaturized tissues for high-

throughput analysis. Miniaturized EHT have several advantages as they reduce cell numbers, cell

culture costs, manual handling requirements and are not restricted by oxygen diffusion limitations,

which is a problem with larger EHT constructs. [47] When developing a model for MI these advantages

are very useful since it makes it possible to produce the model with less cells, lower costs and less

manual handling.

1.6.2 Organ on a Chip Miniaturizing in vitro models has been achieved in recent years through the use of organ on a chip

models. An organ on chip is microfluidic device capable of mimicking key physiology and functions of

specific organs [52]. Many different organ have already been modelled with this technique, including

blood vessels, lung, and liver. Apart from miniaturization being possible, organ on a chips also allow

for culture under perfusion. Therefore concentration gradients of certain molecules (including drugs)

can be applied in an automated and time-controlled manner [53] and cells are not grown in a non-

physiological medium-to-cell ratio anymore [54].

Microfluidic devices are commonly made out of polydimethylsiloxane (PDMS) using soft lithography

because of advantages such as ease of fabrication, good cell culture compatibility, and optical

transparency [55]. The master molds for the PDMS chips are generally made using either micro-milling

or SU-8 photolithography in a cleanroom. Micro-milling can be done on a wide selection of working

materials and has versatile applications and rapid prototyping capability [56]. A design for a mold is

made with computer aided design (CAD) software, in which also a milling procedure is designed. In this

Figure 1.3 Engineered heart tissues. Schematic representations of previously developed EHTs.

A| The first engineered heart tissue (EHT) on Velcro-covered rods (Eschenhagen et

al. [29]) B|A fibrin-based EHT on PDMS (polydimethylsiloxane) racks (Hansen et al.

[48])

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procedure for example the mill/drill diameter, turning speed (spindle speed) and feed rate can be

tailored to ensure a smooth surface, which is necessary for a flawless microfluidic chip. In general, the

surface smoothness increases when the feed rate decrease [57]. The optimal spindle speed will differ

for every milling machine because of system-specific vibrations and resonance frequency. The final

design and optimal procedure is uploaded to a micro-milling device. This device will mill the mold, after

which residual debris can be cleaned from the mold.

Photolithography is a fabrication method that typically involves a photoresist, UV-mask and a silicon

wafer and is performed in a cleanroom. The silicon wafer is the base on which the structures that

comprise the mold are fabricated. This fabrication starts by depositing a photocrosslinkable polymer

(photoresist) onto the wafer. SU-8 is a common photoresist used in mold fabrication because of its

high resolution, mold durability and capacity for high aspect ratios [58]. Subsequently UV-exposure is

performed through a photomask to selectively crosslink the SU-8. Next, the SU-8 can be developed

and only the exposed structures remain. Multi-level structures of SU-8 can be fabricated by repeating

the steps of SU-8 deposition, photomask UV-exposure and SU-8 development.

The molds generated through either milling or photolithography can be used as a master mold onto

which the PDMS is poured and thermally cured. The cured polymer is peeled from the mold surface,

and it contains a replica of the mold. To obtain a microfluidic chip, this PDMS replica can be bonded

onto a glass slide after plasma treatment. The plasma treatment of PDMS introduces polar functional

groups which is mainly the silanol group (SiOH) [59]. This changes the surface properties of PDMS from

being hydrophobic to hydrophilic. However this change is only temporary since the PDMS undergoes

hydrophobic recovery. The main reasons for this are the reorientation of the polar groups from the

surface to the bulk, diffusion of pre-existing low-molecular-weight species from the bulk to the surface

and condensation of the hydroxyl groups [60].

After mold fabrication, soft lithography of PDMS and bonding onto a glass slide a microfluidic chip is

obtained. The combination of this versatile platform with the advantages of the EHT would yield an

EHT that can be cultured under perfusion with contractility tracking as a functional read out.

Scope of the internship The goal of this internship has been the creation of a model for myocardial infarction. An accurate

disease model is comprised of the following: mature cardiomyocytes, a culture platform and

functional read outs. To obtain a model closely resembling the in vivo situation human iPSC can be

differentiated into CM. However these cells show a fetal-like metabolism, using glycolysis instead to

OXPHOS. The first part of the project focusses on the maturatioin these hiPSC-CM through the

change of the available carbon source in the culture medium.

During the second part of the project a suitable culture platform for the hiPSC-CM was designed. This

platform combines the generation of EHTs with the benefits of organ on a chip technology. Allowing

for perfusion culture of a micro tissue, under auxotonic loading, of which contractility can be

measured.

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II. Materials and methods In this section culture procedures are described. Unless stated otherwise, cells were cultured in an

incubator at 37°C at 5% CO2.

2.1 Cell culture

2.1.1 hESC differentiation towards cardiomyocytes One day before differentiation the human embryonic stem cells (hESC) were plated on matrigel

(Corning®) coated 6-wells plate at a density of 250k cells/well. On day 0 BPEL (Bovine serum albumin

Polyvinyl alcohol Essential Lipids) medium containing the growth factors Activin-A (20ng/mL) (Miltenyi

Biotec), Bone Morphogenetic Protein 4 (BMP4) (R&D Systems), and 1.5 µM CHIR99021 (Axon

Medchem) was added. On day 3 the medium was refreshed with BPEL containing Matrigel (1:200) and

5 µM XAV 939 (1:1000). Further the medium was refreshed on day 6 and 9 of differentiation with fresh

BPEL without supplements. The cells were cultured until they start beating spontaneously, around day

13. By now they are ready for experiments and from now on called hESC derived cardiomyocytes

(hESC-CM). They are either immediately used or frozen for later usage. The differentiation and freezing

of these hESC-CM was not part of the project, but it is mentioned here for completeness.

2.1.2 Dissociation of hESC derived cardiomyocytes On the morning of day 13 medium was refreshed with 1mL of BPEL and simultaneously a 96-wells plate

was coated with Matrigel. In the afternoon the hESC-CM were washed with Dulbecco’s Phosphate-

Buffered Saline (DPBS) and dissociated using 1x TrypLE™ select solution (Gibco™) for 10 minutes at

37˚C. Enzymatic activity was neutralized using serum containing medium (MEF) at a ratio of 1:3. 2mL

MEF was added to a tube to which also the dissociated cells are added and 1mL MEF was used to flush

the well and take up the last hESC-CM using low adherence pipet tips. The cell suspension is

centrifuged at 1100rpm for 3 minutes. The supernatant was aspirated and the cells were resuspended

in 1mL BPEL and counted. After counting the cells were centrifuged again (1100rpm, 3min) and seeded

on the coated 96-wells plate at 120.000 cells/well.

2.1.3 Thawing hESC derived cardiomyocytes Before thawing of the hESC-CM a wells plate was incubated firstly with Vitronectin diluted in DPBS

(without calcium and magnesium) for 1 hour at room temperature (RT) and secondly with MEF

medium for 30 minutes at RT. The frozen vial was thawed in a water bath (37˚C), and 500µL BPEL was

added to it dropwise, to prevent osmotic shock. The cell suspension are transferred to a tube with 5mL

BPEL which is rinsed with another 1mL of BPEL. Centrifuge the cells, at 1100rpm for 3 minutes, and

aspirate the supernatant. Resuspend the cells in BPEL containing Revitacell (1:100) and count the cells.

Seed the cells in the desired density on the previously coated plate and refresh medium the next day

with fresh BPEL.

2.2 2D metabolism maturation On day 14 until day 16 medium was refreshed every day, adding 100µL of fresh medium per well of a

96-wells plate. On day 17 390µL BPEL was added to sustain the cells over the weekend. On day 20 the

medium was changed to CM medium with low insulin and galactose (GAL) and glucose (GLU) at

different concentrations. For the concentrations see Table 1. Glucose always had to be present since

previous experiments showed that the cells could not survive solely on galactose. On day 21 the

medium was taken up and prepared for the lactate assay by heat inactivating the Lactate

Dehydrogenase (LDH) for 30 minutes at 56 degrees. After taking up the medium, new medium is added

containing Rotenone (100nM) to block the Electron Transport Chain. At 24 hours of culture in the

medium containing rotenone the ATP assay was performed.

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2.2.1 Lactate Assay A colorimetric lactate assay kit (MAK064, Sigma-Aldrich®) was used. The accompanying protocol was

followed when preparing a standard and performing the assay. To start, the heat inactivated medium

was taken and 1µL per well was pipetted into a clear 96-wells plate containing 49µL of Lactate Assay

Buffer, to make 50µL per well. To this, 50µL of Master Reaction Mix was added before placing the

plate, protected from light with aluminum foil, on a horizontal shaker for 30 minutes. The colorimetric

detection was performed by measuring the absorbance at 570nm (Varioskan Lux, Thermo Fisher).

2.2.2 ATP assay For this assay a luminescent ATP detection kit (ab113849, Abcam) was used. The accompanying

protocol was followed when preparing a standard and performing the assay. This is and end point

assay since it starts by adding 50µL of detergent to every well and placing the plate on a shaker for 5

minutes. This is done to release and stabilize the ATP from the cells. Then 50µL of Substrate Solution

was added to every well before shaking again for 5 minutes. The plate is then dark adapted for 10

minutes, covered by aluminum foil, before measuring luminescence (Varioskan Lux, Thermo Fisher).

2.3 3D disease model

2.3.1 Computer aided design (CAD) With the use of CAD software (Solidworks) different designs were made of the mold for a microfluidic

chip. These designs all started with the basic design constraint of having two pillars inside a microfluidic

channel. The size and geometry of the channel and pillars were tailored to achieve ease of use and

suitable culture environment.

2.3.2 Micro-milling Using Solidworks a milling pattern was generated taking into consideration spindle speed, feed rate

and mill diameter to ensure a smooth surface finish. This is essential since every irregularity generated

in the mold in this production step can cause chip failure at a later stage. The failure can range from

difficulties visualizing the tissue to clogging of the microfluidics. A spindle speed of 20000 rpm is chosen

which was optimal for the milling machine that was used (DATRON Neo). The optimal feed rate differs

for every mill diameter. The bigger mills (4mm in diameter) were set to 600-800mm/min, while the

smaller mills (>1 mm) were set to 400mm/min. For the drilling a high rpm of 30000 was used. To

achieve a drilling depth of 1mm partial retract was used in steps of 100µm with a full retract after

500µm since the drill bits used (100µm) are so small and fragile. When all milling and drilling

parameters were optimized the design and milling pattern were uploaded to the DATRON Neo in which

the milling was performed. All molds were milled out of Poly(methyl methacrylate) (PMMA) since

Table 1: Carbon source concentrations

GAL (mM) GLU (mM) GLU (mM)

10 15 15

10 10 10

10 8 8

10 6 6

10 4 4

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PDMS does not stick to it and it is transparent which is preferential when degassing PDMS because all

bubbles are clearly visible.

2.3.3 Cleanroom wafer production Micro-milling could only be used until a certain size threshold. For the chip design with a square

channel of 200µm the production of a cleanroom wafer was necessary. With this method structures

from 400µm until nanoscale can be produced. To produce a wafer in the cleanroom first a mask has to

be produced. The design for this mask was made in Solidworks, converted to CleWin5 and then ordered

through the MESA+ Nanolab (Appendix).

The protocols for making one and two layer SU-8 structures on a silicon wafer can be found in the

appendix. Briefly, on a silicon wafer the SU-8 (photoresist) is deposited by spin coating. Subsequently

UV-exposure is performed through a photomask to selectively crosslink the SU-8. Next the SU-8 can

be developed and only the exposed structures remain. Multi-level structures of SU-8 can be fabricated

by repeating the steps of SU-8 deposition, photomask UV-exposure and SU-8 development.

To ensure low adhesion of PDMS to the silicon wafer and SU-8 structures and to make it more durable,

the wafers were coated with an anti-stick coating of perfluorodecyltrichlorosilane (FDTS). To apply

FDTS the wafer is placed in a vacuum and flushed with nitrogen for 20 minutes. Vacuum is reapplied

and for 20 minutes FDTS is released onto the wafer. After applying the FDTS another 5 minutes of

nitrogen flushing is performed before the wafer is taken out and ready for use as a mold.

2.3.4 Fabrication microfluidic chips For both the micro-milled mold and the cleanroom wafer the following step is similar. For the sake of

simplicity both will be called mold from now on.

Polydimethylsiloxane (PDMS) (1:10) (Sylgard 184 silicone elastomer kit, Dow Corning) is cast onto the

mold, which is designed to generate a microfluidic chip with a thickness of 3mm. This mold is then

placed in a desiccator for 45 minutes, releasing the vacuum once, to get rid of any trapped air. The

PDMS is cured in an oven at 60°C overnight, to ensure full crosslinking. When fully crosslinked the

PDMS chip can be peeled from its mold gently, to make sure the thin pillars do not break. The base of

the chip is made by spin coating a glass microscope slide (1500rpm, 60s, 1000 rpm·s-1, Spin150, Polos,

The Netherlands) and placing it is a 60°C oven for no less than 2 hours. To bond the PDMS chip onto

the coated glass slide plasma bonding is performed at 50W for 40s (Femto Science CUTE).

2.3.5 Preparation of a fibrin-based hydrogel To produce a fibrin-based hydrogel the following components were mixed in an Eppendorf placed on

ice: 20mg/mL fibrinogen in NaCl 0.9% (1:10 dilution v/v) (Sigma-Aldrich®), 100µL/mL Matrigel (1:10

dilution v/v), 250µg/mL Aprotenin (1:100 dilution v/v) (Sigma-Aldrich®) and two times concentrated

culture medium (1:10 dilution v/v). Lastly cell suspension is added (7:10 dilution v/v). For experiments

without cells normal culture medium is added instead of the cell suspension. This medium consisted

of CM medium with low insulin supplemented with insulin like growth factor-1 (IGF-1) and

triiodothyronine hormone (T3) at a concentration of 1:1000 and 1:300 respectively. As shortly before

seeding as possible 100U/mL thrombin (1:150 dilution v/v) is added to the mixture to start

polymerization.

2.3.6 Seeding microfluidic chips Before seeding of the microfluidic chip the volume per chip was calculated to estimate the amount of

chips that could be seeded in series so that the seeding can be done as quickly as possible. After

seeding of the chips, at a density of 20mln cells/mL, flushing of the seeding channel was performed

| 10

with either air or PBS. The chip is then left 10 min at RT to ensure polymerization. After this medium is

added in the form of medium filled pipet tips. 30µL filled pipet tips (P200) are inserted in one pair of

inlets while the other pair is supplied with 150µL, passive fluid leveling generates a small amount of

flow through the chip. Lastly the chips are placed in the incubator. Medium is refreshed every Monday,

Wednesday and Friday. Assessment of the cultured tissues was done using a Nikon eclipse Ts2.

2.3.7 Crosslinking tailoring To be able to measure the crosslinking time of fibrinogen, to form fibrin, the ability of a droplet of

hydrogel to attach to the wall of an Eppendorf was observed. A fibrin-based hydrogel was prepared to

which different concentrations of thrombin were added (100, 50, 30, 20 U/mL). 25µL of this hydrogel

was pipetted onto the inner wall of a 500µL Eppendorf. At consecutive time points (4, 8, 12, 16, 20 and

30 minutes) the Eppendorf was tilted upright to verify whether the droplet would drop to the bottom

or stick to the side. After this preliminary experiment on Eppendorfs, the experiment was repeated

inside the microfluidic chips. The microfluidic chips were filled with the hydrogel containing cells. The

filling of the chips was performed at 0, 5, 10, 15 and 20 minutes after thrombin addition, to verify

whether it was still possible to seed the already polymerizing hydrogel. 25 minutes after addition of

thrombin, medium was added to all chips to check whether the cells would remain in place, to confirm

appropriate polymerization of the fibrinogen.

2.3.8 Microfluidic surface treatment To keep cells from attaching to the surface of the microfluidic channel, chips were incubated with

Pluronic F127 (1% in water) for 20 minutes at RT. Consecutively the chips were rinsed thoroughly with

PBS twice, to remove any residual Pluronic F127. After rinsing, the chips were filled with PBS until use.

11 |

III. Results During this project a step has been tried towards metabolic maturation of cardiomyocytes, to

eventually be able to model myocardial infarction in vitro using hIPSC-CM.

3.1 2D Maturation Firstly, effort was put into maturing hESC-CMs in a monolayer. This was done by plating the hESC-CMs

in a 96 well plate and using cardiomyocyte medium supplemented with galactose to force the cells

into OXPHOS. To verify whether the hESC-CMs made the switch from glycolysis towards OXPHOS two

different assays were performed, a lactate and an ATP assay.

In normal culture medium around 15mM of glucose is present with most cultured mammalian cells

displaying aerobic glycolysis also known as the Warburg effect [61]. An inefficient energy metabolism,

characterized by increased glucose consumption and lactate production. However, just after birth, for

the first few hours of life, blood glucose levels can range between 1.4 mM and 6.2 mM and by about

72 hours after birth blood glucose levels reach normal infant, child and adult values (3.5-5.5 mM) [62].

For this reason a concentration range between 15 mM and more physiological 4 mM was chosen

supplemented with galactose (10mM). When CMs start using OXPHOS as means of energy production

they lower their lactate production. All pyruvate, resulting from glycolysis, now is converted into Acetyl

CoA instead of lactate. The lactate assay was performed to verify whether a certain concentration of

glucose combined with galactose will induce a shift from glycolysis towards OXPHOS.

Whenever a CM is using OXPHOS to produce energy the ETC is used to convert the energy of free

electrons to a proton gradient over the inner mitochondrial membrane. This gradient is used to power

the ATP-synthase thereby generating ATP (Fig. 2). Rotenone is a proven ETC inhibitor, which blocks the

CMs means of energy production through OXPHOS by inhibition of electron transport in complex I of

the ETC [63]. An ATP assay was performed to check the ATP content of the CMs. When CMs are using

OXPHOS and rotenone is added to the culture, they will only be able to produce limited amounts ATP.

However, when they were using glycolysis, rotenone will have no effect since glycolysis does not need

the ETC.

The lactate assay showed that the hESC-CM in monolayer were equally glycolytic across the different

conditions (Graph 1A). This means that varying the concentrations of glucose with or without galactose

results in a similar conversion rate of glucose into lactate. To corroborate that these cells are still

glycolytic and do not use OXPHOS, the results were compared with the ATP assay. The blocking of the

ETC with rotenone did resulted in different levels of ATP, as can been seen in Graph 1B. From the

results of the ATP assay can only be concluded that whenever the cells are supplied with a higher

amount of glucose they will produce more ATP.

This result could be due to the fact that the switch from OXPHOS back to glycolysis happens on a

smaller timescale than 24 hours. Rana et al. in their experiments on ATP production after carbon

source manipulation showed that differences in ATP production already occurred on a timescale of

minutes rather than hours [26]. Therefore another experiment was performed in which the time of

rotenone incubation was reduced from 24 hours to 10 minutes. This time was chosen since this was

the fasted all the steps of the experiment could be completed. Results of the ATP assay performed are

shown in Graph 2.

| 12

A

B

Graph 1 Monolayer hESC-CM maturation. Every data point is obtained by averaging a

technical duplicate. A| Results of the lactate assay after one day of culture in

CM medium with low insulin and galactose and glucose at varying

concentrations. All different concentrations yielded similar amounts of

lactate. B| The results of the ATP assay show that after 24 hours of

incubation with rotenone the cells that were cultured in lower amounts of

glucose produce less ATP. This holds for both with and without galactose. The

standard for the ATP in the assay kit did not show proper results. Therefore

no conversion of luminescence into ATP concentration could be done.

13 |

A

10min after rotenone addition With galactose

10min after rotenone addition Without galactose

B

Graph 2 ATP assay performed after 10 minutes of rotenone incubation on hESC-CM

cultured in medium with and without galactose. Every data point is

obtained by averaging a technical duplicate. A| After 10 minutes of rotenone

incubation the ATP assay did not show a trend in the results. B| The cells

cultured in galactose even showed higher ATP concentration for the lower

amounts of glucose. The standard for the ATP in the assay kit did not show

proper results. Therefore no conversion of luminescence into ATP

concentration could be done.

| 14

Culture of hESC-CM on medium supplemented with and without galactose showed very diverse results

after 10 minutes of incubation with rotenone. Cells cultured without galactose showed no clear trend

in the ATP contents, for the different glucose concentrations. It would be expected that the hESC-CM

would generate more ATP upon the addition of a higher concentration of glucose since there is more

substrate available for the energy metabolism. However, the lowest amount of ATP was observed in

the cells with the highest amount of glucose. The culture to which rotenone was added indeed showed

a high concentration of ATP upon addition of 15 mM of glucose. This is as expected since there is no

galactose present, to force the cells towards OXPHOS, so addition of rotenone will not affect the ATP

contents of the cells.

The cells cultured with galactose showed a similar trend for the cells incubated with and without

rotenone. However, this trend shows that cells cultured with lower amounts of glucose produce more

ATP even when rotenone was added. It would be expected that when the cells were using OXPHOS,

the amount of ATP would be equal but very low across all the concentrations. Taking into consideration

that the concentration of galactose is equal (10mM) and rotenone blocks its metabolism.

The overall results of the 2D maturation experiment did not show any conclusive results. The lactate

assay showed high amounts of lactate across all culture conditions, showing that all cells were equally

glycolytic. The ATP assay performed after 24 hours of rotenone incubation also showed glycolytic cells

because there was virtually no effect of rotenone. The only trend that was observed was that when

more glucose is added to a culture, cells will produce more ATP. The ATP assay performed shorter after

rotenone addition showed unexplainable results with the lowest amount of glucose producing the

highest amount of ATP.

15 |

3.2 3D disease modelling For over a hundred years, 2D cell cultures have been used as in vitro models to study cellular responses

to biochemical and biophysical cues [64]. These strategies are now well-accepted and have

considerably advanced our knowledge of cell behavior. However, growing evidence is now showing

that 3D cell cultures better recapitulate the in vivo microenvironment [29], [65], [66]. In vivo, cells are

surrounded by neighboring cells and encapsulated in ECM, which defines the nutrient availability,

biomechanics and signaling of this microenvironment. The supply of nutrients and removal of

metabolites behave differently in 3D cell culture as opposed to 2D. In a monolayer cells are always in

contact with the cell culture medium and therefore are able to take nutrients and oxygen from this

and excrete metabolites immediately into it. In 3D cell culture, cell that are on the inside of the tissue

depend on diffusion for receiving oxygen and nutrients and the removal of metabolites. This is vital

when modeling MI since the unavailability of oxygen and nutrients to the tissue is what causes the cell

damage [18], which can only be modelled whenever this availability is similar to the in vivo situation.

Not only is culturing cells in 3D beneficial for generating an accurate disease model, it could also

induce maturation of hIPSC-CM. Ulmer et al. performed a 3D hIPSC-CM culture using the EHT format

and studied the metabolic maturation. Their study provided evidence that a 3D culture format can

induce maturation of energy metabolism toward a more adult-like state [67]. They showed an

increased amount of mitochondria per CM, a 2.3-fold increase in ATP production by oxidation and

upregulation in genes involved in OXPHOS in cells cultured in 3D. This shows that, using

bioengineering to design a platform to produce a more in vivo-like environment could hold the key to

a more mature metabolism.

By incorporating this 3D culture of CM, the EHT, into a microfluidic device spatial and temporal control

over available nutrients and oxygen and over compound exposure can be achieved. It provides the

possibility to culture tissues under constant perfusion. Therefore the uptake of these nutrients, oxygen

and other compounds such as drugs, by the cells, does not create a gradient in the medium as

compared to static culture. Control over the amounts of nutrients and oxygen is vital to inducing MI

since this allows for precise control of its onset.

3.2.1 MicroEHT To be able to culture an EHT in a microfluidic channel flexible pillars are needed around which the EHT

can grow. This was taken as the first design constraint. Furthermore a possibility of seeding the CMs

around the pillars and a perfusion in- and outlet is needed which is the second design constraint. The

third design constraint was taken from Schaaf et al. [68] which is the aspect ratio of the EHT. This is a

3:1 length to width ratio.

Combining these design constraints, a CAD design for an EHT inside a square microfluidic channel was

made (Fig. 3.1) coined the microEHT. This design consisted of an EHT chamber of ratio 3:1 with two

pillars inside it. The space the pillar occupied is added in width of the channel, creating a bulge around

the pillars. This was done to allow enough cells to be seeded around the pillar for the formation of a

tissue around it. Attached to the EHT chamber are 2 in- and outlet pairs, 1 on both sides of the EHT

chamber. One inlet is used to fill the entire chip with CMs suspended in a hydrogel. Consecutively

through both outlets the cell-gel mixture that is not inside the EHT chamber it supposed to be flushed

out. Leaving cells only in the EHT chamber, with the possibility of adding flow through the in- and

outlets.

| 16

To be able to prototype the CAD design a mold was micro milled for casting of PDMS. This type of

prototyping was chosen since it was cheap and readily available. A mold was milled out of PMMA

containing different sizes of the designed microEHT ranging from 200 to 500µm channel width and

height. Onto this mold PDMS was casted and cured in an oven overnight. After curing, the PDMS was

peeled from the mold, cut into the individual microEHTs and plasma bonded onto a PDMS coated glass

slide.

To see whether it was possible to fill up the microEHT and consecutively empty the in- and outlet and

not unnecessarily use up cells, a gel only test was performed on these chips. The fibrin-based hydrogel

was pipetted into the chip after which emptying of the in- and outlet channels was attempted. The

chip could be filled with gel entirely and upon attempting to flush the in- and outlets one-sided flushing

was achieved (Fig. 3.2). However, flushing the second pair of in- and outlet never worked because all

the gel, including the EHT part, now flushed out. This was mainly due to there being nothing to keep

the gel in the EHT chamber in place.

Figure 3.2 Flushing of in- and outlet of the microEHT. A microEHT seeded with gel, containing

blue dye, after flushing with PBS (A) or aspiration (B). Showing the possibility of a

one-side clearing of the in- and outlet. Scale bars are 500µm

B A

Figure 3.1 The microEHT design. A simplified representation of the microEHT platform. 2 in-

and outlet pairs connected to the EHT chamber (red box). Cell seeding occurs

through one of the inlets (*), after which flushing is performed in the direction of

the blue arrows.

*

17 |

Since flushing of the gel from the in- and outlets proved to be the most difficult part, selective filling

of the microfluidic chip was explored. By making a cut-out right below and above the EHT chamber

(Fig. 3.3A) it was possible to pin the gel mixture and only have to flush one side. Also because of this

cut-out the resistance of the gel was high enough so that not the full EHT was aspirated. With

incorporation of these cut-outs, selective seeding of the EHT chamber was achieved in all sizes (200-

500µm) (Fig. 3.3).

3.2.2 MicroEHT array When designing a cell seeding protocol for the microEHT, it was noticed that the amount of cells that

comprise the actual EHT compared to the amount of cells that had to be flushed from the in- and

outlets was quite low. Whenever this platform would be used for disease modelling this is unwanted

because this means that a lot more cells are necessary than are actually used in the test, making it

more expensive than needed. For this reason, more EHTs have to be cultured in parallel to lower the

ratio of flushed cells. To achieve this the microEHT array was designed. An array of microEHTs,

connected at the top and bottom by two perfusion channels as can be seen in Figure 3.3.

Figure 3.3 Selective filling of the microEHT with cut-outs. A| The Solidworks model used to mill

the microEHT with cut-outs. B| A microEHT (200µm) with a selectively gel filled EHT

chamber. Scale bar is 1mm.

Figure 3.3 The microEHT array design. A sketch of the microEHT design with a channel diameter

of 200µm. Seeding occurs in the top channel in the direction of the arrow. After filling

of all 4 EHT chambers the cells in the top channel can be taken up again with the pipet

and seeded into the next array. Making the amount of wasted cells almost zero.

Bottom

Top

| 18

The cut-outs on both sides of the EHT chamber were also implemented in this design. The size of and

geometry of these cut-outs were determined by simply trying different sizes. When calculating the size

of these cut-outs values for hydrophobicity of the microfluidics and viscosity of the gel have to be taken

into account. Both of these values change over time. After plasma bonding the PDMS is hydrophilic

but will gradually become hydrophobic again over time. The viscosity of the gel changes because it is

polymerizing during the seeding process. Both these things make it very difficult to precisely calculate

the size of the cut-outs that is needed for selectively filling the microEHT array.

Cho et al. showed, that sharp cut-out angles are optimal for the selective filling of microfluidics by

pinning the gel at a certain point [69]. However producing a mold with these sharp angles cannot be

done through milling since this production technique does not allow for the generation of sharp angles

because of the round mills. This, and the fact that the designed microEHT array has a height and width

of only 200µm, means that the production of the mold for this chip has to done in the cleanroom.

The mold for the microEHT array was made using photolithography. A silicon wafer served as a base

onto which the photoresist (SU-8) was spin coated. Using a mask (Fig. 3.4A, Appendix) the SU-8 was

selectively crosslinked by UV exposure. Lastly the SU-8 was developed leaving only the exposed

structures. During production of the wafers it was soon realized that the usual method for developing

did not fully clean the uncured resist from the pillar holes. To get rid of this, the wafers were submerged

in an ultrasonic bath. This has to be done with considerable care so that SU-8 structures do not come

loose from the wafer.

3.2.3 MicroEHT array cell culture To verify the design, cardiomyocytes (of around 80% purity) were mixed into the fibrin-based hydrogel.

This cell-gel mixture was gently seeded into the microEHT array. This had to done gently to not break

the fluid pinning at end of the EHT chamber. The hydrophilicity of the chip plays a major role in the

success of the seeding procedure. When the chip is to hydrophilic, the mixture can easily surpass the

bottom cut-out and enter the bottom channel. However, when the chip is to hydrophobic the mixture

will not properly enter the EHT chamber.

The balance of hydrophilicity and hydrophobicity can be tailored by varying the time of the waiting

step between plasma bonding of the PDMS and the cell seeding. Experimental testing showed that 1

hour of waiting time resulted in optimal seeding conditions. This leaves enough time after bonding of

the chip to get the cell-gel mixture prepared before seeding. Figure 3.6A shows the result of the first

Figure 3.5 MicroEHT array. A| The mask that was used in cleanroom production of the

microEHT array mold. B| A selectively filled microEHT array with gel in the EHT

chamber and a still dry bottom channel. Scale bar 1mm

B

19 |

seeding in the microEHT array. These microEHTs were kept in culture for over two months (Fig. 3.6B)

proving that long term culture of EHTs in the microEHT array is possible.

The mold for these chips was made in one layer in the cleanroom, meaning that all features have the

same height. This causes the pillars and the channel to be of equal length. This consequently causes

the pillars to attach to the bottom of the channel during the bonding of the chip. Because of this the

EHTs are not able to bend the pillars, and no force readings can be performed.

To overcome the problem of the pillars bonding to the coated glass slide, two cleanroom masks were

made, one with and one without pillar holes. By first using the mask without pillar holes and

consecutively aligning the second one with pillars, the gap between the pillars and the bottom of the

channel could be tailored. A gap of 20µm was considered to be enough to not let the pillars bond, but

not enough for the microEHT to slip underneath the pillars. To corroborate whether a gap of 20µm

would provide enough space for the pillars not to attach to the bottom during bonding, 0.2µm

fluorescent beads (F8807, Invitrogen) were added to the microEHT array showing that they could go

underneath.

Figure 3.6 Seeding and culture inside the microEHT array. A|A freshly seeded microEHT array

(20 mln cells/mL), before medium addition. Seeding occurred through the top

channel, with the bottom channel staying nearly completely dry. After 10 minutes

of polymerization of fibrin, medium was added to both top and bottom channels.

B| A microEHT array after two months of culture. Showing the possibility of long

term culture. Scale bars are 400µm.

B

A Top channel (wet)

Bottom channel (dry)

| 20

3.2.4 Upscaling microEHT size The microEHT array with a channel diameter of 200µm have an EHT chamber volume of 0.03mm3.

Seeded at 20mln cells/mL yields microEHTs of around 600 cells. 600 cells is very little and some

experiments might require more cells. Therefore the possibility of upscaling the size of the microEHT

array was explored. Two different channel widths were tested (400µm and 1mm), with the same length

to width ratio of 3:1 (Fig. 3.7). The molds for these designs were made through micro milling since

these sizes fall outside of the possibilities of the cleanroom.

Through Solidworks the volume of the different size EHT chambers was calculated. From this a graph

was plotted showing the amount of cells necessary to generate EHTs in the different sizes of the

microEHT array. Values shown are within the tested boundaries and if seeded at 20mln cells/mL. This

was done so that whenever an experiment requires a certain amount of cells, the size of the EHT

chamber can easily be found and a tailor made microEHT array can be produced.

Figure 3.6 The fluorescent bead experiment. The white dots (fluorescent beads) shows that

there is enough space underneath the pillar (red circle) for it not to bond to the

bottom. Scale bar is 100µm.

A

B

Figure 3.7 Up scaled microEHT array. A|A microEHT array with a 400µm channel width after

seeding. Scale bar is 400 µm. B|. A microEHT array with a 1mm channel width after

seeding. The microEHT already shows the ability of bending the pillars. Scale bar is

1mm.

21 |

3.2.5 Fibrinogen crosslinking time When seeding cells into the microEHT array it has to be taken into account that during this time the

fibrinogen keeps crosslinking, because of the added thrombin. At a certain time point it will therefore

not be possible anymore to continue seeding, because the cell-gel mixture becomes to clumpy. To be

able to upscale the production of EHTs in the microEHT array it is necessary to be able to determine

the time it will take before the fibrinogen is crosslinked. This is what will determine how many

microEHT arrays than can be seeded consecutively.

With the current seeding protocol the fibrinogen is crosslinked in around eight minutes. With seeding

being possible until around 4 minutes after addition of thrombin. After this clumps of cells start

forming blocking the microfluidics. Within 4 minutes it is possible to seed around 12 microEHT arrays

(48 individual EHTs). However different microfluidic large-scale integration chips [70], [71] and a

microfluidic circuit board [72] are being developed which could upscale and automate EHT production.

Intuitively, when seeding a higher amount of EHTs, the seeding takes longer. To be able to perform the

chip seeding over a longer time the crosslinking time of the fibrinogen has to be extended.

There are different strategies to lengthen crosslinking time of fibrinogen, including changing the pH

and ionic strength of the fibrinogen mixture [50]. However, the easiest way is to lower the ratio of

thrombin to fibrinogen. Since there is less thrombin available to cleave the fibrinogen, fibrin strands

will form at a lower rate. To test the effect of different concentrations of thrombin on the crosslinking

time a simple experiment was performed. A droplet containing the fibrin-based hydrogel with different

concentrations of thrombin (100, 50, 30, 20 U/mL) was pipetted onto the inner side of an Eppendorf

laying on its side. At predetermined time points this Eppendorf was tilted upright. Whenever the

droplet slid to the bottom the hydrogel was considered as non-crosslinked. Table 2 shows the results

of this experiment. Each time a droplet fell to the bottom it was scored as a 1. Every time point and

condition was repeated 3 times. This showed that it is possible to extend the crosslinking time to

around 16 to 20 minutes.

To test whether these times hold in the microEHT array, cell seeding was performed using thrombin at

a concentration of 30 U/mL. The microEHT arrays were seeded after 0, 5, 10, 15 and 20 minutes after

Graph 3 Cells per EHT chamber. Whenever an experiment requires an EHT with a certain

amount of cells this graph can be used to determine the size of the EHT chamber.

| 22

addition of the thrombin to the cell-gel mixture. At 25 minutes after addition of the thrombin medium

was added to the chips to check whether gel had crosslinked and stayed inside the EHT chamber. This

showed that it is possible to seed polymerizing gel for up to 10 minutes after thrombin addition. At 15

minutes the gel had already polymerized too much. After addition of medium to the chips seeded at

0, 5 and 10 minutes all of the microEHTs stayed in the EHT chamber and could be kept in culture.

3.2.6 Fouling of the microEHT array When the microEHT array is kept in culture for a prolonged amount of time at a certain point, some of

the EHTs start attaching to the walls of the EHT chamber, especially the large EHTs (Fig. 3.8A). To

counteract this the chips were incubated, for 20 minutes, with Pluronic 127 to generate a non-fouling

surface. Pluronic has a triblock structure consisting of one hydrophobic polypropylene oxide (PPO)

group flanked by two hydrophilic polyethylene oxide (PEO) units (Fig. 3.8B) [73]. The hydrophobic PPO

chain is attached to a hydrophobic surface, and the hydrophilic PEO chains self-assemble into a cilia-

like surface that effectively repels proteins. Due to this no cells are able to attach. After incubation

with Pluronic the EHTs did not attach to the walls of the channel anymore as shown in Figure 3.8C.

Table 2: Fibrinogen polymerization

Time (min)

Thrombin concentration (U/mL)

100 50 30 20

4 3 3 3 3

8 1 2 3 3

12 1 1 3 3

16 0 0 2 3

20 0 0 0 3

30 0 0 0 3

C B A

Figure 3.8 Pluronic coating of PDMS. A|A microEHT attached to the bottom of the

channel after 2 days of culture. B| The structural formula of Pluronic and how

it is physisorbed onto a hydrophobic surface (PDMS). C| A microEHT 2 days

after seeding, showing no attachment to the channel because of the Pluronic

coating. Scale bars are 1mm.

23 |

IV. Discussion During this project, firstly and attempt was made to mature hESC-CM in a monolayer by switching the

carbon source away from glucose. Secondly, a microfluidic platform was developed which allows the

culture of an engineered heart tissue inside a microfluidic chip. This platform is made out of PDMS,

with an array of channels with flexible pillars. Cells were cultured in this platform for over two months.

In the upcoming section, the performed experimental procedures and obtained results will be

discussed and compared to literature.

4.1 2D maturation By addition of galactose to the culture medium no clear metabolic maturation of the hESC-CM was

obtained with Graph 1 showing that the cells still had a tendency towards glycolysis. Shortening the

timeframe of the ATP assay also did not yield the desired results. However, this could be due to the

fact that the metabolic switch does not occur simultaneously in all CMs. Therefore it could be the case

that part of the CM already switched towards glycolysis, while the other part is still using OXPHOS. If

the switching is occurring right around or within 10 minutes this could be an explanation for the erratic

results. To improve the measurements in following experiments a seahorse assay can be performed

(Seahorse XF Analyzers). The seahorse assay measures oxygen consumption rate (OCR) and

extracellular acidification rate (ECAR) of live cells in a multi-well plate. The OCR is an indicator of the

mitochondrial respiration and ECAR is largely due to glycolysis. Since this assay is performed on live

cells and rotenone addition can be done by the machine, measurements can be performed in real time

[26].

Another factor that could influence the results is the CM purity after differentiation. The purity of the

culture could influence the outcome of the experiment because you may be measuring the changes in

metabolism happening in other cells instead of the hESC-CM. Therefore it is recommended that

fluorescence-activated cell sorting (FACS) is performed to check the percentage of CM present after

differentiation.

Lastly as Veerman et al. stated in a review on hIPSC-CM immaturity, it possible that, since each of the

features of maturity may be independently regulated, strategies focusing on one particular aspect of

the maturation alone may be insufficient to improve cardiomyocyte maturity [74]. So focusing solely

on changing the metabolism may not be an effective way of maturing the cells. Combined approaches

that impact multiple parameters simultaneously could be more effective in achieving this.

4.2 3D disease modelling After establishment of a proof of concept for culturing an EHT in a microfluidic channel it should now

be corroborated whether this platform yields the capability of maturing the hESC-CM into a more

mature metabolism. By addition of galactose as a carbon source and culturing of the platform in

hypoxia it can be verified whether the cells are now dependent on oxygen for energy production.

Cells that use OXPHOS as a means of energy production will suffer cell damage or even cell death

upon removal of oxygen.

Even if maturation of hESC-CM is not achieved in the microEHT array it can still hold great value as a

platform for high-throughput analysis. Miniaturized EHTs have several advantages as they reduce cell

numbers, cell culture costs, manual handling requirements and are not restricted by oxygen

diffusion limitations, which is a problem with larger EHT constructs [47].

The currently cultured microEHTs were kept in culture for over two months. Over this two month

period the microEHTs had a survival rate of 50% (10 out of 20) after 1 month and only 20% (4 out of

20) after two months. Failure mostly happened in the form breakage of the EHTs. This breakage

| 24

could be due to the seeding density being too low or the percentage of CMs versus other cell types

being non-physiological. The cellular composition of the adult heart is widely debated topic with

estimations of CM percentages around 70-85% [75]–[77] with fibroblasts (FBs), endothelial cells

(ECs), and peri-vascular cells occupying the rest of the heart. Although these non-myocytes are a

relatively small fraction, they are essential for the adult heart, providing the ECM, intercellular

communication, and vascular supply. They will therefore play an important role in the structural

integrity of the microEHT. Cardiac fibroblasts are responsible for proper maintenance of the ECM

through a process of synthesis and remodeling [78]. They also contribute to the homogeneous

distribution of mechanical stress, which could be the key for survival of the microEHTs. Optimizing

the seeding density and percentages of cell types present in the culture is therefore essential in

obtaining and maintaining microEHTs.

Measuring the contractility of the microEHTs is a good functional read out. However, up until now this

has not been possible since the microEHTs either broke or were not able to bend the PDMS pillars. To

estimate the diameter of the pillar that a microEHT is able to bend, the spring constant of the PDMS

pillars can be calculated using the following formula [79]:

𝑘 =3𝜋𝐸𝐷4

64𝐿3

Here E represents the young’s modulus of PDMS (1.7MPa [80]), D is the diameter of the pillar and L

the length. To be able to calculate the pillar deflection (d), the force generated by the microEHT (F)

should be divided by the spring constant.

𝑑 =𝐹

𝑘

However estimating the force that will be generated by a microEHT is very difficult. Therefore it is

recommended to make microEHT arrays with different pillar diameters to measure the deflection a

microEHT can generate and from here calculate the force. This force calculation can be used to then

tailor the diameter of the PDMS pillar.

4.3 Fibrinogen crosslinking time The crosslinking time of fibrinogen was doubled from 8 to 16 minutes by adding 30 instead of 100

U/mL of thrombin. A problem that arises when the cell-gel mixture is left to crosslink in the EHT

chamber for a long time is that it starts drying out. When a closed circuit is used like the, microfluidic

large-scale integration chips [70], [71] and a microfluidic circuit board [72], this will not occur since

the liquid is sealed and cannot evaporate. A solution for this problem in the microEHT array could be

that after seeding the in- and outlets are sealed by placing the chip underneath a PDMS slap, blocking

the possibility of evaporation.

4.4 Fouling of the microEHT array It is important to note that, while Pluronic is established as a non-fouling agent, it degrades in a cell-

independent manner [81], so it does have a finite lifespan. By incubating for 20 minutes, the Pluronic

is physisorbed onto the PDMS, but the presence of serum proteins in the culture medium will

eventually replace the polymer from the surface [82]. This replacement takes place within two days

of coating. After this short time the microEHTs did still not show signs of attaching which may be due

to the fact that they are attached to the pillars and therefore not touching the surface. To be sure

that the coating with Pluronic lasts longer it can be incorporated into the PDMS. Embedding Pluronic

in PDMS allows gradient induced migration to the surface of the microfluidic channel. Hereby the

surface could be modified to become non-fouling for over 10 days [83].

25 |

V. Conclusion In this project unfortunately no measurable metabolic maturation of hESC-CM was achieved. After

adding galactose to the culture medium cells still showed a glycolytic metabolism. This could be due

to the cells not maturing or the currently used assay not being capable of measuring the actual switch

between oxidative phosphorylation and glycolysis.

Furthermore a microfluidic device was designed with the capabilities of culturing an engineered heart

tissue inside. This was achieved by adding pillars in a microfluidic chamber which could be selectively

filled with cardiomyocytes. Compared to the current state of the art, the microEHT array benefits from

the general advantages of microfluidic devices, such as the more representative 3D microenvironment

for the cells, control over compound exposure and the small size. This 3D microenvironment has

previously been shown to help mature the cardiomyocytes and with the control over compound

exposure concentrations of carbon sources can be kept constant and hopefully this will help in

metabolically maturing the cardiomyocytes.

Even if the cardiomyocytes cannot be matured inside the microEHT array, the platform still holds great

value. Having a miniaturized heart tissue with contractility as a functional read-out can be

implemented as a high-throughput analysis platform slowly shifting away from animal testing.

VI. Future perspective Firstly it has to be tested whether the culture of cardiomyocytes on the microEHT array improves

maturation and allows for the culturing of hIPSC-CM with a more mature metabolism. This could be

tested by addition of galactose as a carbon source and culturing of the platform in hypoxia. Cells that

use OXPHOS as a means of energy production will suffer cell damage or even cell death upon removal

of oxygen. This can consecutively be measured by performing a live death staining.

Whenever this is achieved the next step would be to check whether MI can be induced by culturing

the mature hESC-CM in a hypoxia chamber. When successful the platform can be used the perform

studies on the mechanisms behind MI and the possibility of prevention or drug intervention. However,

for the implementation of the platform in drug testing the device cannot be made out of PDMS

anymore because it can absorb small molecules such as drugs [84]. Therefore research will have to be

put into finding an alternative. An option for this could be the photosensitive thermoset OSTEMER

322-40 which has shown to have good optical properties and biocompatibility with low vapor

permeability [85].

To be able to do high throughput analysis the amount of microEHTs per device should be increased.

This can be done by incorporating the design of microEHT array into a microfluidic circuit board. A

small step has already been taken towards this by lengthening the time in which seeding can occur.

| 26

Acknowledgements

First of all I would like to thank Prof. Passier for giving me the opportunity of performing my graduate

project at the applied stem cell technologies group. I just came walking into the group in search of a

project and after a fairly short conversation it was decided, I could start the next day. I felt immediately

welcomed and at home. For my daily supervision I thank Marcelo Ribeiro. His close supervision was

very helpful and whenever I was sat staring at a problem for too long he was always prepared to listen.

The project he provided me with was challenging and a lot of fun. Through his knowledge I learned a

lot, not only about the cardiac field but also about hard work. Apart from the daily supervision I thank

Wesley, my daily sparring partner. Whenever a problem arose that needed some engineering, Wesley

was there to think along with me and off course help me mill the idea afterwards. Off course also a big

thank you to the rest of AST, specifically room ZH115. Who were always there whenever a cup of coffee

and a good conversation was exactly what I needed.

For production of all the devices I designed I would like to thank Johan Bomer, Rolf Slaats and Jose

Rivera Arbelaez.

To all the friends I met during my time studying in Enschede, thank you for providing me with the

sometimes necessary and sometimes abundant times of distraction from studying. This is what kept

me going and made sure I reached the finish line.

The time has come to express my utmost gratitude to my family, who always supported blindly. I know

it was not always easy raising me, but you made me the man I am today. You always listened, with

great interest, to the stories I brought home from the far away Enschede. I hope I have made you all

proud.

Lastly I would like to thank the loveliest girl I met during my years of studying. Milou thank you for

showing me how big the world is and what can happen if you look further than the railroad between

Enschede and Roermond.

27 |

Bibliography

[1] “Cardiovascular diseases (CVDs).” [Online]. Available: https://www.who.int/news-room/fact-sheets/detail/cardiovascular-diseases-(cvds). [Accessed: 30-Sep-2019].

[2] C. D. Mathers and D. Loncar, “Projections of global mortality and burden of disease from 2002 to 2030,” PLoS Med., vol. 3, no. 11, pp. 2011–2030, Nov. 2006.

[3] T. Thom et al., “Heart disease and stroke statistics - 2006 Update: A report from the American Heart Association Statistics Committee and Stroke Statistics Subcommittee,” Circulation, vol. 113, no. 6. Feb-2006.

[4] G. Minotti, Cardiotoxicity of Non-Cardiovascular Drugs. John Wiley and Sons, 2010.

[5] J. P. Piccini et al., “Current challenges in the evaluation of cardiac safety during drug development: Translational medicine meets the Critical Path Initiative,” American Heart Journal, vol. 158, no. 3. pp. 317–326, Sep-2009.

[6] N. Milani-Nejad and P. M. L. Janssen, “Small and large animal models in cardiac contraction research: Advantages and disadvantages,” Pharmacology and Therapeutics, vol. 141, no. 3. Elsevier Inc., pp. 235–249, 01-Mar-2014.

[7] K. Takahashi and S. Yamanaka, “Induction of Pluripotent Stem Cells from Mouse Embryonic and Adult Fibroblast Cultures by Defined Factors,” Cell, vol. 126, no. 4, pp. 663–676, Aug. 2006.

[8] P. W. Burridge et al., “A universal system for highly efficient cardiac differentiation of human induced pluripotent stem cells that eliminates interline variability,” PLoS One, vol. 6, no. 4, 2011.

[9] X. Lian et al., “Directed cardiomyocyte differentiation from human pluripotent stem cells by modulating Wnt/β-catenin signaling under fully defined conditions,” Nat. Protoc., vol. 8, no. 1, pp. 162–175, Jan. 2013.

[10] H. S. V. Chen, C. Kim, and M. Mercola, “Electrophysiological challenges of cell-based myocardial repair,” Circulation, vol. 120, no. 24. pp. 2496–2508, Dec-2009.

[11] W. C. Stanley, F. A. Recchia, and G. D. Lopaschuk, “Myocardial substrate metabolism in the normal and failing heart.,” Physiol. Rev., vol. 85, no. 3, pp. 1093–129, Jul. 2005.

[12] G. D. Lopaschuk, R. L. Collins-Nakai, and T. Itoi, “Developmental changes in energy substrate use by the heart,” Cardiovascular Research, vol. 26, no. 12. Oxford University Press, pp. 1172–1180, 1992.

[13] Y. M. Cho et al., “Dynamic changes in mitochondrial biogenesis and antioxidant enzymes during the spontaneous differentiation of human embryonic stem cells,” Biochem. Biophys. Res. Commun., vol. 348, no. 4, pp. 1472–1478, Oct. 2006.

[14] G. D. Lopaschuk, M. A. Spafford, and D. R. Marsh, “Glycolysis is predominant source of myocardial ATP production immediately after birth.,” Am. J. Physiol., vol. 261, no. 6 Pt 2, pp. H1698-705, Dec. 1991.

[15] S. Chung, P. P. Dzeja, R. S. Faustino, C. Perez-Terzic, A. Behfar, and A. Terzic, “Mitochondrial oxidative metabolism is required for the cardiac differentiation of stem cells.,” Nat. Clin. Pract. Cardiovasc. Med., vol. 4 Suppl 1, no. Suppl 1, pp. S60-7, Feb. 2007.

[16] “Oxidative phosphorylation | Biology (article) | Khan Academy.” [Online]. Available: https://www.khanacademy.org/science/biology/cellular-respiration-and-fermentation/oxidative-phosphorylation/a/oxidative-phosphorylation-etc. [Accessed: 14-Oct-2019].

[17] J. M. Berg, J. L. Tymoczko, and L. Stryer, “Oxidative Phosphorylation,” 2002.

[18] “Heart Attack (Myocardial Infarction) - Harvard Health.” [Online]. Available: https://www.health.harvard.edu/a_to_z/heart-attack-myocardial-infarction-a-to-z. [Accessed: 14-Oct-2019].

| 28

[19] H. S. Gillespie, C. C. H. Lin, and J. M. Prutkin, “Arrhythmias in Structural Heart Disease,” Current Cardiology Reports, vol. 16, no. 8. Current Medicine Group LLC 1, 2014.

[20] M. S. Martínez et al., “Energetic metabolism in cardiomyocytes: molecular basis of heart ischemia and arrhythmogenesis,” Vessel Plus, vol. 1, no. 12, Dec. 2017.

[21] T. Boudou et al., “A Microfabricated Platform to Measure and Manipulate the Mechanics of Engineered Cardiac Microtissues,” Tissue Eng. Part A, vol. 18, no. 9–10, pp. 910–919, May 2012.

[22] H. T. Heidi Au, B. Cui, Z. E. Chu, T. Veres, and M. Radisic, “Cell culture chips for simultaneous application of topographical and electrical cues enhance phenotype of cardiomyocytes,” Lab Chip, vol. 9, no. 4, pp. 564–575, 2009.

[23] N. L. Tulloch et al., “Growth of engineered human myocardium with mechanical loading and vascular coculture,” Circ. Res., vol. 109, no. 1, pp. 47–59, Jun. 2011.

[24] A. Mihic et al., “The effect of cyclic stretch on maturation and 3D tissue formation of human embryonic stem cell-derived cardiomyocytes,” Biomaterials, vol. 35, no. 9, pp. 2798–2808, Mar. 2014.

[25] M. C. Ribeiro et al., “Functional maturation of human pluripotent stem cell derived cardiomyocytes in vitro – Correlation between contraction force and electrophysiology,” Biomaterials, vol. 51, pp. 138–150, May 2015.

[26] P. Rana, B. Anson, S. Engle, and Y. Will, “Characterization of Human-Induced Pluripotent Stem Cell–Derived Cardiomyocytes: Bioenergetics and Utilization in Safety Screening,” Toxicol. Sci., vol. 130, no. 1, pp. 117–131, Nov. 2012.

[27] C. C. Veerman, G. Kosmidis, C. L. Mummery, S. Casini, A. O. Verkerk, and M. Bellin, “Immaturity of human stem-cell-derived cardiomyocytes in culture: fatal flaw or soluble problem?,” Stem Cells Dev., vol. 24, no. 9, pp. 1035–52, May 2015.

[28] T. Kamakura et al., “Ultrastructural maturation of human-induced pluripotent stem cell-derived cardiomyocytes in a long-term culture.,” Circ. J., vol. 77, no. 5, pp. 1307–14, 2013.

[29] T. Eschenhagen et al., “Three-dimensional reconstitution of embryonic cardiomyocytes in a collagen matrix: a new heart muscle model system.,” FASEB J., vol. 11, no. 8, pp. 683–694, Jul. 1997.

[30] T. Shimizu et al., “Fabrication of pulsatile cardiac tissue grafts using a novel 3-dimensional cell sheet manipulation technique and temperature-responsive cell culture surfaces.,” Circ. Res., vol. 90, no. 3, 2002.

[31] Y. S. Zhang et al., “Bioprinting 3D microfibrous scaffolds for engineering endothelialized myocardium and heart-on-a-chip,” Biomaterials, vol. 110, pp. 45–59, Dec. 2016.

[32] W. H. Zimmermann et al., “Tissue engineering of a differentiated cardiac muscle construct,” Circ. Res., vol. 90, no. 2, pp. 223–230, Feb. 2002.

[33] C. Correia et al., “Distinct carbon sources affect structural and functional maturation of cardiomyocytes derived from human pluripotent stem cells,” Sci. Rep., vol. 7, no. 1, Dec. 2017.

[34] C. Aguer et al., “Galactose enhances oxidative metabolism and reveals mitochondrial dysfunction in human primary muscle cells,” PLoS One, vol. 6, no. 12, Dec. 2011.

[35] M. N. Hirt, A. Hansen, and T. Eschenhagen, “Cardiac tissue engineering : State of the art,” Circulation Research, vol. 114, no. 2. Lippincott Williams and Wilkins, pp. 354–367, 2014.

[36] H. C. Ott et al., “Perfusion-decellularized matrix: Using nature’s platform to engineer a bioartificial heart,” Nat. Med., vol. 14, no. 2, pp. 213–221, Feb. 2008.

[37] A. Khademhosseini et al., “Microfluidic patterning for fabrication of contractile cardiac organoids,” Biomed. Microdevices, vol. 9, no. 2, pp. 149–157, Apr. 2007.

[38] A. Agarwal, J. A. Goss, A. Cho, M. L. McCain, and K. K. Parker, “Microfluidic heart on a chip for higher

29 |

throughput pharmacological studies,” Lab Chip, vol. 13, no. 18, pp. 3599–3608, Sep. 2013.

[39] J. Jang et al., “3D printed complex tissue construct using stem cell-laden decellularized extracellular matrix bioinks for cardiac repair,” Biomaterials, vol. 112, pp. 264–274, Jan. 2017.

[40] H. Naito et al., “Optimizing engineered heart tissue for therapeutic applications as surrogate heart muscle,” Circulation, vol. 114, no. SUPPL. 1, Jul. 2006.

[41] M. Tiburcy et al., “Defined engineered human myocardium with advanced maturation for applications in heart failure modeling and repair,” Circulation, vol. 135, no. 19, pp. 1832–1847, May 2017.

[42] T. Shimizu, M. Yamato, Y. Isoi, T. Akutsu, and T. Setomaru, “Fabrication of Pulsatile Cardiac Tissue Grafts Using a Novel 3-Dimensional Cell Sheet Manipulation Technique and,” Circ. Res., pp. 1–9, 2002.

[43] J. P. Guyette et al., “Bioengineering Human Myocardium on Native Extracellular Matrix,” Circ. Res., vol. 118, no. 1, pp. 56–72, Jan. 2016.

[44] A. V. Borovjagin, B. M. Ogle, J. L. Berry, and J. Zhang, “From Microscale Devices to 3D Printing: Advances in Fabrication of 3D Cardiovascular Tissues,” Circulation Research, vol. 120, no. 1. Lippincott Williams and Wilkins, pp. 150–165, 06-Jan-2017.

[45] L. A. Hockaday et al., “Rapid 3D printing of anatomically accurate and mechanically heterogeneous aortic valve hydrogel scaffolds,” Biofabrication, vol. 4, no. 3, Sep. 2012.

[46] R. J. Mills and J. E. Hudson, “Bioengineering adult human heart tissue: How close are we?,” APL Bioeng., vol. 3, no. 1, p. 010901, Mar. 2019.

[47] R. J. Mills, H. K. Voges, E. R. Porrello, and J. E. Hudson, “Disease modeling and functional screening using engineered heart tissue,” Current Opinion in Physiology, vol. 1. Elsevier Ltd, pp. 80–88, 01-Feb-2018.

[48] A. Hansen et al., “Development of a drug screening platform based on engineered heart tissue.,” Circ. Res., vol. 107, no. 1, pp. 35–44, Jul. 2010.

[49] K. Y. Ye, K. E. Sullivan, and L. D. Black, “Encapsulation of cardiomyocytes in a fibrin hydrogel for cardiac tissue engineering,” J. Vis. Exp., no. 55, 2011.

[50] H. A. Scheraga, “The thrombin-fibrinogen interaction,” Biophys. Chem., vol. 112, no. 2-3 SPEC. ISS., pp. 117–130, Dec. 2004.

[51] K. Breckwoldt et al., “Differentiation of cardiomyocytes and generation of human engineered heart tissue,” Nat. Protoc., vol. 12, no. 6, pp. 1177–1197, Jun. 2017.

[52] C. Han, J. M. Labuz, S. Takayama, and J. Park, “Organs-on-a-Chip,” in Tissue Engineering: Second Edition, Elsevier Inc., 2014, pp. 717–746.

[53] A. Polini, L. Prodanov, N. S. Bhise, V. Manoharan, M. R. Dokmeci, and A. Khademhosseini, “Organs-on-a-chip: A new tool for drug discovery,” Expert Opinion on Drug Discovery, vol. 9, no. 4. Informa Healthcare, pp. 335–352, 2014.

[54] I. K. Zervantonakis, C. R. Kothapalli, S. Chung, R. Sudo, and R. D. Kamm, “Microfluidic devices for studying heterotypic cell-cell interactions and tissue specimen cultures under controlled microenvironments.,” Biomicrofluidics, vol. 5, no. 1, p. 13406, Mar. 2011.

[55] G. M. Whitesides, “The origins and the future of microfluidics,” Nature, vol. 442, no. 7101. pp. 368–373, 27-Jul-2006.

[56] D. J. Guckenberger, T. E. de Groot, A. M. D. Wan, D. J. Beebe, and E. W. K. Young, “Micromilling: a method for ultra-rapid prototyping of plastic microfluidic devices.,” Lab Chip, vol. 15, no. 11, pp. 2364–78, Jun. 2015.

[57] J. Z. Zhang and J. C. Chen, “The development of an in-process surface roughness adaptive control system in end milling operations,” Int. J. Adv. Manuf. Technol., vol. 31, no. 9–10, pp. 877–887, Jan.

| 30

2007.

[58] V. Faustino, S. O. Catarino, R. Lima, and G. Minas, “Biomedical microfluidic devices by using low-cost fabrication techniques: A review,” J. Biomech., vol. 49, no. 11, pp. 2280–2292, Jul. 2016.

[59] H. Hillborg, J. F. Ankner, U. W. Gedde, G. D. Smith, H. K. Yasuda, and K. Wikström, “Crosslinked polydimethylsiloxane exposed to oxygen plasma studied by neutron reflectometry and other surface specific techniques,” Polymer (Guildf)., vol. 41, no. 18, pp. 6851–6863, 2000.

[60] S. H. Tan, N. T. Nguyen, Y. C. Chua, and T. G. Kang, “Oxygen plasma treatment for reducing hydrophobicity of a sealed polydimethylsiloxane microchannel,” Biomicrofluidics, vol. 4, no. 3, 2010.

[61] O. Warburg, F. Wind, and E. Negelein, “The metabolism of tumors in the body,” J. Gen. Physiol., vol. 8, no. 6, pp. 519–530, Mar. 1927.

[62] M. Güemes, S. A. Rahman, and K. Hussain, “What is a normal blood glucose?,” Arch. Dis. Child., vol. 101, no. 6, pp. 569–574, 2016.

[63] S. Heinz, A. Freyberger, B. Lawrenz, L. Schladt, G. Schmuck, and H. Ellinger-Ziegelbauer, “Mechanistic Investigations of the Mitochondrial Complex I Inhibitor Rotenone in the Context of Pharmacological and Safety Evaluation.,” Sci. Rep., vol. 7, p. 45465, 2017.

[64] R. G. Harrison, M. J. Greenman, F. P. Mall, and C. M. Jackson, “Observations of the living developing nerve fiber,” Anat. Rec., vol. 1, no. 5, pp. 116–128, 1907.

[65] L. G. Griffith and M. A. Swartz, “Capturing complex 3D tissue physiology in vitro,” Nature Reviews Molecular Cell Biology, vol. 7, no. 3. pp. 211–224, Mar-2006.

[66] K. Duval et al., “Modeling physiological events in 2D vs. 3D cell culture,” Physiology, vol. 32, no. 4. American Physiological Society, pp. 266–277, 14-Jun-2017.

[67] B. M. Ulmer et al., “Contractile Work Contributes to Maturation of Energy Metabolism in hiPSC-Derived Cardiomyocytes.,” Stem cell reports, vol. 10, no. 3, pp. 834–847, 2018.

[68] S. Schaaf, A. Eder, I. Vollert, A. Stöhr, A. Hansen, and T. Eschenhagen, “Generation of strip-format fibrin-based engineered heart tissue (EHT),” Methods Mol. Biol., vol. 1181, pp. 121–129, 2014.

[69] H. Cho, H. Y. Kim, J. Y. Kang, and T. S. Kim, “How the capillary burst microvalve works,” J. Colloid Interface Sci., vol. 306, no. 2, pp. 379–385, Feb. 2007.

[70] R. Gómez-Sjöberg, A. A. Leyrat, D. M. Pirone, C. S. Chen, and S. R. Quake, “Versatile, fully automated, microfluidic cell culture system,” Anal. Chem., vol. 79, no. 22, pp. 8557–8563, Nov. 2007.

[71] M. Blazek, S. Santisteban, and M. Meier, “Lab on a Chip single cells on a microfluidic chip †,” Lab Chip, vol. 15, pp. 726–734, 2015.

[72] A. R. Vollertsen et al., “Standardized, modular parallelization platform for microfluidic large-scale integration cell culturing chips,” 22nd Int. Conf. Miniaturized Syst. Chem. Life Sci. MicroTAS, pp. 2199–2201, 2018.

[73] P. Alexandridis and T. Alan Hatton, “Poly(ethylene oxide)poly(propylene oxide)poly(ethylene oxide) block copolymer surfactants in aqueous solutions and at interfaces: thermodynamics, structure, dynamics, and modeling,” Colloids and Surfaces A: Physicochemical and Engineering Aspects, vol. 96, no. 1–2. pp. 1–46, 10-Mar-1995.

[74] C. C. Veerman, G. Kosmidis, C. L. Mummery, S. Casini, A. O. Verkerk, and M. Bellin, “Immaturity of Human Stem-Cell-Derived Cardiomyocytes in Culture: Fatal Flaw or Soluble Problem?,” Stem Cells and Development, vol. 24, no. 9. Mary Ann Liebert Inc., pp. 1035–1052, 01-May-2015.

[75] G. R. Guo, L. Chen, M. Rao, K. Chen, J. P. Song, and S. S. Hu, “A modified method for isolation of human cardiomyocytes to model cardiac diseases,” J. Transl. Med., vol. 16, no. 1, Oct. 2018.

[76] Y. Tang, J. R. Nyengaard, J. B. Andersen, U. Baandrup, and H. J. G. Gundersen, “The application of

31 |

stereological methods for estimating structural parameters in the human heart.,” Anat. Rec. (Hoboken)., vol. 292, no. 10, pp. 1630–47, Oct. 2009.

[77] P. Zhou and W. T. Pu, “Recounting Cardiac Cellular Composition,” Circ. Res., vol. 118, no. 3, pp. 368–370, Feb. 2016.

[78] S. Psarras, D. Beis, S. Nikouli, M. Tsikitis, and Y. Capetanaki, “Three in a Box: Understanding Cardiomyocyte, Fibroblast, and Innate Immune Cell Interactions to Orchestrate Cardiac Repair Processes,” Front. Cardiovasc. Med., vol. 6, Apr. 2019.

[79] N. E. Oyunbaatar, D. H. Lee, S. J. Patil, E. S. Kim, and D. W. Lee, “Biomechanical characterization of cardiomyocyte using PDMS pillar with microgrooves,” Sensors (Switzerland), vol. 16, no. 8, Aug. 2016.

[80] I. D. Johnston, D. K. McCluskey, C. K. L. Tan, and M. C. Tracey, “Mechanical characterization of bulk Sylgard 184 for microfluidics and microengineering,” J. Micromechanics Microengineering, vol. 24, no. 3, 2014.

[81] C. M. Nelson, S. Raghavan, J. L. Tan, and C. S. Chen, “Degradation of micropatterned surfaces by cell-dependent and -independent processes,” Langmuir, vol. 19, no. 5, pp. 1493–1499, Mar. 2003.

[82] J. T. Li and K. D. Caldwell, “Plasma protein interactions with Pluronic®-treated colloids,” Colloids Surfaces B Biointerfaces, vol. 7, no. 1–2, pp. 9–22, Jul. 1996.

[83] W. Qiu, X. Sun, C. Wu, K. Hjort, and Z. Wu, “A Contact angle study of the interaction between embedded amphiphilic molecules and the PDMS matrix in an aqueous environment,” Micromachines, vol. 5, no. 3, pp. 515–527, 2014.

[84] B. J. van Meer et al., “Small molecule absorption by PDMS in the context of drug response bioassays,” Biochem. Biophys. Res. Commun., vol. 482, no. 2, pp. 323–328, Jan. 2017.

[85] D. Sticker, M. Rothbauer, S. Lechner, M. T. Hehenberger, and P. Ertl, “Multi-layered, membrane-integrated microfluidics based on replica molding of a thiol-ene epoxy thermoset for organ-on-a-chip applications,” Lab Chip, vol. 15, no. 24, pp. 4542–4554, 2015.

Appendix: UV-masks and cleanroom protocol for fabrication of

the microEHT array

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Name of process flow: 2-step SU-8 lithographtPlatform: FluidicsCreation date: 2017-01-24Personal informationUser name: Slaats, R.H.Email address: r.h.slaats@utwente.nlCompany/Chair: ASTFunction: PhdProject: I/R-on-a-ChipName of supervisor: Robert PassierProcess planningProcess start: Process end: StatusName of advisor: Last revision: 2019-08-07Approval: Approval date: Expiration date:

ILP: In-line Processing MFP: Metal-freeProcessing

UCP: Ultra CleanProcessing Removal of Residues

Step Level Process/Basic flow Usercomments

1 Substrate Silicon (#subs107)

NL-CLR-Wafer Storage CupboardOrientation: <100>Diameter: 100mmThickness: 381µm ± 15Polished: Single side polished (OSP)Resistivity: 1-10 ΩcmType: N/phosphor

litho1830: Lithography SU-8 5 (negative resist - ILP) 2 ILP Dehydration bake for SU-

8 (#lith050)

NL-CLR-WB24 Dehydration bake on hotplate• Temperature: 120°C• Time: 10min

3 ILP Coating SU-8 5 (#lith163)

NL-CLR-WB24 SüssMicroTec Spinner Delta 20Resist: MicroChem NANOTM SU-8 5

• Static dispense of resist from the bottle to cover of the substrate• Select one of the dynamic spin programs (see table below): - ramp of 100rpm/sec to 500rpm for 10-15s to cover the entiresurface of the substrate with resist - ramp of 300rpm/sec to final spin speed and hold for 30s

SU-8 5 spin speed characteristics:

Print date: 2019-10-15

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Spin Program Spin Speed(rpm)

Thickness(μm)

1 1000 13.62 1500 8.43 2000 7.44 2500 6.05 3000 5.26 3500 4.67 4000 4.1

4 ILP Softbake SU-8 5 (#lith164)

NL-CLR-WB24 • Hotplate• For spin programs 4-7• start @ 25 oC• 1 min @ 50 oC• 1 min @ 65 oC• 3 min @ 95 oC• 5°C/2min ramp down to 25°C

5 ILP Alignment &Exposure SU-8 5(#lith165)

NL-CLR-EV620 mask alignersUse contact mode

Spinprogram

Exposuretime [s]

1 132 133 134 105 106 107 10

6 ILP Post exposure bake SU-85(#lith166)

NL-CLR-WB24• Hotplate• For spin program 4-7:• Start @ 25 °C• 1 min @ 50 °C• 1 min @ 65 °C• 2 min @ 80 °C• 5°C/2min down to 25°C

7 ILP Development SU-8 5(#lith167)

NL-CLR-WB 24 • TCO Spray Developer• Developer: PGMEA (RER600, ARCH Chemicals)

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• Time 3:30 min with spray-gun• Rinse with RER600• Rinse with IPA• Spin dry• Check result and perform extra cycles if not complete

8 ILP Hard bake SU-8 (#lith168)

NL-CLR-WB24• Hotplate• 2hr @ 120 °C

litho1832: Lithography SU-8 100 (negative resist - ILP) 9 ILP Dehydration bake for SU-

8 (#lith050)

NL-CLR-WB24 Dehydration bake on hotplate• Temperature: 120°C• Time: 10min

10 ILP Coating SU-8 100(Delta20)(#lith125)

NL-CLR-WB24 SüssMicroTec Spinner Delta 20Resist: MicroChem NANOTM SU-8 100

SU-8 100 spin speed characteristics:

Spin Program Spin Speed(rpm)

Thickness(μm)

8 1000 3809 1500 23010 2000 20011 2500 15012 3000 12013 3500 10014 4000 85

11 ILP Prebake SU-8 100(#lith126)

NL-CLR-WB24Hotplate xxFor spin programs 8 - 10:• 25°C• 10min @ 50°C• 30min @ 65°C• 210min @ 95°C• 5°C/5min ramp down to 25°C

For spin programs 11- 14:• 25°C• 10min @ 50°C• 30min @ 65°C• 120min @ 95°C• 5°C/5min ramp down to 25°C

12 ILP Alignment & ExposureSU-8 100 (#lith127)

NL-CLR-EV620 mask alignersUse contact mode

Spinprogram

Exposuretime [s]

8 1209 7510 6711 6012 4513 3714 37

15-10-2019 https://mesaplusnanolab.ewi.utwente.nl/MICMEC/tts/proc_show.php

https://mesaplusnanolab.ewi.utwente.nl/MICMEC/tts/proc_show.php 4/4

13 ILP Post exposure bake SU-8100(#lith128)

NL-CLR-WB24/hotplatesFor spin programs 8 - 10:• 25°C• 10min @ 50°C• 10min @ 65°C• 50min @ 75°C• 5°C/5min ramp down to 25°C

For spin programs 11- 14:• 25°C• 10min @ 50°C• 10min @ 65°C• 35min @ 75°C• 5°C/5min ramp down to 25°C

14 ILP Development SU-8 100(#lith129)

NL-CLR-WB24 • Developer: PGMEA (RER600, ARCH Chemicals)• Time: ~3:30min with spray-gun• Rinse with RER600• Rinse with IPA• Spin dry• Check result and perform extra cycles if not complete

15 ILP Lithography - Hard bakeSU-8 100(#lith131)

NL-CLR-WB24-hotplateStart @ 25°C• 10min @ 50°C• 10min @ 65°C• 10min @ 100°C• 2hrs @ 120°C• 5°C/10min down to 25°C