Vein-On-A-Chip: A Microfluidic Platform for …...1.3.1 Immune Function and Selective Barrier Function of Venous System The vein wall functions as a barrier between the intra- and

Vein-On-A-Chip: A Microfluidic Platform for Functional

Assessments and Staining of Intact Veins

by

Zhamak Abdi Dezfooli

A thesis submitted in conformity with the requirements

for the degree of Master of Applied Science

Institute of Biomaterials and Biomedical Engineering

University of Toronto

© Copyright by Zhamak Abdi Dezfooli (2015)

ii

Vein-On-A-Chip: A Microfluidic Platform for Functional Assessment

and Staining of Intact Veins

Zhamak Abdi Dezfooli

Master of Applied Science

Institute of Biomaterials and Biomedical Engineering

University of Toronto

2015

Abstract

The ability to perform systematic studies on vein function and structure is of key importance for

understanding many vascular diseases and the underlying mechanisms governing transport of

cells and macromolecules through the blood vessel wall, both of which are beneficial for drug

discovery. Available experimental methods are limited by poor optical access and limited

microenvironmental control for in vivo studies, and a partial representation of blood vessel

structure in the case of in vitro approaches. In this thesis, I present a microfluidic platform for the

in vitro investigation of intact mouse mesenteric vein segment (200-300 µm diameter and 1.5-2

mm length) function under near-physiological conditions. The lumen of the pressurized vein

segments are stained on-chip for CD31 to highlight the capacity of our platform for in-situ

immunofluorescence staining without the otherwise required tissue processing and sectioning.

The selected approach is automatable and reduces the time and staining solution consumption by

an order of magnitude compared to traditional methods.

iii

Acknowledgments

First and foremost, I would like to express my gratitude to Prof. Axel Guenther for introducing

me to this exciting subject, guiding me towards meeting my goals, and supervising me

throughout the project.

I would also like to thank our collaborator, Prof. Steffen-Sebastian Bolz, for his guidance and

valuable inputs. I am thankful for all the support I received from the members of Bolz lab

particularly, Dr. Firhan Malik for helping me in the development of the on-chip

immunofluorescence staining protocol.

I would like to express my appreciation to my committee members, Prof. Aaron Wheeler, Prof.

Jonathan Rocheleau, Prof. Dan Dumont, and Prof. Myron Cybulsky for their invaluable advice

and scientific guidance and contribution, as well as to Dr. Paul Van Slyke for his helpful

experimental recommendations with regards to working with leukocytes.

I would also like to thank the supervisory committee at The Centre for Microfluidic Systems,

specifically Ms. Lindsey Fiddes for providing technical support in regards to device fabrication

and imaging.

Moreover, my special thanks to all the members of Guenther lab for their suggestions and

support, specifically Dr. Ali Oskooei for helping me in the design of the temperature control

system.

Last but not least, I am very much thankful to my family. To my parents and my sister who have

always believed in me and for their support in every sense of the word throughout my whole life.

I am indebted to my two amazing children for being so patient with me and for supporting me

throughout the years of my studies.

iv

Table of Contents

Acknowledgments .......................................................................................................................... iii

Table of Contents ........................................................................................................................... iv

List of Tables ................................................................................................................................. vi

List of Abbreviations .................................................................................................................... vii

List of Figures ................................................................................................................................ ix

List of Appendices .......................................................................................................................... x

Chapter 1 : Introduction .................................................................................................................. 1

1.1 Hypothesis ........................................................................................................................... 2

1.2 Specific Aims ...................................................................................................................... 3

1.3 Background ......................................................................................................................... 3

1.3.1 Immune Function and Selective Barrier Function of Venous System .................... 3

1.3.2 Microfluidic Studies in Blood Vessels ................................................................... 5

1.4 Thesis Structure .................................................................................................................. 7

Chapter 2 : Device Study .............................................................................................................. 8

2.1. Device Design and Fabrication ........................................................................................... 8

2.2. Physical Characterization .................................................................................................... 9

Chapter 3 : Experimental Methods ............................................................................................. 16

3.1. Microfluidic platform ........................................................................................................ 16

3.2. Vein isolation and functional response assessment .......................................................... 18

3.3. Immunofluorescence Staining Procedure ......................................................................... 20

Chapter 4 : Results ...................................................................................................................... 22

4.1. Functional Responses ........................................................................................................ 22

4.2. Immunofluorescence Staining .......................................................................................... 24

Chapter 5 : Discussion ................................................................................................................ 27

v

Chapter 6 : Conclusion ................................................................................................................ 31

Chapter 7 : Future Directions ...................................................................................................... 32

References ..................................................................................................................................... 34

Appendices .................................................................................................................................... 40

vi

List of Tables

Table 1: Young’s modulus measurement of chip-hosted mouse mesenteric veins at different

pressure ...…………………………………………………………………………………….. 15

Table 2: Protocol for on-chip immunofluorescence staining of intact vein endothelium ……... 21

vii

List of Abbreviations

Endothelial cell EC

Smooth muscle cell SMC

Acetylcholine Ach

Phenylephrine PE

Bovine serum albumin BSA

3- (N-morpholino) propanesulfonic acid MOPS

Norepinephrine NE

Leukotrienes LTs

polymorphonuclear leukocytes PMNLs

cluster of differentiation 31 CD31

Tumor necrosis factor TNF-α

Interleukin 8 IL-8

Intercellular adhesion molecule 1 ICAM1

Junctional adhesion molecule JAM

Endothelial cell-selective adhesion molecule ESAM

Platelet/endothelial-cell adhesion molecule 1 PECAM-1

Vascular cell-adhesion molecule 1 VCAM1

Immunoglobulin superfamily Igs

Poly-(dimethylsiloxane) PDMS

Particle Image Velocimetry PIV

Paraformaldehyde PFA

Phosphate Buffered Saline PBS

viii

TRITC Tetramethylrhodamine

TxRed Texas Red

Nitric Oxide NO

Lipopolysaccharides LPS

ix

List of Figures

Figure 1: A schematic illustration of the leukocyte migration cascade in venules …………… 4

Figure 2: Microfluidic device design …………………………………………………………. 9

Figure 3: Flow characterization within the microfluidic device ……………………………… 11

Figure 4: Demonstration of sheath flow in the microchannel .………………………………... 13

Figure 5: Fixation channel design …………………………………………………………….. 15

Figure 6: Illustration of the experimental setup ………………………………………………. 17

Figure 7: Vein isolation and functional response assessment ………………………………… 19

Figure 8: On-chip functional assessment of mouse mesenteric vein …………………………. 23

Figure 9: Immunofluorescence staining of CD3 ……………………………………………… 25

Figure A1: Experimental setup- photograph and side view illustration ……………………… 40

Figure A2: Illustration of temperature control setup …………………………………………. 42

Figure A3: Dose response data ……………………………………………………………….. 43

Figure A4: Staining data ……………………………………………………………………… 45

Figure A5: Bubble Formation Test ………………………………………………………….... 48

x

List of Appendices

A1. Experimental setup ………………………………………………………………………... 40

A2. Temperature control setup…………………………………………………………………. 41

A3. Dose Response Data ………………………………………………………………………. 42

A4. Staining Data ……………………………………………………………………………… 44

A5. Bubble Formation Test ……………………………………………………………………. 47

1

Chapter 1 : Introduction

Veins are the blood vessels of the vascular system that are responsible returning blood flow to

the heart. Vein wall consists of three main layers: an outer layer that is called tunica adventitia

and is composed of connective tissue, nutrient vessels and autonomic nerves; a middle layer

called the tunica media is made up of smooth muscle cells (SMCs), elastic fibers and connective

tissue and is the thickest layer that provides structural support, vasoreactivity and elasticity; an

inner layer called the tunica intima with endothelial cells (ECs) and a small amount of sub-

endothelial connective tissue. The diameter of the lumen and the thickness of the wall of venous

blood vessels vary between 3 cm and 1.5 mm (i.e. vena cava) to 20 µm and 2 µm (i.e. venule),

respectively, depending on the position of the vascular bed. Therefore, compared to arteries,

veins have thinner wall and larger inner diameter due to significantly less SMCs and elastic

tissue in the structure of the middle layer- the tunica media.1-3 Veins play significant roles in

various physiological processes, respond to chemical and physical stimuli, synthesize bioactive

substances within the vascular wall, and either promote or prevent the transport of cells and

biomolecules. The soft wall of veins makes them vulnerable to irregular dilation, compression

and penetration by tumour and inflammatory processes. Diapedesis of leukocytes and plasma

macromolecule extravasation during inflammation are two examples of the transport processes

through the vein wall during inflammation.2, 4-6 However, the level of systematic research in the

field remains limited, in part because of experimental challenges related to the work with intact

organs. This is due to the very fragile and fine structure of the vein tissue and susceptibility of

the blood vessel to different external stimuli.

Common in vivo methods such as non-invasive, invasive and in situ cannulation techniques for

studying veins are challenging because of different technical and physiologic related limitations.

The complex interplay between different parameters e.g. signals from surrounding tissues and

the inability to independently alter chemical cues in the vessel microenvironment are limiting

factors concerning in vivo platforms. Also, limited spatial resolution due to high signal-to-noise

ratio when imaging at a distance and through centimetres of tissue in the body, as well as lack of

optical access due to limitation in imaging depth are other technical challenges of in vivo

approaches. For example, acquiring precise images by imaging modalities such as single- and

two-photon techniques usually can only be achieved to a depth of 100 µm from the surface of the

2

tissue, while much deeper penetration is needed for in vivo imaging. Another important

limitation is physiologic movement from breathing, cardiac contractions and the pulsatile flow of

blood, which negatively affect the sharpness of images compared to in vitro methods. However,

in vivo techniques are valuable because of the ability to maintain physiological conditions.7-11 On

the other hand, existing assays for in vitro studies of blood vessels for evaluating physiological

functions or drug development often use a confluent layer of isolated endothelial cells (ECs).

These models, to a limited extent, recapitulate important pathophysiological features experienced

by veins in vivo, e.g., correct cell alignment or perfusion at a defined shear rate. Such assays

include perfusion chambers, microfluidic devices and Boyden chambers.12, 13

Current in vitro technologies to study intact vein function and structure, either mounting the

vessel segment on two wires (isometric approach) or cannulation and perfusion with glass

micropipettes (isobaric approach),14-16 require skilled personnel and are not scalable; i.e.,

increasing the throughput proportionally consumes more time and workload. Techniques

available for whole tissue staining require the expertise to perform complex, multi-step

procedures, which consume both time and staining solution. The time and solution consumption

for whole mount fluorescent immunohistochemistry are on average 3-5 days for sample

preparation and on the order of milliliters, respectively, including 2 hours to one day fixation and

1-4 days incubation with the primary antibody.17, 18 Moreover, in vitro staining of the intact

blood vessel lumen is very challenging and difficult to perform using existing methods due to

inaccessibility of the cells. Operating an experiment with conventional methods that involves

functional assessment, staining, and imaging of an intact vein segment requires manipulation of

the specimens between three to four different instruments: the pressure myograph, tissue

processor, microtome (if tissue sectioning is required), and confocal microscope. Hence, an in

vitro approach that is capable of maintaining an intact and functional vein under physiological

conditions and which is also scalable i.e., increasing the throughput does not equivalently

increase the reagent, time and workload consumption, is a promising tool for the investigation of

blood vessel function at the cellular level.

1.1 Hypothesis

I hypothesize that a microfluidic-based approach to pressure myography assures the maintenance

of an isolated intact small vein under physiological environmental conditions, and makes in situ

3

functional assessment and immunofluorescence staining of the blood vessel lumen possible with

1 to 5 days reduction in time, 10 fold reduction in solution consumption, and reduced manual

operation than that associated with conventional techniques, and controlled physical and

chemical microenvironment.

1.2 Specific Aims

The following experimental strategy was carried out in order to probe the hypothesis:

i. The design, fabrication and testing of the microfluidic device for hosting and reversibly

fixating an isolated mouse mesenteric vein.

ii. The development of a protocol for the functional assessment of the chip-hosted vein to

apply abluminal flow of vasoactive substances under physiological conditions.

iii. The development of a protocol for immunofluorescence staining of the lumen in a chip-

hosted vein segment.

1.3 Background

1.3.1 Immune Function and Selective Barrier Function of Venous System

The vein wall functions as a barrier between the intra- and extravasal spaces; yet, it is rather

locally selective, and regulated. It is well known that the post-capillary venules involve the

exchange processes of the tissues and contribute in the resorption of interstitial fluid and

transportation of large molecules.19, 20 These vessels are composed of endothelial cells, pericytes

and a basement membrane.21 Studies show that the vascular endothelium permeability to

macromolecules increases and is dependent on the activation of a receptor-mediated

physiological mechanism.22 Inflammatory mediators such as histamine, serotonin, TNF and

bradykinin promote the extravasation of macromolecules from venules.23-25 The flux of

macromolecules through the blood vessel wall is associated with the formation of leakage sites in

the post-capillary venules.4

Furthermore, venules and post-capillary venules regulate the diapedesis of polymorphonuclear

leukocytes (PMNLs) and are the primary sites for leukocytes extravasation.26 Studies have

shown that there are specific locations in the basement membrane of venules that are closely

associated with gaps between pericytes and contain low extracellular matrix protein. These sites

are preferred pathway for transmigrating leukocytes during their passage through venules.21, 26, 27

4

Leukocyte migrate through a multiple cascade paths, mediated by fluid shear stress along with

chemical cues, and consists of the following steps (Fig. 1): The first step is referred to as

margination. Interaction between leukocytes and the large number of disk-shaped erythrocytes

that are contained in the complex fluid blood traveling through post-capillary venules initiates

the circulating leukocytes to migrate from the venule centre towards the wall, thereby initiating

surface interaction between leukocytes and endothelial cells. The interactions initially take place

via selectins (i.e., E- selectin, P-selectin and L-selectin) and their counter-ligands and are strong

enough to counter lift forces but too weak to firmly attach the leucocytes to the endothelial layer

in the presence of fluid shear. Leukocyte rolling is then followed by firm adhesion to the

endothelium. The final stage is the transendothelial migration which is either paracellular or

transcellular. The paracellular migration is through intraendothelial junctions. Transcellular

migration is through transcytotic pathways in the continuous endothelium. These pathways are

responsible for the transport of cells and macromolecules across the endothelial barrier by a

vectorial transport mechanism into the inflamed tissue.28-31

Figure 1: A schematic illustration of the leukocyte migration cascade in venules. The migration

process consists of margination, rolling, adhesion and transendothelial migration. The adhesion

comprises of attachment and intravascular crawling and transendothelial migration is done

through the venule wall and their respective molecular components involved.

5

1.3.2 Microfluidic Studies in Blood Vessels

In the past decade, several microfluidic tools have been developed for the study of different

aspects of blood vessel physiological function.32-34 In comparison to conventional cell and tissue

culture methods, these platforms and assays have a higher throughput with a smaller sample and

reagent consumption. These methods also provide precise control over the physical and chemical

characteristics that are relevant to vascular function, such as flow, shear stress, and reagent

concentration of the microenvironment. They assure better optical access, as well as more

physiological and pathological relevant conditions.

Most of the reported approaches are based on the mono- or co-culture of EC on either a stiff

microchannel wall or an engineered gel within a microfluidic device. Khan et al. studied the

inflammatory state of endothelium by assessing the effects of disturbed flow on ECs. To

generate disturbed flow, they designed a microfluidic chip which consisted of patterned channels

lined with ECs. Under constant flow, altering the geometry of channels will change the shear

stress applied on the ECs, which resulted in a change in activation and expression of cell

adhesion molecules such as VCAM-1 and ICAM-1 on the endothelial cell layer. They also

quantified the adhesiveness of the ECs to leukocytes by measuring the number of adherent

cells.35 Han et al. reported a platform that simulated the 3D configuration of neutrophil

transmigration during inflammation by placing an extracellular matrix with collagen type I in a

microfluidic device. Using a chemo-attractant gradient in this microfluidic device, they

quantitatively visualized the effect of chemoattractants e.g., IL-8 and fMLP on neutrophil

transmigration through a layer of cultured ECs. The researchers noted that the number of

migrated neutrophils and the distance they travelled through the collagen gel decreased in the

absence of intact endothelium, despite the presence of an optimal fMLP concentration gradient.

They also observed that the number of migrated neutrophils showed a reverse correlation with

the stiffness of the type I collagen.36 Schaff et al. studied the effect of shear stress on the

interaction of neutrophils and cultured ECs in a microfluidic platform, and measured neutrophil

rolling velocity and the adhesion rate under a defined shear stresses. Using their platform, they

confirmed that the rate of the neutrophil recruitment to the substrate was very sensitive to

disturbances in flow streamlines as it was increased at the entrance of the microchannels. They

also showed that dose response of stimulation of the ECs by the chemokine IL-8 can trigger β2

integrin activation.37 Zhang et al. developed an in vitro vascular system that recapitulated the

6

geometry of native vasculature with a circular microchannel. This was used to investigate the

relationship between shear stress applied on the ECs cultured in the channels and dose of the

applied pharmacological agent. They validated their platform for drug screening by examining

the effects of four vasoactive drugs on the secretion of nitric oxide from the endothelial cells

under physiological conditions: acetylcholine, phenylephrine, atorvastatin, and sildenafil. This

study demonstrated the importance of physiologically-realistic endothelial cell geometry and

flow rates in drug validation applications. The researchers also demonstrated a monocyte

adhesion assay, which could be used for the assessment of efficiency of anti-inflammatory

drugs.38 Namdee et al. explored how particle size, along with hemodynamics (blood shear rate

and vessel size) and hemorheology (blood hematocrit) affect the capability of spheres to

marginate (localize and adhere) to inflamed endothelium in a microfluidic model of human

microvessels. They used cell free plasma from whole blood and showed that microspheres

present excessively higher margination than nanospheres in all hemodynamic conditions.39 Tsai

et al. reported a microfluidic vasculature model that recapitulate platelets, leukocytes, and

endothelial cells activation under controlled flow conditions. The microfluidic device enabled

quantitative study on how biophysical alteration, for instance shear stress, in hematologic

diseases such as sickle cell disease (SCD) and hemolytic uremic syndrome (HUS), cause

vasculature occlusion and thrombosis. It consisted of bifurcated microchannels that were

luminally covered with monolayer of endothelial cells that resembled a microvasculature system.

The researchers used the model for studying how the drug hydroxyurea quantitatively affects

microvascular obstruction in SCD by applying blood sample from patient with SCD to the in

vitro microvasculature and investigating the effect of an antiplatelet drug on aggregation of

platelets with changing shear. In addition, they used their microsystem as an in vitro model of

HUS and showed that shear stress influences microvascular thrombosis/obstruction and assessed

the effectiveness of the drug eptifibatide, which decreases platelet aggregation in patients with

HUS.40

All of the preceding studies were based on the culture of EC monolayers on either a stiff

microchannel wall, or a gel surface within a device in the absence of other components of the

blood vessel wall i.e. perivascular basement membrane and the connective tissues. Hence, the

cultured monolayers cannot form physiologically relevant levels of junctional complexes.

7

Compared to cell culture-based platforms, tissue and organ-based microfluidic approaches have

also been reported that have better representation of in vivo conditions.

Our research group previously developed a microfluidic platform for the assessment of isolated

resistance artery structure and function under physiological conditions, 37 °C temperature and 45

mmHg transmural pressure.41 Small arteries were isolated from a mouse mesentery, loaded onto

the microfluidic device, and immobilized by applying sub-atmospheric pressure on the blood

vessel wall through microfluidic channels. These arteries were kept intact in well-defined

microenvironments that reflected the in vivo conditions of terminal arteries. This platform

isolated the luminal and abluminal surfaces of the blood vessel, which allowed for localized

stimulation of the blood vessel. This capability was validated by dose-response measurements to

the abluminal application of phenylephrine and acetylcholine, which matched the results from

conventional myography techniques. An automated microfluidic platform has further been

developed by our research group that is capable of on-chip staining of smooth muscle cells and

functional assessment of cerebral resistance arteries under physiological environment

conditions.42 Using this platform, the abluminal structure of the blood vessel was visualised by

imaging the position of the smooth muscle cell nuclei, actin filaments and voltage gated calcium

channels. The smooth muscle cell function and intracellular calcium were assessed

simultaneously by staining the SMCs with FURA-2AM during exposure to varying

concentration of PE (a vasoactive mediator) with and without a calcium blocker incubation,

which provided a better understanding of vasomotor dynamics and calcium response.

1.4 Thesis Structure

This thesis is organized as follows: parametric study on the microfluidic device is carried out in

the Chapter 2. Chapter 3 conveys the experimental methods employed, including the chip design

and experimental setup, the device characterization and the protocols for functional assessment

and immunofluorescence staining. Chapter 4 presents the obtained results from the experimental

strategy. Chapter 5 communicates the discussion on the approach and the results. Finally,

Chapter 6 and 7 summarize the findings and suggest potential applications of the described

approach, respectively.

8

Chapter 2 : Device Study

2.1. Device Design and Fabrication

The design of the microfluidic device is a modified version of the devices previously introduced

by our group for the functional assessment of mouse mesenteric and cerebral resistance

arteries.41, 42 The design was prepared using a computer aided design program (Autocad 2011,

San Rafael, CA, U.S.A), which consisted of a loading well and four microchannel networks as

follows (Fig. 1): a network of fixation channels (indicated in blue color) which held the vein

segment in the desired position within the inspection region. The second channel network

(indicated in yellow and green and color) comprised two lines, including one core channel and

two side channels that merged upstream of the microchannel and connected to the blood vessel

luminal side. The inlet of the core-channel of the perfusion line (yellow) was connected to a

single reservoir to facilitate the blood vessel loading procedure. The side-channels inlet (green)

was connected via a selector valve to one of six individual reservoirs and allowed for the

controlled luminal perfusion of different solutions. Besides, the two channels were designed to

produce co-flow of two different solutions using flow focusing technique. This facilitates further

application of the platform in investigation of the leukocyte-ECs interaction by flowing the cells

through the side channels and positioning them at the vicinity of the endothelium. The third

microchannel network, superfusion channels (indicated in red color), defined the vein

microenvironment on the abluminal side and allowed physiological buffer, vasoactive drugs and

inflammatory mediators to be superfused on two opposing sides, for application of different

concentrations. Finally, a vacuum line was connected to two previously reported bubble traps43

that were located in the superfusion channels which prevented bubbles from getting through the

inspection area and damaging the blood vessel segment. All channels were 150 µm deep and

smaller than the diameter of the pressurized and unconstricted vein to prevent the blood vessel

segment from passing through the inspection area during loading. The width of the different

microchannels varied between 30 µm at the four fixation locations and 500 µm at the fixation

and superfusion channels. The microfluidic device was fabricated in PDMS

(polythimethysiloxane), using standard single-layer soft lithography techniques, and permanently

sealed on a microscope glass slide (thickness 1mm).

9

Figure 2: Microfluidic device design. The device comprised a loading well and four sets of

microchannel networks: fixation channels presented in blue for holding the vein segment in the

inspection area, perfusion channels with two inlets indicated in yellow and green merge upstream

of the microchannel that connect to the blood vessel luminal side and the loading well,

superfusion channels (colored in red) that had two separate pairs of inlets and are connected to

one outlet with two bubble traps on each line, and one vacuum line for the two bubble traps.

2.2. Physical Characterization

To experimentally examine the shear stress exerted on the walls in our system, flow velocity in

different regions of the inspection area was measured by micro- particle image velocimetry

(micro-PIV). A PDMS device was designed and fabricated to mimic the geometry of the original

microfluidic device with a feature of pressurized blood vessel on the perfusion line and two

inlets for the side flow and one inlet for the core flow (Fig. 2A). The micro-PIV measurement

was done by flowing 1.1 µm fluorescent particles through the core channel of the perfusion line

from a reservoir connected to the inlet. The two side channels were blocked and the inlet

pressure was maintained at 4.5 mmHg, using hydrostatic pressure to represent the transmural

pressure in the vein-on-chip microfluidic platform. The device outlet was connected to a syringe

pump in withdrawal mode with a constant flow rate of 1 mL/hr (3×10-4 cm3/s). Fluorescent

particles were illuminated by a laser (λ=550 nm) and visualized by an inverted microscope with

20× objective (field of view: 435µm×325µm). Images were recorded at a rate of 5 frames per

second.

10

In order to determine the flow parameters in the perfusion channel, we assumed that the flow

was fully developed. To justify this assumption, the hydrodynamic entrance length Ln for the

flow through the perfusion channel was calculated according to the equation (1):44

(1)

In the part of the channel before the expanded section with hydraulic diameter (Dh) =168 µm and

the Reynold’s number (Re) = 2.5, Lh was equal to 414 µm. This shows that the flow was fully

developed when it passed through the 22.5 mm long channel. In addition, according to equation

(1) hydrodynamic entrance length for the expanded section was equal to 373 µm, while the

length of the section was 550 µm. This number could vary between 420 µm (i.e. Dh = 150 µm)

and 383 µm (i.e. Dh = 194 µm) for flow through the chip-hosted vein, depending on the size of

the blood vessel which influences the width of the expansion in the blood vessel due to the

applied transmural pressure. Therefore the flow gradually became fully developed after half way

through the section. For simplicity, this divergence was not considered in the analytical solution

and calculation of the shear stress. Assuming fully developed flow, i.e., a parabolic velocity

distribution, the cross sectionally averaged velocity obtained from micro-PIV measurements in

the channel at the expanded section which was representative of a pressurized blood vessel 300

µm wide was recorded to be 6.5 mm/s (Fig. 2B). This number is very close to the result from

analytical calculations which was 6.2 mm/s. However, the difference between the two parabolas

in the figure, is as a result of the channel width extension from 200 µm to 300 µm and the

variation in the entrance length, as discussed above.

The shear stress applied to the walls of a micro-channel with rectangular cross-section due to a

fluid flow was calculated by .45 In this equation, (dyne/cm2) is the fluid viscosity, Q

(cm3/s) is the flow rate, and w (cm) and h (cm) are the width and height of the blood vessel in the

inspection area, respectively. Considering a fluid viscosity of 0.007 dyne s/cm2 for the buffer, the

mentioned velocity caused shear stress of about 1.6 dyne/cm2. This number falls in the minimum

range of shear stress applied on the wall of blood vessel in an inflamed tissue.46, 47

11

Figure 3: Flow characterization. (A) Schematic of the experimental setup for micro-PIV

measurement. The pressurized blood vessel-feature is magnified. (B) Analytical and

experimental velocity profile resulted from the micro-PIV measurement.

Moreover, the physical parameters of the flow-focusing geometry employed in the design of the

perfusion channel were characterized as follows: the same microfluidic device used for flow

characterization in the previous experimental setup was used. Pure 3- (N-morpholino)

propanesulfonic acid (MOPS) was perfused through the core channel of the perfusion inlet at a

flow rate of QB via a syringe pump. Fluorescent buffer (rhodamine in MOPS) was flowed

through the two side channels of perfusion inlet from a reservoir. Because the two channels were

identical, the flow rate in each channel was half the flow rate of the reservoir, QL. Outlet of the

perfusion channel was connected to a syringe pump in withdrawal mode and the flow rate was

set at Q, which was also equal to the total rate of QL+QB. Therefore, altering the core channel

flow rate, QB, resulted in a change in the side channels flow rates, QL. A relationship between the

relative flow rates, QL/Q, and relative thickness, δ/w (where δ and w are the side flow width and

the channel width at the expanded area, respectively, as are shown in Fig 3A) can be derived

from the parabolic velocity profile:

32

46

wwQ

QL (2)

Equation (2) indicates that altering the core channel flow rate (QB) results in a change in the

thickness of the side flow solution, δ. Fluorescent images were taken while increasing the flow

rate QB to experimentally approve the control over thickness of the side flows, and the resulting

12

data was plotted (Fig. 3A and 3B). Experimental results from varying relative thickness (δ/w) vs.

relative flow rate (QL/Q) was in agreement with theory, as shown in Fig. 3C. This characteristic

gives rise to the ability of controlling the concentration of the side flow solutions. Also,

thicknesses of the fluid in the side channels (δ) were measured for different core channel flow

rates (QB) which confirmed the consistency between the top layer and the bottom layer (Fig. 3D).

As shown in Fig 3B, the velocity gradient close to the wall of the microchannel is similar to the

one in the dimension of small venules, thus the shear stress applied to the wall by the fluid flow

is the same as in vivo condition in small venules. All of the above mentioned flow characteristics

of the device support further application of the platform as a tool for studying leukocyte

migration.

13

14

Figure 4: Sheath flow. (A) Fluorescence imaging of the channel with a feature of pressurised

blood vessel in the middle that shows the control of the thickness of cell populated side regions.

(B) Velocity profile in pressurized blood vessel shows the velocity gradient in the vicinity of the

vessel wall is the same in both vein and postcapillary venule. (C) Experimental results of

changes in relative thickness (δ/w) vs. relative flow rate (QL/Q) was in agreement with the

analytical prediction. (D) Thickness of the fluid in side channels (δ) were measured for different

core channel flow rates (QB) and showed consistency between the top layer and the bottom layer.

Furthermore, the design of the fixation channels was justified for veins by calculating the width

of the channels from Young’s modulus ( E ) based on half-space model as follows:48

(3)

where hD is hydraulic diameter of the fixation channels, P is the difference between the luminal

and the fixation pressures, AL is the length of the blood vessel wall that penetrates into the

fixation channel after applying pressure to the channel and is a dimensionless constant

parameter (wall function) which is set to 2.1.48

A vein segment was loaded onto the chip device (loading procedure will be described in the next

chapter) and the pressure applied to the fixation channels was incrementally changed and the

bright field images were recorded. In Fig. 5, the images labelled (a) to (d) show the increased

aspiration length (i.e. length of the blood vessel wall entered into the fixation channels) by

increasing the fixation pressure from 0 mmHg to 22 mmHg. The images were processed and the

Young’s modulus was calculated according to equation (3); The results shown in Table 1 are in

agreement with the data obtained literature.49 Then the fixation pressure was set at 22 mmHg and

the transmural pressure was incrementally changed by increasing the height of the hydrostatic

head (Fig 5, images e and f). Circumferential elastic modulus (Eθ) of the blood vessel wall was

calculated by measuring the inner and outer diameter according to the following equation:50

)(

)1(222

00

2

0

2

i

ii

RRR

RRPE

(4)

where ΔPi is the transmural pressure increment, R0 and Ri are the external and internal,

respectively, ΔR0 is the change in external diameter due to the change in transmural pressure,

and δ is the Poisson ratio which is 0.45 for isotropic elastic material. The circumferential elastic

modulus of the vein segment was calculated based on equation (4) and was 3.7 kPa and 5 kPa for

4.5 mmHg and 11 mmHg change in transmural pressure, respectively.

15

Figure 5: Fixation channel design. Images labelled as (a) to (d) show unpressurized vein

segment for fixation pressure from 0 mmHg to 22 mmHg, respectively. The aspired section of

the blood vessel wall is marked in red. Images (e) and (f) indicate pressurized vessel at 4.5

mmHg and 11 mmHg transmural pressure, respectively. Scale bar represents 200 µm.

Table 2: Young’s modulus measurement of chip-hosted mouse mesenteric veins at different

pressure (n=3)

The angle of the fixation channels applied a tension to the wall of the vein segments to mimic the

in vivo tension on the blood vessel wall. Moreover, veins are less stiff than arteries because their

walls are covered with less SMCs, therefore they tend to collapse easily and it is challenging to

keep the lumen open. The direction of the angles in the design of the fixation channels helped to

overcome this problem by pulling the tissue towards the two ends of the vessel segment at each

side.

Ffix

(mmHg)

E (kPa) Ref. #6

E

(kPa)

7.4 3.9±0.6 4.8±0.7

14.7 11.1±0.8

22.1 18.1±0.9

16

Chapter 3 : Experimental Methods

3.1. Microfluidic platform

The experimental platform for on-chip functional assessment and immunofluorescence staining

of vein comprises of the previousely described microfluidic device which was connected to five

computer controled syringe pumps (Harvard Apparatus, Holliston, MA) four of which were

connected to the superfusion inlets and run in perfusion mode for delivery of different

concentration of drugs and physiological buffers to the abluminal side of the blood vessel and the

remaining one was connected to the perfusion outlet and operated in withdrawal mode. The inlet

of the perfusion side channels was connected to a computer controlled selector valve (VICI

Valco Instruments Canada Corporation, H-C25Z-3186EUHA) and the other perfusion inlet (the

core channel) was connected to a reservoir. The superfusion channel outlet and the fixation

channel inlets were connected to three open, MOPS filled reservoirs. The height difference

between the two reservoirs would define the fixation pressure for holding the blood vessel. The

two bubble traps in the superfusion lines were connected to a miniaturized vacuum pump

(Parker’s CTS Micro Diaphragm Pump, USA) via the manifold. A stereo microscope (model:

Nikon) and an inverted fluorescence microscope (model Ti Eclipse, Nikon, Japan) were

connected to a CCD camera (model EXi Blue, QImaging, Surrey, BC, Canada) for blood vessel

loading and imaging, respectively. An automated temperature control system maintained the

desired temperature during different stages. A peristaltic pump (Fisher Scientific 13-875-410

DIG. FH100 PUMP 14-4000ML/MIN EA) was employed as a heat sink for the cooling system

(details in Appendices A1). A commercially available manifold with ten fluidic inputs/outputs to

interface with the PDMS device by means of luerlock connectors (Idex Health & Science, Oak

Harbor, WA) was employed to connect the fluidic inlets and outlets with tubing.

17

Figure 6: Experimental setup. Schematic illustration of the vein-on-a chip platform consisting

of a microfluidic device inserted into the manifold (1), five computer controlled syringe pump

four of which are connected to the superfusion inlets and one is connected to the perfusion outlet

(2), a computer controlled selector valve connected to the perfusion inlet (3), an automated

temperature control system (4) and a vacuum system (5).

The physiological pressure and temperature were provided as previously described.41 Briefly, the

temperature was preserved using a thermoelectric element (model TE-35-0.6-1.0, TE

Technology) attached to a sapphire glass which was placed on top of the device, where the blood

vessel segment was positioned. A thermistor (model MP-2444, TE Technology) was embedded

into the microfluidic device at the close vicinity of the vein segment during the PDMS moulding

that allowed for on-chip control of the temperature in the inspection area. In order to achieve a

cold temperature (4˚C) for the purpose of immunofluorescent staining, an aluminium cooling

jacket was designed and attached to the TE element to operate as a heat sink to cool down the

warm side of the TE element. Details of the set up for maintaining a target temperature are

illustrated in Appendix A2. Physiological transmural pressure of 4.5 mmHg for small veins51

was maintained by applying hydrostatic head at the both perfusion inlets (Fig. 5).

18

3.2. Vein isolation and functional response assessment

For functional assessment, vein segments were isolated from 4-8 month old C57BL/6 mice. A

segment (approximately 1-2mm in length) of mouse mesenteric vein (diameter approximately

200-300 μm) was isolated by removing the adhering fat and connective tissue (Fig. 6A and 6B).

The order of the mesenteric veins refers to the bifurcating branches inside the mesentery. The

major vein supplying the mesentery is the first-order vein and subsequent branches accordingly

increase in their order.52 The isolated vein segment was reversibly mounted onto the microfluidic

device as previously described by Guenther et al.41 In summary, the vessel was placed in the

loading well and was drawn through the perfusion channel to the inspection area where it was

reversibly immobilized at eight points using negative hydrostatic head, which were set at 22

mmHg. While maintaining physiological temperature (37C) and transmural pressure of 4.5

mmHg assumed for small vein, viability of the blood vessel’s structural component was

confirmed by dose response measurements performed with vasoactive substrates (Fig 6C).

Intactness of the smooth muscle cells (SMCs) was confirmed through contractile responses in the

presence of increasing concentration (0.5, 2.5, 5, 10µM) of phenylephrine (PE) that is superfused

at a constant flow rate of 1 mL/hr. Subsequently, the endothelial cells (ECs) were assessed

through the dilatory response of the pre-constricted vessel in the presence of increasing

concentrations (0.25- 1 µM) of acetylcholine (Ach). Bright field images of the vein segments

were obtained using a CCD camera (1392 × 1040 pixel resolution, model EXi Blue, QImaging,

Surrey, BC, Canada).

19

Figure 7: Vein isolation and functional response assessment. (A) Illustration of the order of

branching microvessels through schematic and bright-field micrograph of mouse mesentery.

Roman Numbers in the right image indicate the blood vessel order. Scale bar 1 mm (B) Bright-

20

field micrograph illustrating mesenteric vein isolation. Scale bar 250 µm (C) Schematic of the

experimental setup for dose response assessment. (D) Schematically illustration of the top view

(middle image) and the vertical cross section (bottom) and the horizontal cross section (right) of

the chip hosted, pressurized vessel.

3.3. Immunofluorescence Staining Procedure

To examine the microfluidic device functionality for luminal staining, immunofluorescence

staining for CD31 on endothelium was performed. In this set of experiments, six reservoirs were

connected to the perfusion channel. Via a selector valve, the reservoirs opened to the perfusion

channel one at a time, for the purpose of executing immunofluorescence staining protocols on

the vein segments. After loading the vein segments onto the chip, the vessels were perfused and

superfused with MOPS at a constant flow rate of 4 µL/min for 2 hours at 37 °C to avoid any

probable stimulation related to isolation or loading. Then the chip was cooled down to 4 °C and

vein segments were fixed by perfusion of 4% paraformaldehyde (PFA) for 30 minutes, blocked

with 3% bovine serum albumin (BSA) in phosphate-buffered saline (PBS) for 1 hour and

incubated with 1:100 rat anti-mouse CD31 antibody (BD Biosciences) over night. All the steps

were performed with a flow rate of 4μl/min, except for incubation with primary antibodies,

where the flow was stopped after 5 minutes perfusion of the antibody. The temperature was

maintained at 4 °C during all the steps. Then the veins were washed with perfusion of PBS for 30

minutes and incubated with 1:200 goat anti-rat TRITC (in some cases TxRed) for one hour at

room temperature. Finally the veins were individually imaged using a confocal microscope

(model Ti Eclipse, Nikon, Japan) with magnification 10× (working distance: 4 mm, numerical

aperture: 0.45); and magnification 40× (working distance: 3.6 mm, numerical aperture: 0.6). To

obtain control data, a blood vessel segment was perfused with the secondary antibody for one

hour right after the blocking and washed with PBS for 10 min before imaging. Summary of on-

chip staining procedure is presented through table 2.

21

Table 2: Protocol for on-chip immunofluorescence staining of intact vein endothelium

Step Solution Time Temperature

(° C)

Perfusion

Flow rate

(µl/min)

Total

Volume

(µl)

Fixing 4% PFA in

PBS

30 min 4 4 137

Blocking 3% BSA 60 min 4 4 257

Primary antibody 1:100 rat anti-

mouse CD31

antibody

12 hrs 4 4 and 0 100

Wash PBS 30 min 25 4 137

Secondary antibody 1:200 goat

anti-rat

TRITC

60 min 25 2 137

Wash PBS 30 min 25 4 137

Total time 15.5 hrs

22

Chapter 4 : Results

4.1. Functional Responses

After loading the vein segment onto the microfluidic chip and heating the vessel to 37 °C and

pressurizing to 4 mmHg, the functional response of the SMC and EC layers was verified through

superfusion of varying concentration of PE and ACh respectively.

Figure 8 A and C, respectively, represent images from chip-hosted vein segments, abluminally

exposed to PE and Ach through the superfusion channels at the top and the bottom. Summary of

the results from the vein functional response to PE (n=5) and Ach (n=4) are shown in Fig. 8 B

and D. Captured images were then manually processed using ImageJ (ImageJ 1.48v, U.S.A.) to

quantify the contractile responses as explained in the discussion section.

23

Figure 8: On-chip functional assessment of mouse mesenteric vein. Representative images of

vein functional response to (A) PE and (C) Ach. (B) and (D) Dose response plots to PE (n=5)

and Ach (n=4), respectively. Scale bars are 200 µm. Error bars are standard error.

24

4.2. Immunofluorescence Staining

After completing the staining procedure, the manifold was detached from the fluidic connections

and transferred to the confocal microscope to picture the expression of CD31 on the ECs of the

blood vessel walls. Figure 9A shows a representative confocal image of intact whole vein

segment luminally stained for CD31. The intensity of the “top” and “bottom” walls of the vein

segment was measured from the orthographic projection of the confocal and plotted for four

stained sample and one control sample (Fig 9 B and C) for quantifying the data.

25

Figure 9: Immunofluorescence staining of CD31. (A) Confocal image of an intact vein

segment stained luminally for CD31 with 10× (bottom) and 40× (top, inset), respectively.

Arrows indicate the endothelial cell boundaries. Scale bars are 50μm (top) and 100μm (bottom).

26

(B) and (C) Orthographic projection and the respective intensity profiles of the confocal images

of stained (scale bar is 50 µm) and control (scale bar is 100 µm) samples obtained with 10×

objective. (D) Scattered plot of the two maxima obtained from the intensity profiles.

27

Chapter 5 : Discussion

The main advancement achieved with the presented platform is to keep an isolated small mouse

mesenteric vein segment alive and intact under well-controlled physical (including the

temperature, transmural pressure and perfusion flow rate) and chemical microenvironment

conditions, for functional assessment and immunofluorescence staining of the lumen.

Figures 8A-D show the dose response results from the chip-hosted vein segments, abluminally

exposed to phenylephrine (PE) and acetylcholine (Ach) through the superfusion channels at the

top and the bottom, respectively. Bright field images of the samples (Fig A3) indicated that the

contractile responses of mouse mesenteric veins to vasoactive drugs were different from the

contractile responses of arteries from the same vascular bed. Unlike the sample of arteries,41 in

our results vein wall did not thicken after exposure to PE, but collapsed. This is probably because

of the differences in the wall structure of the two types of blood vessel, i.e. the amount of the

SMC layers around the entire vein is much less than that of the arteries which affects the

vasoconstriction.1, 53 Therefore, it was not possible to differentiate between the inner and outer

borders of the blood vessel wall and only the outer lines of the vein wall were counted for

quantifying the functional responses. Also, as described in the experimental method section for

functional assessment, the isolated vein segments (diameter approximately 200-300 µm) were

loaded onto the microfluidic chip and sandwiched within the PDMS channel with 150 µm height

and the glass slide and the cross-section of the samples was imaged from bottom as

schematically illustrated in the Fig 7 D. Moreover, confocal images of the constricted vein

segments showed that not the entire vascular wall but the sides that were exposed to the

vasoactive drugs exhibit functional responses. Therefore, change in the width of the blood vessel

segments which represented the distance between the side walls (denoted by D), in response to

different concentrations of the vasoactive drugs, was used as a measure of percentage

constriction and dilation. In order to measure D (or D0) the images were post processed in

ImageJ by drawing a line that connected the top and bottom side of the vessel, as illustrated in

the figure. The constriction and dilation percentage of vessels were calculated by normalizing D

to the maximum and minimum vessel width (denoted by D0) i.e. before applying the constrictor

and dilatory mediators, respectively. Therefore the contraction percentage is estimated as (D0-

28

D)/D0. In our experiments, all the functional veins showed contractile response to PE and Ach

with the highest constriction at 10 µM and highest dilation at 1 µM, respectively.

Phenylephrine is known for its effect on SMCs to induce vessel contraction by binding to

adrenergic receptors on the SMCs. This results in the buildup of internal calcium which causes

increased tension within the cells around the intact blood vessel, leading to the entire vessel

constricting.54, 55 The results from on-chip measurements showed an increasing trend in the

contraction of the vein walls in response to PE. The maximum constriction was about 72% which

was achieved by superfusion of 10 µM PE. The maximum constriction is commensurate with

other reported data for rat mesenteric small veins, which were produced using pressure

myography technique, in response to 10-4 M norepinephrine at 2 mmHg and 6 mmHg transmural

pressure, i.e. ~85% and ~70%, respectively.52, 56

Acetylcholine has effects on stimulating intracellular nitric oxide (NO) production and release in

endothelial cells during blood vessel relaxation.57 Chip-hosted veins showed weak responses to

low concentrations of Ach and the maximum dilation was about 40% in response to 1 µM. This

number is less than the reported results from on-chip dose response measurements for arteries

(i.e. ~75%).41 This difference might be due to more endothelium-derived NO in arteries than in

veins. Other studies on arterial and venous grafts in patients after coronary bypass operations

seem to support our rationale.58

One important feature of our approach is the capability of in situ luminal staining of the intact

veins right after applying chemical (such as inflammatory mediators and drugs) or physical (such

as shear stress) stimuli on the blood vessel. To confirm this, endothelium of the chip-hosted vein

segments were stained for CD31. CD31 is a glycoprotein commonly used as an endothelial cell

marker. It forms a large fraction of endothelial cell intracellular junctions and is localized in the

lumen surface of blood vessels.59, 60 Fig. 3A and B demonstrates representative confocal images

of a luminally stained vein segment. Arrows in the top image of Fig. 3A indicate cell boundaries

which were fluorescently labelled. Configuration and alignment of the labelled cells validate that

the endothelial cells of the vein segment were stained. The intensity histogram from the

orthographic projection of the confocal images peaked at the region with low intensity and the

second peak fell in the region with higher intensity. The second peak of the counts fell in the

region with approximately 4 times higher intensity in the stained vessels compared to the control.

29

Quantitative data was obtained by measuring the mean intensity of “top” and “bottom” walls of

the blood vessels through orthographic projection of the confocal images. In order to quantify the

data from fluorescent intensity measurement, orthographic projection of the confocal images

were taken with the best visualized cross-section in the XZ direction and the intensity profiles

were created along arbitrary lines across the images as is shown in Fig. 3C and 3B. The maxima

in the intensity profiles represent the “top” and “bottom” walls of the vein segments. The top and

bottom maxima were normalized by subtracting and dividing by the intensity at the mid-point

between them, i.e. the lumen of the blood vessel. Fig. 3D shows the data obtained from the

maximum intensities of intensity profiles for four stained samples and one negative control.

Stained blood vessels demonstrated on average four times higher intensity in the regions where

CD31 was expressed compared to the negative control. Nonetheless, the inconsistency in the

intensity of the fluorescent images among the samples is mostly because of the fragility of the

vein wall, which at times resulted in the blood vessel segment collapsing in the microchannel.

This effect is due to small changes in the transmural pressure (e.g. Fig. A4 B), which needs to be

addressed in future experiments in order to improve the yield of the platform.

Finally, the described approach is advantageous over conventional whole tissue

immunofluorescence staining methods because it provides the possibility of in situ luminal

staining of the intact vein right after applying chemical (such as inflammatory mediators and

drugs) or physical (such as altering the shear stress) stimuli on the blood vessel. In contrast,

procedures for conventional whole tissue staining need primary preparation including sectioning

and paraffin embedding of the blood vessels. Moreover in our approach the consumed solution is

ten times less than in the conventional methods. Finally, our method is faster since on average

there is a three days reduction in the work process compared to whole tissue staining methods.

The introduced platform has a number of challenges which some have been resolved and some

need further investigation and effort in order to improve the yield. One important experimental

problem was the appearance of bubbles in the inspection area, despite the presence of the bubble

traps in the superfusion lines. The bubbles appeared either inside or outside of the blood vessel

during the experiments that required long time i.e. > 12 hours perfusion through the blood vessel

which often damaged the endothelium of the blood vessel or resulted in collapsing the lumen as

is shown in Fig A5. A and B. Four main reasons were identified through a set of tests that could

cause the appearance of bubble: a) Volume of the bubble trapes was 0.5 µl which would fill up

30

by large size bubbles and result in non-functioning. Implementing multi and parallel bubble traps

in the channels could be one approach to overcome this issue which has not been done yet. b)

Surface profilometry showed that the surface roughness of the sections of the manifold that were

in contact with the PDMS around the inlet holes was one source of the bubble formation.

Polishing the surface and preventing any damage to the surface improved the system in that

regards. c) Gas permeability of the fluidic tubing and syringes was another cause for bubble

formation which was resolved by changing the materials to less permeable substances, i.e. using

PEEK tubing (permeability: < 1 cm2/s) instead of Tygon tubing (permeability: 12- 30 cm2/s), and

gas-tight glass syringes instead of plastic syringes. d) Air diffusion through the manifold inlet

and the tubing connectors was another reason for the formation of bubbles in the channels, which

was temporary resolved by filling the gaps with epoxy glue. Details of the experimental setup

and conditions are explained in the Appendix A5.

Another problem regarding the experimental approach was linked to transport of the chip

containing a pressurized vein segment between different imaging modalities. As it was

mentioned earlier in this section, sudden change in the transmural pressure often resulted in

collapsed veins due to the very thin and limp wall of the blood vessel. The fluctuation in the

transmural pressure was mostly because of the change in the hydrostatic pressure applied to the

inlet of the perfusion channel which was maintained by holding a reservoir at a defined height.

One way of resolving this problem is to attach the perfusion reservoir to the manifold, thus

reduce the risk of any sudden fluctuation to the inlet pressure. Also, different approach for the

maintenance of the trasmural pressure, such as attaching a pressure transducer to the inlet of the

perfusion channel could potentially be helpful to overcome this problem.

Chip cracking was the other experimental difficulties which affected the yield of the platform.

The main reason behind this problem was the uneven force applied to the microscope slide

which was bonded to the PDMS device from the manifold. Although, this problem was less

notable when the thickness of the PDMS device was homogeneous along the chip which was an

issue relating the PDMS fabrication.

31

Chapter 6 : Conclusion

We have developed an automated microfluidic approach for studying the intact small veins that

enables functional investigations while maintaining the near-physiological conditions as well as

immunofluorescence staining of the lumen surface. Parametric study of the platform is

performed and the shear stress applied to the wall of the blood vessel is measured for a defined

flow rate identical to the flow rate perfused through the vein segment in blood vessel

experiments. Moreover, capability of the platform for generating sheath flow and control over

the thickness of the solutions within the on-chip pressurized blood vessel is confirmed. Also, the

design of the fixation channels for immobilizing a vein segment is justified through the

measurement of the elasticity of the veins in the chip. Furthermore, performance of the vein-on-

a-chip is evaluated through the following procedure: the mouse mesenteric veins are isolated and

manually loaded onto the microfluidic platform. Viability of the vein segments is assessed

through superfusion of varying concentrations of phenylephrine and acetylcholine. Functional

assessments of the endothelium and the smooth muscle cells show 72% and 40% constriction

and dilation of the veins, respectively. Finally, a protocol for in situ immunofluorescence

staining of the lumen of the chip-hosted veins is developed. Using this protocol, pressurized

veins are stained on-chip for CD31 markers confirming the capacity of our platform for in-situ

immunofluorescence staining, without the need for tissue processing and sectioning. The

selected approach is automatable and reduces the time for staining procedures by on average 3

times compared to traditional methods. Moreover, it requires ten times less staining solution and

antibodies, i.e., 100-300 µL for the vein-on-a-chip vs. 1 mL, for the current conventional

techniques, and is capable of performing all the experimental phases from functional assessment

to immunofluorescence and imaging in less than two days on each intact vein segment.

32

Chapter 7 : Future Directions

The established platform in this work can be used in further applications such as inflammation

studies at a molecular level. It is known that during inflammation, the endothelium of small veins

is activated due to the expression of selectins, their counter ligand, intracellular adhesion

molecule-1 (ICAM-1) and vascular cell adhesion molecule-1(VCAM-1) in response to chemical

and physical cues. Subsequently, recruitment of leukocytes from the blood stream into the

inflamed tissue takes place.5, 30, 61A proposed strategy could be to use the platform for

inflammatory activation of intact veins to quantify the expression of cell adhesion molecules

which could be exploited for detailed study of inflammation in cellular and molecular level.

One approach is superfusion of inflammatory mediators and chemical stimulation of the blood

vessel by the drugs from abluminal side. Because there are separate superfusion channels on

each side of the vein segment, the blood vessel can be activated by exposure to an inflammatory

mediator (e.g. TNF-α, LPS, fMLP) 62-65 on one side while the other side is superfused with a pure

buffer (e.g. MOPS). Then in situ immunofluorescence staining protocol can be performed for

visualizing the expression of respective cell adhesion molecules. Therefore, the activation and

control experiment can be achieved on one blood vessel in a homogeneous physical condition.

Another proposed application is to use the platform for creating controlled perfusion flowrates

and as a way to precisely regulate the applied shear stress to the blood vessel wall. This makes

the platform an ideal tool for investigating shear-dependent endothelium activation in molecular

level, and the related vascular disorders. This will be of relevance specifically in situations with

chronic venous diseases such as vasculitis and varicose veins.8, 66, 67

As we have previously mentioned an important characteristic of this platform is the possibility of

generating sheath flow within the blood vessel. This feature of the platform supports the

margination of leukocytes near the blood vessel wall which mimics in vivo leukocyte

margination during inflammation that promotes leukocyte- EC interaction.68, 69 Thereby, it

triggers the initiation of leukocyte extravasation from luminal flow to the abluminal

environment. Therefore another application of this platform is to quantitatively visualize

leukocyte extravasation cascade in vitro on an intact vein portion, and the effects of chemokines

on leukocyte rolling and adhesion can be studied. In order to achieve high-resolution cell-level

33

imaging with this platform, a microfluidic device designed on a thinner substrate than the current

microscope slide (i.e. ~1.5 mm) such as coverslip glass is preferred. This would decrease the

working distance, by increasing magnification and numerical aperture.

As it was mentioned before, one important feature of this platform is the ability to apply

controlled perfusion luminal flow. This can be simultaneously done by probing the functional

responses of the blood vessel as well as the expression of molecules and genes on the wall which

are caused by abluminal perfusion of specific drugs (through the superfusion channels). That

feature when coupled with the micro-scale volume of reagent consumption suggests that the

proposed platform can be a suitable candidate for applications used in drug developments.

34

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Appendices

A1. Experimental setup

Figure A1: Experimental setup. (A) Photograph and (B) schematic illustration of the vein-on-

a-chip platform consisted of: (1)Microfluidic device with manifold (2) Selector valve and

reservoirs (3) Syringe pumps (4) Temperature control system (5) Peristaltic pump (6) Vaccum

41

system for operating the bubble raps (7) Sterio microscope (8) Inverted microscope (9) CCD

camera (10) Computer

A2. Temperature control setup

The desired temperatures during different stages of experiment are achieved and controlled at the

inspection area of the chip as illustrated in Fig. A1. thermoelectric element (model TE-35- 0.6-

1.0, TE Technology) connected to a computer-controlled single mode temperature controller is

used to generate warm and cold temperatures of interest at the organ bath. A sapphire disk

(diameter: 25mm, thickness: 1mm, thermal conductivity: 35 W/m. K, model WSR-251, UQG

Optics, Cambridge, UK) is placed in between the TE element and the PDMS device using a

thermally conductive glue (thermal conductivity: 7.5 W/m. K) to produce a horizontally uniform

and constant temperature across the inspection area. To accurately generate and control

temperature at the inspection section, a thermistor (model MP-2444, TE Technology) was

embedded into the chip at the vicinity of vein segment location during the molding process of

PDMS. Heating (37 °C) and cooling (4 °C) is feedback controlled from thermistor. Temperature

control is automated in a custom software program (LabView, National Instruments, Austin, TX,

USA). In order to dissipate heat from the side of the TE element that is not in contact with the

PDMS device, a custom made cooling element is machined in aluminium and attached. The

cooling jacket is placed on top of the TE element and perfused with water at a rate of 30 mL/min,

using a peristaltic pump (Fisher Scientific 13-875- DIG. FH100 PUMP 14-4000ML/MIN EA).

Water perfusion produces an on-chip temperature change from 37 °C to 4 °C at a rate of 2

°C/min.

42

Figure A2: Temperature control setup. Schematic illustration of the automated temperature

control setup consisted of: a TE element glued to a sapphire dick which was placed on top of the

organ bath, a thermistor embedded in the PDMS device at the vicinity of the vein segment

(distance: 1.5 mm), a temperature controller connected to the TE element and the thermistor, a

cooling jacket which was placed on top of the TE element to dissipate heat produced during the

cooling mode operation, and a peristaltic pump connected to the cooling jacket for the water

perfusion.

A3. Dose Response Data

Figure A2 (A) and (B) represent chip-hosted vein segments, abluminally exposed to

phenylephrine (PE) and acetylcholine (Ach) through the superfusion channels at the top and the

bottom, respectively.

43

44

Figure A3: Dose response data. Chip-hosted mouse mesenteric vein segments (A) constricted

in response to incrementally increasing phenylephrine (PE) concentration, and (B) dilated in

response to incrementally increasing acetylcholine (Ach) concentration. Contrast is adjusted to

keep consistency in the images. Scale bars are 200μm.

A4. Staining Data

This section shows the images and respective intensity plots obtained from four stained samples.

45

46

Figure A4: Staining data. Orthographic projections and the respective intensity profiles of the

confocal images of four samples stained for CD31. Scale bars represent 50 µm for images A and

B, and 100 µm for images C and D.

47

A5. Bubble Formation Test

In order to investigate the source of bubble formation in the inspection area (i.e. where the vein segment

was held) during long run experiments (i.e. >12hours) a set of tests were performed. The experimental

platform was set up as follows: a PDMS device that contained of all the channel networks of the original

microfluidic device, except for the fixation channels, was used within the manifold. All of the fluidic

connections were implemented similar to the original vein-on-a-chip platform. Perfusion and superfusion

flow rates were set at 1 mL/hr. The bubble traps were connected to a miniature vacuum pump which was

operating throughout the experiment. The chip was incrementally heated up to 37 °C and images were

taken at a rate of 1fps for 16 hours from underneath using a CCD camera (model EXi Aqua,

QImaging, Surrey, BC, Canada) attached to a telecentric lens (zoom 7000, magnification: 18-108

mm, working distance: 127 mm, Navitar, Japan). Initially five main sources for the presence of

bubbles were assumed: a) heating the chip, b) roughness of the rings on the manifold which were used to

seal the fluidic inlets and outlets c) the small capacity of the bubble traps, d) gas diffusion through the

tubing and the syringes, and e) infusion of gas to the channels through the manifold inlets and connectors.

The assumptions were tested by running stepwise experiments. First, we performed the experiments at

room temperature and 37 °C, which rejected the first assumption as variation in the temperature did not

affect the bubble creation. In the next step, we polished the surface of the manifold that were in contact to

the PDMS for the sealing purposes. Then we changed the tubing material from plastic to PEEK and used

gas tight glass syringes instead of plastic ones. Next we sealed the manifold inlets by filling the gaps

between the tubing. After each step we repeated the experiment. Fig. A5 shows image data obtained from

the experiments. The results showed that the tubing and syringe material influence the bubble formation.

As well, polished surface and proper sealing between the manifold, the PDMS, and the tubing had effects

on the presence of bubbles in the microchannel. Fig. A5. F shows the number of bubbles presented in the

microchannels in each experiment plotted over time. The presence and absence of bubbles were indicated

as 1 and 0, respectively.

48

Figure A5: Bubble formation test. A and B) Images showing the presence of bubbles in the

superfusion and perfusion microchannels, respectively. Scale bar: 200 µm. C) Image of the chip

without bubble. Scale bar: 2 mm. D) Image of chip with bubbles in the superfusion inlet and the

bubble trap. Circles indicate the location of the bubbles inside the microchannels. Scale bar: 2

49

mm. E) Schematic of the manifold (left) and bottom surface of the connection inset (right). One

of the rings on the manifold which support the seal between the manifold and the PDMS device

is denoted by a circle. F) Bubble presence plotted over time for four experimental conditions as

follows: Test 1: before polishing, tygon tubing, plastic syringe, room temperature. Test 2: before

polishing, tygon tubing, plastic syringe, 37 °C. Test 3: before polishing, peek tubing, plastic

syringe. Test 4: after polishing, peek tubing, gastight syringe.