Development and Testing of a Microfluidic Device for Studying Resistance Artery Function

DEVELOPMENT AND TESTING OF A MICROFLUIDIC DEVICE FOR STUDYING RESISTANCE ARTERY FUNCTION

Andrei Vagaon

A thesis submitted in conformity with the requirements for the degree of Master of Science

Graduate Department of Physiology

University of Toronto

MSc thesis, 2010, Andrei Vagaon, Department of Physiology at the University of Toronto

ABSTRACT

Introduction: Hypertension is the number one risk factor for cardiovascular diseases. Total

peripheral resistance (TPR) is strongly involved in blood pressure homeostasis. TPR is primarily

determined by resistance arteries (RAs). Pathogenic factors which change RA structure are

associated with cardiovascular disease. Despite this, methods employed in the study of RAs lack

efficiency.

Methods: A polymer microfluidic device (Artery‐on‐a‐Chip Device, AoC) made from

polydimethylsiloxane (PDMS) was developed. RAs from CD1 mice were measured on the

device. Their responses to phenylephrine (PE), acetylcholine (Ach), FURA‐2 imaging, and 24‐h culture were assessed.

Results: Following several modifications, vessel function on the AoC device was successfully

measured. Robust PE constriction and Ach‐induced vasodilation were observed. AoC arteries

were viable after 24‐hour culture, and FURA‐2 was successfully imaged.

Conclusions: The AoC device is a viable alternative to cannulation myography. The AoC can

greatly increase the efficiency of RA studies, while also decreasing training time and difficulty.

TABLE OF CONTENTS

Chapter 1. INTRODUCTION ...... 1

1.a Clinical background ...... 1

1.b Factors involved in resistance artery (RA) tone regulation ...... 3

1.c Current techniques employed in RA research ...... 4

1.d Difficulties in the study of RAs ...... 6

1.e Microfluidic devices ...... 10

Chapter 2. MATERIALS AND METHODS ...... 14

2.a Solutions and substances ...... 14

2.b Vessel isolation and preparation ...... 15

2.c Vessel loading onto the AoC device ...... 15

2.d AoC device fabrication ...... 16

2.e Vessel fixation onto the AoC device ...... 17

2.f Substance delivery to the AoC device ...... 18

2.g Diameter measurement and imaging of the arteries loaded onto the AoC ...... 18

2.h Statistics and diameter change measurements ...... 19

Chapter 3. RESULTS ...... 21

3 Vessel loading ...... 21

3.a Vessel fixation ...... 21

3.a.1 Vetbond tissue adhesive – complete bonding ...... 22

3.a.2 Vetbond tissue adhesive – partial bonding ...... 24

3.a.3 Vessel fixation via negative hydrostatic pressure (NHP) ...... 27

3.b The vessel organ bath, substance delivery and removal ...... 29

iii

3.b.1 Organ bath with flow over the vessel (design 1) ...... 29

3.b.2 Organ bath with flow over the vessel (design 2) ...... 31

3.b.3 Organ bath with flow on the vessel lateral surfaces, NHP fixation ...... 32

3.c The mixing cell ...... 34

3.d Separation of the perfusion and superfusion streams ...... 36

3.e Phenylephrine dose response curves (DRCs) on the AoC device ...... 37

3.f Acetylcholine dose response curves on the AoC device ...... 40

3.g Cultured vessels on the AoC device ...... 41

3.h One‐sided PE constrictions ...... 42

3.i FURA‐2 on‐chip measurements ...... 43

Chapter 4. DISCUSSION ...... 45

PDMS – AoC fabrication material ...... 45

4.a Vessel fixation and pressurization ...... 46

4.b The organ bath ...... 50

4.c The on‐chip mixing cell ...... 55

4.d Phenylephrine and Acetylcholine dose response curves ...... 56

4.e Cultured vessels on the AoC device ...... 59

4.f Novel experiments using the AoC device ...... 60

Chapter 5. CONCLUSIONS ...... 62

Future directions ...... 67

5.a Basic science ...... 67

5.b Clinical applications ...... 68

Chapter 6. REFERENCES ...... 69

LIST OF ABBREVIATIONS

Ach acetylcholine

AoC artery‐on‐a‐chip device

CAM calmodulin cAMP cyclic adenosine monophosphate cGMP cyclic guanosine monophosphate

CO cardiac output

DRC dose response curves

ECs endothelial cells

EDHF endothelium‐derived hyperpolarizing factor

LOC lab‐on‐a‐chip

MLCK myosin light chain kinase

NHP negative hydrostatic pressure

NO nitric oxide

NOS nitric oxide synthase

PDMS polydimethylsiloxane

PE phenylephrine

PGH prostaglandin H

PGI2 prostacyclin

PKG protein kinase G

PLC phospholipase C

RAs resistance arteries

SMCs smooth muscle cells

TPR total peripheral resistance

LIST OF IMAGES AND FIGURES

Image 1: Chip fabrication by soft lithography ...... 11

Image 2: Representative image of the AoC inspection area ...... 16

‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐

Figure 1: The V1 chip design: VetbondTM tissue adhesive – complete bonding ...... 22

Figure 2: The V2 chip design: VetbondTM tissue adhesive – partial bonding ...... …… 24

Figure 3: The V2 chip design: test ...... 26

Figure 4: The S1 chip design: Vessel fixation via Negative Hydrostatic Pressure ...... 27

Figure 5: The BM 1.0 chip design: Organ bath with flow over the vessel ...... 29

Figure 6: The SY 1.1 chip design: Organ bath with flow over the vessel (design 2) ...... 31

Figure 7: The AoC 1.0 design: Organ bath with flow on vascular lateral sides and NHP .... 32

Figure 8: The on chip mixing cell used in AoC devices ...... 34

Figure 9: Organ bath washout, and separation of the perfusion and superfusion ...... 36

Figure 10: AoC arteries: PE dose response curves ...... 38

Figure 11: AoC arteries: Ach dose response curves ...... 40

Figure 12: AoC arteries: 24‐h cultured vessel ...... 41

Figure 13: AoC arteries: One‐sided PE constriction curves ...... 42

Figure 14: FURA‐2 imaging experiments on the AoC ...... 43

Figure 15: Flowchart of the AoC development process ...... 65

Figure 16: Final version of the AoC device ...... 66

1. Introduction

1.a Clinical background

Hypertension (high blood pressure) is regarded as the number one risk factor for cardiovascular

diseases1, which are considered to be the primary cause of death in the North American population2. Blood pressure homeostasis is primarily regulated through two mechanisms:

cardiac output (CO) and total peripheral resistance (TPR). While diseases of the heart, and their

contributions to hypertension, are intensively researched (through methods such as Doppler

echocardiography3, thermodilutions4, or the Langendorf perfused heart preparations5),

mechanisms that modulate TPR (the sum of the resistance of all peripheral vasculature) need to be better examined.

TPR is largely determined by resistance arteries (RAs). These resistance vessels are highly specialized structures, with diameters ranging from 30 µm to 300 µm, located in end sections of the arterial vascular tree6. The luminal side is composed of one layer of endothelial cells (ECs)

which are enveloped by several layers of circumferentially oriented smooth muscle cells

(SMCs), which can undergo rapid contraction or relaxation to regulate tissue blood flow7. As

resistance is inversely proportional to the radius^4, explained by the Hagen‐Poiseullie equation,

ΔP = 128µLQ/Πd4, this means that a change in radius will exponentially alter the resistance to

flow in end‐tree vasculature8,9. Consequently, resistance arteries have mechanisms which

enable them to adapt to changes in blood pressure, in order to maintain organ perfusion at constant levels. An example of one such mechanism is the myogenic response, also known as

1 the Bayliss effect10,11,. In this response, the vascular smooth muscle cells, within resistance

arteries, respond to stretch generated as a result of increased pressure. As the muscle

membrane is distended, stretch‐activated ion channels open and vasoconstriction ensues.

While this response maintains organ perfusion homeostasis, and protects sensitive capillary

beds, the decrease in diameter elevates TPR, resulting in an overall increase in blood pressure.

12 Over time this leads to, or exacerbates, a hypertensive state . This specific effect is largely

mediated at the level of small resistance arteries. Overall, it serves as an example of the importance of arterial diameter, and the implication of resistance arteries in hypertension.

The individual capacity for each resistance artery to develop tone increases as its diameter decreases13. As a result, arteries and arterioles with diameters ranging from 10 to 200 µm are

the principal vessels involved in controlling resistance, and therefore arterial blood pressure.

These vessels are highly innervated by autonomic nerves and respond to changes in nerve

activity, circulating hormones (i.e. vasopressin), or endogenously released factors in either

constriction or dilation14. As such, tone, defined as amount of constriction relative to the maximal arterial diameter, is established through such diffusible neurotransmitters as noradrenaline, which promote vasoconstriction, as well as vasodilatory factors from

endogenous endothelial cells, such as prostacyclin15 and nitric oxide16. These factors are

released in response to stimuli such as fluid‐induced shear stress, pressure, strain17,18, and

chemical stimuli (i.e. prostacyclin, nitric oxide, and endothelium‐derived hyperpolarizing factor19). Therefore, vascular tone is the net result of these multiple inputs acting on resistance

arteries in a highly controlled and dynamic microenvironment. Subsequently, changes in the

structure and function of resistance arteries are important pathogenic factors which both promote and sustain cardiovascular disease.

1.b Factors involved in RA tone regulation

Two factors which regulate vascular tone are often used in resistance artery research: phenylephrine and acetylcholine. In resistance arteries, PE promotes contraction of the smooth muscle cells, and subsequently vasoconstriction. It is primarily an α1‐adrenergic receptor

agonist. α1 Receptors are members of the G‐protein coupled receptor family, and are associated with Gq. Upon PE binding, the Gq receptor activates phospholipase C (PLC), which

2+ 2+ subsequently leads to an increase in IP3 and Ca . This leads to an overall increase in Ca , which

binds to calmodulin (CAM), and then activates myosin light chain kinase (MLCK). MLCK

phosphorylates the myosin light chain at residue 19, enabling the myosin to bind actin, leading

to crossbridge formation6,7. The overall result is vasoconstriction.

Ach, another neurotransmitter commonly used in vascular research, promotes vasodilation in end‐tree vasculature resistance arteries. Ach primarily acts by increasing cellular levels of prostacyclin, endothelium‐derived hyperpolarizing factor (EDHF), and nitric oxide (NO). These factors increase in different ratios in various vascular beds. Prostacyclin (PGI2) is produced from

prostaglandin H (PGH) in endothelial cells. PGI2 binds to its receptors, increasing cytosolic cyclic

adenosine monophosphate (cAMP) levels. cAMP then activates protein kinase A (PKA), and

subsequently inhibits MLCK, causing vasodilation19.

A second mechanism of vasodilation is EDHF. This term however encompasses a group of yet‐

undetermined vasodilatory factors. EDHF came about as a result of experiments which reported

that when prostacyclin and NO were inhibited there was still a factor causing vasodilation19.

The third mechanism of Ach concerns the activation of nitric oxide synthase (NOS), and subsequent increase in NO. This in turn causes an increase in cyclic guanosine monophosphate

(cGMP) causing an increase in the activity of protein kinase G (PKG). The overall effect is either

a reduction in the activity of MLCK, or an increase in MLCP via a decrease in Rho‐associated protein kinase (ROCK), causing vasodilation6,14,15,16. Due to the fact that their pathways are fairly

well defined, both PE and Ach are often used as ‘gold standards’ to test resistance artery

experimental viability in cardiovascular research. It is important to note that both of these

neurotransmitters focus on diameter regulation, as it is the primary mechanism through which

resistance arteries regulate blood flow.

Despite its obvious importance, small resistance artery research is often impeded by limitations in the experimental setups used for these studies. Current methods are expensive, slow, training intensive, highly inflexible, lack scalability, and most importantly do not permit a microenvironment control, which would appropriately mimic in‐vivo conditions. Presently,

there are two methods widely employed in the study of isolated intact resistance arteries: wire myography and pressure myography.

1.c Current techniques employed in RA research

The wire myograph was first developed in 1972 by Bevan and Osher20. In this method, vessels

up to 5mm in length are isolated and thin platinum wires threaded through the vessel lumen and attached to force transducers in order to measure vascular responses. Although this method has been greatly improved upon since development, and measurement sensitivity has greatly increased, the basic concept remains the same21. In the most widely used wire

myography system, vascular responses are measured as the force necessary to counteract a

force exerted by a resistance artery against wires attached to the transducer and kept at a fixed

distance. As there is no change in the length or shape of the vessel, as well as no change in the

distance between the wires, the artery is considered to be measured under isometric (constant

length) conditions22. This enables the precise and standardized measurement of forces

developed by isolated resistance arteries prepared either as intact vessels, strips, or ring preparations23,24. The wire myography setup however does not accurately reflect the in‐vivo environment. Due to its design, the setup does not allow for the separate treatment and/or analysis of the intra and extralumenal compartments. As a result, spontaneous constriction does not occur until a force equivalent to 60‐100 mmHg of intralumenal pressure is applied and, more importantly, myogenic responses are not observed, even when the vessels are subjected to forces equivalent to baseline in‐vivo pressures25,26. Furthermore, the setup allows

mechanical forces within the artery to act in only one direction27. Vessel constriction and relaxation in a myograph thus take place primarily in the radial direction, unlike in‐vivo

conditions where this happens in the circumferential direction23. Lastly, the general

morphology and conformational structure under which the artery is being measured can be different than in‐vivo28.

In order to enable the study of isolated resistance arteries under conditions closer to those

observed in‐vivo, an alternative method to wire myography, pressure myography, was

developed. A pressure myography setup uses two cannulas (usually made out of glass) on

which the vessel is pulled on and held in place using sutures. Experimental results are shown directly as changes in resistance artery diameter, as recorded with a CCD camera and measured using edge‐detection methods (video myography). On a pressure myograph buffer solutions are readily perfusable into the vessel lumen, and intralumenal pressure can then be established using either a liquid column (hydrostatic pressure) or a servo‐controlled peristaltic pump29,30. As

a result, these cannulation setups are able to maintain isobaric (constant pressure) conditions

in the vessel lumen, and more closely mimic the in‐vivo environment of the resistance artery being studied. Myogenic responses are readily observed on the cannulation setup at all physiological pressure levels31, with constriction taking place primarily in the circumferential

rather than the radial plane23. As a result, various experiments have shown that resistance

arteries studied on pressure myographs are significantly more sensitive to stimuli than when studied using wire myography.22,23,26,27,32. The EC‐50 concentration of PE and Ach needed to

elicit a vascular response of similar magnitude was 4‐5 times lower in the pressure myograph

compared to the wire myograph33.

1.d Difficulties in the study of RAs

Despite the notable advancement represented by employing pressure myography in resistance

artery studies, several disadvantages are associated with this technique. Custom‐made

mechanical setups used to cannulate the vessels are expensive (costing upwards of $5000

each), deteriorate over time (requiring maintenance), and have very limited abilities for

integration into other analysis systems due to their weight and bulky composition (as even the

smaller custom‐made micromanipulators, used to position the pipettes, are large in size and

need to be attached to a stable base). Commercially available pressure myography systems, such as the latest DMT models, 202‐CM and 204‐CM34, are even bulkier and more difficult to

integrate into custom applications, and are often prohibitively expensive, with costs of $40 000

per system. Although companies such as DMT, and organizations such as the Consortium for

Integrative Cardiovascular Research, have made numerous improvements to pressure myography setups, one remaining constant is the target diameter of the resistance arteries

being studied. Due to various limitations, arteries with a diameter notably smaller than 60 µm

are very difficult to study on current pressure myographs, as the design of the setups is not

scalable. As a result, vessels cannulated on pressure myography setups come from a limited

number of vascular beds. This is a serious limitation, as smaller distal resistance arteries, which

are potentially the primary players in blood pressure regulation37, cannot be studied on

presently existing setups. While arteries as small as 35 µm have been cannulated and pressurized in our laboratory, on custom made setups, the success rate of these cannulations has been very low. More so, the technical skills required to cannulate such small vessels are extensive and difficult to acquire.

Alongside accessibility and flexibility challenges, there are also numerous practical obstacles in pressure myography. The organ bath where the vessel is being studied has a minimum volume

of 2 mL and is prone to evaporation, requiring extensive substance volumes to be used over the course of a long experiment, or whenever a dose or solution is replaced. As it is not desirable

for a resistance artery to be exposed to air, in order to maintain a constant environment, new

solutions are often mixed with a determined amount of a previous solution whenever dose

response curves are performed. As such, replacement of a substance that the vessel is exposed

to takes place by strongly diluting it (through repeated washes) rather than by performing a complete washout. Another mechanical limitation of cannulation setups is that, over time,

waste products need to be removed and the nutrients in the organ bath need to be

replenished, as the solution bathing the vessel is stagnant. As these washes need to be ideally

performed every 15‐20 minutes, based on results observed in our laboratory, the vessel

measurement process continuously requires user input. While the newest available pressure

myography model from DMT, the 204‐CM myograph, is equipped with the ability to

automatically washout the vessel chamber superfusion area (enabling long‐term vessel culture

experiments), it is a costly setup and does not address the aforementioned problems of large

chamber volumes and ‘imperfect’ washes. The minimum volume of the 204‐CM organ bath is

still large, at 2 mL, and residues still accumulate at the bottom of the bath due to its cup‐like

design. Therefore, as a tight control of the microvascular environment is intimately linked with

proper arterial function, these disadvantages limit the research flexibility of even the newest

commercially available pressure myographs.

Further impeding the widespread use of pressure myographs as vascular research tools is that,

alongside the aforementioned financial, mechanical, and experimental limitations involved with

using the cannulation setups, there are multiple end‐user limitations. Based on observations

made in our laboratory, an average user requires 2‐3 months of training to cannulate vessels on

the pressure myography setup while still maintaining intact smooth muscle function (evaluated

35 as percent constriction of the vessel to phenylephrine, an α1‐adrenergic receptor agonist ) and while also maintaining intact endothelial function (evaluated as percent dilation of the vessel by exposure to acetylcholine, a muscarinic receptor agonist36). Even when a high success rate

(~80%) of preserving vascular function is attained, the user continues to improve their techniques over the course of a project, adding a degree of variability. Although quality control methods are put in place (such as excluding arteries that exhibit less than 30% constriction of

maximum diameter at 3.0 µM PE, or less than 40% reversal of a 1.0 µM PE constriction when

exposed to 10.0 µM Ach) these substances cannot test for the proper function of all

physiological responses within a resistance artery. As a result, due to the fact that there is no

clear way to set a particular baseline in the skills of the user, it can often be difficult to directly

compare current results with those obtained when the user was newly introduced to the system without accounting for this variable. More importantly, it can often be difficult to compare results obtained from different users within the same laboratory, and especially

between different laboratories, as there is currently no set standard for cannulation procedures and techniques that every pressure myograph user needs to adhere to. As a result, standardization of results and experimental reproducibility are often difficult.

1.e Microfluidic devices

In order to overcome the limitations of pressure myographs, and to increase the efficiency of microvascular research and advance the field, it is desirable that a new device which improves

upon the current systems should be developed. The new device would have to be highly cost

efficient, scalable, flexible, easy to integrate with other research tools, user friendly, and

simplify the implementation of experimental protocols used to study resistance arteries. The aim of my thesis project was to develop such a novel interface, a prototypic microfluidic tool:

the AoC (Artery‐on‐a‐Chip) device. The field of microfluidics involves the precise manipulation

of sub‐milliliter quantities of fluids within a carefully controlled environment. Such “lab‐on‐a‐ chip” (LOC) devices scale down several laboratory functions onto a chip, made of various polymers, and measure only a few square centimeters in size37. LOC devices have been

successfully used in various biological applications, such as electrophoresis studies, cell culturing, and rapid cell counting and sorting38.

Although microfluidic devices are fabricated from a wide array of materials, PDMS

(polydimethylsiloxane) is often the material of choice due to advantages conferred by its

unique properties. PDMS chips are fabricated quickly (going from the design phase to the quality control phase within 24 hours) and are relatively inexpensive compared to other

materials such as glass. Furthermore, PDMS is transparent, making it ideal for many imaging techniques employed in the study of resistance arteries. The material is also non‐toxic to cells

and has a low thermal conductivity of 0.2 W/(m.K). While the low conductivity means a PDMS

device requires 4‐5 minutes to reach body temperature when heated, this ensures that the

interior environment is kept at a fairly constant temperature, as very little heat is dissipated to

the external environment37. PDMS also easily binds to many surfaces, such as glass, through plasma treatment which oxidizes the surface and alters its chemistry to allow for bonding

without changing the properties or shape of the design39.

The PDMS chips used in this study were fabricated using rapid prototyping soft lithography

methods (Image 1).

Image 1. Chip fabrication by soft lithography. The initial CAD design is printed on a high resolution transparency, creating the photomask, which is then used in contact photolithography by spin‐coating two layers of photoresist (and by exposing it to UV light and developer) to yield a master. PDMS was then molded onto the master and bonded to a glass microscope slide.

In this process, a design created in a CAD (computer assisted design) program is printed on a high‐resolution transparency, creating a photomask40, which is then used in contact photolithography to yield a master. The master is subsequently molded with PDMS, and the mold is then plasma treated, and bound to standard glass microscope slides. This yields a device bound by glass on the bottom surface and PDMS on the top, with a network of channels

enclosed within the two layers. Fluidic connections are then attached to the completed device

to facilitate the addition and removal of various substances to and from the organ bath where

the vessels are measured.

The aims of developing the AoC device are to (i) design and fabricate a chip which would permit

the reversible loading and mounting of resistance arteries, and (ii) transfer all current resistance artery diameter and calcium measuring techniques onto the AoC, while attempting to develop novel protocols for vascular studies which cannot be carried out using the standard pressure myography cannulation setup.

The AoC technology must successfully complete experimental protocols while also maintaining

complete vascular integrity (defined by smooth muscle and endothelial function as previously described under quality control). The design needs to be such that resistance arteries can be loaded onto the chip easily, and to the same standard, by users with varying degrees of

experience. After loading, the device would need to be able to maintain the resistance artery in

place, and allow it to be exposed to a wide range of intralumenal pressures, over extended

periods of time. Substance delivery and removal to the organ bath where the vessel is being studied needs to be such that it improves upon the current cannulation setup by carrying out faster washouts without having any leftover ‘residues’ from the previous substance used.

Lastly, the AoC design should adequately separate the perfusion and superfusion streams.

The AoC project was developed in collaboration with members of Dr. Axel Guenther’s MIE

laboratory at the University of Toronto. While I provided feedback and suggestions regarding

the chip designs, and while I conducted the on‐chip experiments in this project and was involved in the design process, fabrication of the PDMS devices was exclusively handled by the

members of the Guenther lab, according to the specifications jointly agreed upon. The overall

project was a joint effort, with weekly meetings taking place over the duration of the project,

which served as a forum for updates and the exchange of new ideas. For the purposes of this

experiment it was hypothesized that resistance arteries can be successfully held in place and

measured on the AoC device.

2. Materials and Methods

2.a Solutions and Substances

All of the vessels tested were isolated in pH 7.4 MOPS‐buffered salt solution containing

[mmol/L]: NaCl 145, KCl 4.7, CaCl2 3.0, MgSO4•7H2O 1.17, NaH2‐PO4•2H2O 1.2, pyruvate 2.0,

EDTA 0.02, MOPS (3‐morpholinopropanesulfonic acid) 3.0, and glucose 5.0, kept at 4oC for vessel isolation and 37oC for functional measurements. Vessels were fixated in place using 3M

Vetbond Tissue Adhesive 1469SB (3M Innovations – kindly provided by the University Of

Toronto Division Of Comparative Medicine). SU‐8 2050 and SU‐8 25 photoresist, both obtained

from Microchem, were used in AoC device fabrication, along with SU‐8 developer and polydimethylsiloxane (PDMS) (also obtained from Microchem). The culture medium used in culture experiments consisted of Leibovitz medium (L15), containing 20,000 U/l penicillin and

20 mg/l streptomycin, and supplemented with 15% heat inactivated newborn calf serum

41 (Invitrogen), as previously described . Phenylephrine (PE) or Acetylcholine (Ach), employed in functional experiments to test smooth muscle and endothelial integrity, were obtained from

Sigma‐Aldrich and prepared and diluted to the necessary stock concentrations, 3 µM PE and

10µM Ach respectively. The FURA‐2 AM (from Molecular Probes), used to test the optical properties of the AoC device, was dissolved in DMSO, stored as a 1mM stock solution in 10‐µL

aliquots, and diluted to a final concentration of 2 mM, as previously described41.

2.b Vessel isolation and preparation

Resistance artery segments used on the AoC device were isolated from CD1 mouse (Charles

River) from the second and third‐order branches off the superior mesenteric artery by careful

microdissection (using a Leica MZ16 stereomicroscope with a Planapo 1.0X lens), and the rough

connective tissue was removed. Throughout all the experiments arteries were isolated and kept

in MOPS‐buffered salt solution during isolation and at all points prior to measurement.

2.c Vessel loading onto the AoC device

Despite various design changes in AoC devices throughout the duration of the study, the method of vessel loading remained basically the same in all iterations tested. Vessels were

loaded onto the AoC device using suction applied through a microfluidic channel which was

perpendicular to the channels leading into and out of the organ bath/inspection area (where vessel diameter was measured, Image 2). The suction was therefore applied longitudinally,

parallel to the longitudinal axis of the vessel. The lumen of all of the arteries loaded onto the

AoC device was opened on both ends prior to loading, in order to prevent the vessel from

collapsing due to the suction pressure applied. The loading channel underwent significant

narrowing halfway throughout its length, in all AoC designs, at the point where the artery was

supposed to stop in order for functional measurements to be performed.

Image 2. Representative drawing (not to scale) of the AoC inspection area, seen from above. The vessels are loaded through the bottom channel by applying suction in the direction of the arrow. Arteries are then fixated in place and exposed to various substances via the lateral channels. The inspection bath is enclosed between a glass microscopy slide and a PDMS layer.

2.d AoC Device Fabrication

Although fabrication was carried out by members of Axel Guenther’s MIE laboratory, at the

University of Toronto, a brief overview of the fabrication process shall be provided. The PDMS

chips were fabricated using rapid prototyping soft lithography methods. In this process, designs

created in a CAD program were printed on high‐resolution transparencies with a 10 µm

resolution by CAD/ART Services Inc., creating a photomask. The photomasks were then used in

contact photolithography, to make the masters, by spin‐coating two layers of SU‐8 2050 photoresist from Microchem at 1800 rpm on top of one seed layer of SU‐8 25 spun at 2000rpm,

giving a total feature height of approximately 150 microns. The product was then exposed with

38 milliwatts/cm2 UV light at 365 nm for 6.3 seconds, and developed with SU‐8 developer

(Microchem) to yield a master, which was subsequently molded with (PDMS). The mold was then removed from the master, plasma treated, and bound to a standard glass 2” X 3” microscope slide. The device interface was then made using 0.5 inch long, type 304 stainless steel 23 gauge pins (New England Small Pin Corp). Tygon S‐54‐HL tubing (Upchurch Scientific) with inner diameters of 1/32” and 1/16” was attached to the pins in order to complete the device interface and to facilitate the addition and removal of various substances to and from the AoC device. Chip depths and feature sizes were subsequently measured (to confirm the

accuracy of fabrication) by members of the laboratory of Dr. Axel Guenther (University of

Toronto – MIE) using a profilometer, which provided measurements with an accuracy of ± 2 µm in the vertical plane.

2.e Vessel Fixation onto the AoC Device

Vessels were fixated in place using two methods, Vetbond and negative hydrostatic pressure

(NHP) fixation. In the first method, 1 mL of Vetbond was delivered to the AoC device at a flow

rate of 4 mL/h using Aitecs 2015 syringe pumps (Viltechmeda, Lithuania). In the second method

vessels were fixated by suction, applied at specific points to the vessel wall, generated using negative hydrostatic pressure. BD 60 mL Luer‐Lok Tip syringes connected to Tygon 1/16” diameter tubing, filled with MOPS, and lowered to an equivalent ‐45 mmHg of pressure

(relative to the height of the AoC device) were used to generate the necessary NHP.

2.f Substance Delivery to the AoC Device

Fluids delivered to vessels were perfused using either Aitecs 2015 syringe pumps (for the PE dose response curves) or the more sensitive Harvard Apparatus (U.S.A.) MRI 70‐2131 syringe pumps (for the Ach dose response measurements). BD 5mL and 10mL Luer‐Lok Tip syringes

were loaded onto the syringe pumps and used to deliver substances to the AoC device through

Tygon® Formulation S‐54‐HL tubing with a diameter of 1/32 inches.

In 24h vessel culture experiments the vessels were perfused with Leibovitz L15 culture medium

through the lumen (perfusion) and on the outside of the vessel (superfusion) continuously at a

flow rate of 1mL/h using the aforementioned Aitecs 2015 syringe pumps.

2.g Diameter Measurement and Imaging of Arteries Loaded on the AoC Device

All vessel diameter measurements were conducted at body temperature. The arteries were

warmed up to 37oC (using a Linkam Scientific Instruments MC60 heating plate) in approximately

4oC steps (5 minutes/step) over the course of 20 minutes, and while hydrostatically setting the

intralumenal pressure to 45 mmHg.

Functional measurements were conducted either by exposing the vessels to Phenylephrine (PE)

or Acetylcholine (Ach) obtained from Sigma‐Aldrich and prepared and diluted to the necessary

stock concentrations, 3 µM PE and 10µM Ach respectively. All functional measurements were

conducted with vessels pressurized to 45 mmHg, and with no flow through the lumen.

Cannulation setup experiments were performed using a custom pressure myography setup

developed internally in the lab of Dr. S.S. Bolz, and the vessels were pressurized using a hydrostatic column; diameter data was read and interpreted using raster‐line edge‐detection equipment (Crescent Technologies, Windsor, ON, Canada).

Functional responses of vessel diameter in the AoC were measured on a Nikon Eclipse E600

microscope (10X/0.3 PlanFluor objective, DIC L/N1 ∞/0.17 WD 16) which had a Panasonic CCTV

WV‐1410 Camera attached. The camera was connected to a Picolo Euresys capture card, which

recorded the measurements and delivered them for processing to the visual display and

diameter analysis program Diamtrak3+ (version 3.5, ©T.O. Neild, 2000,2003). High resolution

vessel images were captured using a Pixelink Capture 6.0 Mpixel camera. The images were then

analyzed either in SimplePCI (version 5.3, C‐Imaging Systems), for high resolution fluorescent

images (obtained using Fluorescein and Rhodamine dyes) or in ImageJ (version 1.41, N.I.H.‐

U.S.A.) for one‐sided PE constriction vessel measurements. For one‐sided responses

constriction was measured as wall displacement relative to the vessel wall position in MOPS

buffer.

FURA‐2 images were obtained using a Photometrics QuantEM 512SC camera connected to a

Zeiss Observer.Z1 microscope (20XPlan‐NeoFluar objective) setup by 3I (Intelligent Imaging

Innovations).

2.h Statistics and Diameter Change Measurements

o Vessel tone is calculated as percent of the maximal diameter (Diamax) recorded at 37 C in MOPS

solution individually for each vessel. For Ach experiments percent dilation is the percent reversal of a 1 µM PE initial constriction. For one sided PE constriction experiments all changes

in diameter, and vessel wall displacement, are reported by using the edges of the non‐ constricted vessel, in MOPS solution at 37o C, as a set point of reference. Data shown

represents means ± SE, for n‐values, where “n” represents the number of resistance arteries investigated. An unpaired Student’s t‐test was used to examine PE constriction and Ach dilations in vessels cultured on the AoC device, to indicate a significant Ach dilation. Diameter

curves for one‐sided PE constrictions were analyzed using ANOVA, to indicate diameter

differences between the two vessel walls. Differences were interpreted as significant at P <

0.05. The data were analyzed using Graph Pad Prism 5, with the abovementioned parameters.

3. Results

Vessel loading

Arteries were reversibly loaded onto the AoC device through a microfluidic channel, that was open to the outside MOPS‐filled loading bath, and on which suction was applied parallel to the longitudinal axis of the vessel, in order to draw it into the device (Figure 1). This method of

loading was used successfully on all AoC devices. Vessel loading on the AoC device was

significantly faster than vessel cannulation on glass pipettes. As a noteworthy observation,

users that tested the AoC device for the first time were able to successfully load vessels within

the same day.

3.a Vessel Fixation

In order to be properly pressurized, and for their diameter to be measured under in‐vivo like conditions, resistance arteries first had to be fixated in place on the AoC device. Two methods

were considered for this purpose: Vetbond tissue adhesive and negative hydrostatic pressure

(NHP). 3M Vetbond polymerizes in seconds when it comes in contact with tissue and fluids42, provides a strong hold, and it is non‐toxic, readily available and fairly cost‐efficient, and its blue colour enables easy tracking of delivery in a clear device, such as the AoC. Although Vetbond had numerous advantages, it was ultimately unusable in the AoC device. Due to its binding method, upon coming in contact with fluids, Vetbond desiccated and strongly damaged the

vessel beyond repair upon reaching the vessel surface (n=23). Attempts to dilute the Vetbond

proved unsuccessful. Modifications in chip design, aimed at reducing the amount of Vetbond

reaching the vessel walls, also proved unsuccessful. A subsequent development then focused on fixation using suction applied on the vessel walls via negative hydrostatic pressure (NHP)

(n=17).

3.a.1 VetbondTM Tissue Adhesive – Complete Bonding

In an attempt to fixate the vessel in place inside the AoC device, to enable pressurization,

Vetbond from 3M was used in a modified version of the AoC device (Figure 1).

b a

Figure 1. The V1 chip design. The vessel, represented by a contoured outline, was loaded onto the device with negative pressure applied in the direction of the arrow. Vetbond was delivered around the vessel, after loading occurred, from channel (a); any excess amount was removed through channel (b).

The Vetbond‐1 (V1) chip consisted of a simple single layer design, with four main channels. The

loading channel, which ran vertically and contained the vessel, had a diameter of 200 microns.

The pressurization channel, indicated by the arrow in Figure 1, had a diameter of 80 microns.

Channels (a) and (b), used to deliver the Vetbond solution, had a diameter of 150 microns.

Channel (a) was a composite channel, with three separate entry ports if necessary, in order to

enable the dilution of the Vetbond substance as needed at the point of contact with the vessel.

The depth throughout the chip was uniform for each chip, and chips of 100, 150, and 220

microns depth were tested.

The chip was flushed with and submersed in MOPS buffer at room temperature (24oC). Second

and third order mesenteric CD1 resistance arteries were isolated and loaded onto the chip.

Vessels were loaded through the bottom channel, by using suction applied via a syringe in the

direction indicated by the arrow in Figure 1, and positioned as shown by the representative

vessel outline. Vetbond was applied by syringe pump infusion from the rightmost channel, (a),

over and under the vessel, and out through channel (b). A flow rate of less than 3.5 mL/h in the

syringe pumps resulted in the Vetbond bonding to the channels before reaching the vessel

surface, and a flow rate of 4.0 mL/h was used as a result.

In the V1 chips with depths of 100 microns (n=6) and 150 microns (n=5) the Vetbond was

unsuccessful in crossing over the vessel and exiting through channel (b). The Vetbond delivered

in these chips exited through the bottom channel (used to load the vessel) upon making contact

with the vessel surface. In the chips of 220 microns in depth (n=7) Vetbond appeared to successfully flow over and under the vessel and out through channel (b). In these experiments

however the vessel was quickly desiccated and broken down upon contact with the Vetbond on

all sides. This created a solid polymer plug where the vessel was, between channels (a) and (b) and blocked off the channel. Attempts to dilute the Vetbond before reaching the vessel surface

(by delivering Vetbond through the top and bottom of channel (a) and MOPS through the

middle of channel (a)) were unsuccessful, as it would polymerize at the exit point of channel (a), before making contact with the vessel surface. Vetbond was unsuccessful at fixating the vessel

in place in the V1 chip design.

3.a.2 VetbondTM Tissue Adhesive – Partial Bonding

Partial bonding experiments were conducted using a modified version, the V2 design (Figure 2). i ii

a a

b b

Figure 2. The V2 chip design. Suction was applied in the direction of the arrow (i) to load the vessel on the AoC device. Trypan blue was successfully perfused through channels (a), from a common reservoir, and out through channels (b) for removal from the vessel surface. No spilling into any other channels seemed to occur. Figureb 2‐ii shows a vessel successfully loaded onto the V2 chip.

The V2 chip design consisted of single layer chips, with 6 channels. Channels (a) and (b) had a

diameter of ~50 microns. The loading channel had a diameter of 200 microns which narrowed

to 80 microns at the adhesion point where all the channels met. The suction/pressurization

channel, indicated by the arrow, had a diameter of 40 microns at the point where the loaded

vessel stopped, and then gradually increased to a diameter of 100 microns, in the direction of

the arrow. The depth was uniform throughout the chip, at 150 microns.

The vessel was loaded in the same fashion as the V1 chip, through the bottom vertical channel

of the chips, using pressure applied via syringe in the direction of the arrow shown in Figure 2i.

Vetbond was delivered through the channels labeled (a) in Figure 2i, which were connected to a

joining port and a single syringe outside the chip, at a flow rate of 4mL/h. The Vetbond exited

the chip via channels labeled (b) (which also joined to a single port and one syringe outside the

chip) using a syringe pump suctioning at a matching rate of 4 mL/h. Experiments using Trypan

blue, as shown in Figure 2i, indicated that at a flow rate of 4mL/h the dye successfully flowed in and out of the chip without visible spillovers into the loading channel where the vessel would be located. Figure 2ii shows a vessel that was successfully loaded and positioned into the V2 chip, in a MOPS buffered filled environment, before delivery of the Vetbond solution.

Attempts were then made to deliver smaller amounts of Vetbond, in a controlled fashion,

similar to the way in which the Trypan blue was delivered. The adhesive, in this design, was to

‘touch‐and‐go’ on the surface of the artery, rather than to flow over it.

a Figure 3. The V2 chip design. The vessel was loaded through the vertical channel (a), and the flow in the channel was reversed in the direction of the arrow, to pressurize the vessel, after Vetbond was delivered to the vessel surface. Trypan blue was injected through the lumen of the vessel, with limited success.

Upon delivery, the Vetbond quickly polymerized when in contact with the vessel surface, as

show in Figure 3. Although the Vetbond did not polymerize before reaching the vessel surface

in the delivery channels, it quickly polymerized afterwards rendering the chip unusable for a

second experiment. At the vessel surface the Vetbond held the vessel in place for the first

minute of the experiment. The flow was reversed in the channel labeled (a) in Figure 3, which

was previously used to load the vessel. Trypan blue dye was delivered through the lumen of the

vessel, in the direction of the arrow shown in Figure 3, using a hydrostatic column containing

Trypan blue stained MOPS, at a pressure of 45 mmHg. Although the lumen of the vessel was initially open, and Trypan blue flowed through the vessel, the Vetbond adhesive quickly

desiccated and broke down the vessel around one to two minutes after reaching the vessel surface (n=5). This resulted in Vetbond leaking into the vertical channels, used to load and pressurize the vessel, and quickly polymerizing and blocking them. Any attempts to dilute the

Vetbond before delivery proved unsuccessful.

3.a.3 Vessel Fixation Via Negative Hydrostatic Pressure (NHP)

Another line of experiments conducted in the development of the AoC device involved fixating

the vessel in place inside the chip by using negative hydrostatic pressure, using a design shown

in Figure 4.

i ii + 45 mmHg

‐45 mmHg ‐45 mmHg a b a b ‐45 mmHg ‐45 mmHg

Figure 4. The S1 chip design. Suction was applied to the top of the vessel at 4 contact points, (a) and (b), using negative hydrostatic pressure, to fixate the vessel in place. Channels (a) connected to a different reservoir than channels (b), which enabled independent control of suction on the left and right sides of the vessel. The vessel was at pressurized at 45 mmHg and the negative hydrostatic pressure was set to match the perfusion pressure (ii).

The S1 chip design, shown in Figure 4, was made up of six channels. The loading channel had a

diameter of 150 microns, and the channels labeled (a) and (b) had a diameter of 120 microns

which narrowed to a diameter of 40 microns at the vessel contact points. Depth was uniform

throughout the device, and chips with depths ranging from 120 to 220 microns were used.

Chips with depths around 180 microns (or greater) caused the vessels to bend in the Z plane when suction was applied, and the lumen was no longer perfusable. As a result, all functional experiments were obtained using S1 chips with a depth of 150 microns. The channels labeled

(a) in Figure 4i joined up outside the field of view, and were both connected to a single common outlet which was attached to a 60 mL syringe. The channels labeled (b) in Figure 4i were

identical to the (a) channels, and were connected to a separate 60 mL syringe. Both syringes were lowered below the level of the chip to an equivalent of ‐45mmHg, creating negative hydrostatic pressure and suction in the four channels located on the left and right of the vessel.

In a similar fashion to the other AoC devices the loading channels ran longitudinally, and the vessels were loaded by applying syringe suction in the direction of the arrow shown in Figure 4i.

A pressurized vessel which was successfully held in place using the S1 chip design is shown in

Figure 4ii. To pressurize the vessel, the flow in the channel previously used to load the vessel into the AoC device was reversed, indicated by the direction of the white arrow. The vessel was pressurized to 45 mmHg, via hydrostatic pressure. The other end of the vessel, outside the field of view, was kept open to allow the blood to exit the lumen as an indicator of vessel

pressurization. The vessel was successfully held in place at 100% of the time at various pressure changes, ranging from 20 to 100 mmHg, over the pre‐set experimental time of 30 minutes

(n=5). Experiments conducted with Trypan blue indicated that the lumen was open and that an estimated 5‐10% of the total fluid perfused through the vessel was lost to the suction channels

before it entered the lumen.

3.b The Vessel Organ Bath, Substance Delivery and Removal

3.b.1 Organ Bath With Flow Over the Vessel (Design 1)

The initial organ bath design, where the vessel would be measured and where various

substances were to be delivered: AoC design BM 1.0 (Figure 5).

a b

Figure 5. The BM 1.0 chip design. The vessel was loaded through the vertical channel (c) which led to the organ bath (highlighted by the dotted box). Substances were delivered via the superfusion lines to the surface of the vessel by inflow from channel (a) and outflow through channel (b). A non‐functional vessel is shown to indicate the orientation of the organ bath.

The BM 1.0 design was developed in parallel with the V1 chip (as previously described), with the main focus of controlling substance delivery (such as PE and Ach) to and from the vessel surface. Substances were to be delivered via syringe pump at a flow rate of 4 mL/h from

channel (a) in Figure 5, and out through channel (b). Channel (c), as previously described, was

used to load and pressurize the resistance arteries. The diameter of the loading channel was

250 microns and the diameter of channels (a) and (b) was 80 microns.

The organ bath, outlined in Figure 5, presented several challenges. Bubble formation in

channels (a) and (b) always occurred at the exit points near the vessel surface. Removing these

bubbles was only possible by manually flushing the AoC channels at a pressure which would also cause irreversible damage to the device, by delaminating the PDMS material from the

bottom glass slide. The 90 degree angled corners of the inspection area, where the vessel was

located, were also sites of nucleation and air bubble formation. Loading the vessel was very

challenging, with the vessels visibly damaged (as shown in Figure 5), due to the large diameter

disparity between the bottom loading channel and the channel used to apply the suction necessary for loading (channel (c)).

3.b.2 Organ Bath With Flow Over the Vessel (Design 2)

a b

Figure 6. The SY 1.1 chip design. Suction was applied through negative hydrostatic pressure at 4 points which pinch the vessel at the top and another 4 that pinch it at the bottom. The organ bath was designed with rounded edges and substances were delivered via the superfusion lines to the surface of the vessel by inflow from channel (a) and outflow through channel (b).

The SY 1.1 chips (Figure 6) were designed using the S1 chip holding mechanism (previously

shown in Figure 4) and with a modified organ bath. All dimensions were kept the same as in the

S1 chip, with the addition of the delivery channels (a) and (b), for delivery and removal of

substances, as shown in Figure 6. An hourglass shape was introduced at the end of the vessel

pressurizing channel, towards the top suction points, which greatly facilitated loading by

stopping the vessel once it reached the correct location for measurement. As the organ bath

was designed with rounded edges this eliminated the problem of bubble formation near the vessel surface.

Resistance arteries from CD1 mice were isolated and loaded as previously described. Vessels were successfully subjected to pressure changes from 20 to 100 mmHg and fixated in place over a pre‐set time of 30 minutes (n=14). However, flowing substances over the vessel, at any

flow rate, either by manual delivery or by syringe pump was not possible (n=12). The average

depth of the chips tested was 150 microns (measured using profilometry). Delivery was possible

with chips that had depths greater than 200 microns (n=4). However, at chip depths where flow

over the vessel was possible the fixation mechanism at the top and bottom of the vessels was

unable to fixate the resistance arteries once the intralumenal pressure exceeded 30 mmHg.

3.b.3 Organ Bath With Flow on the Vessel Lateral Surfaces (Negative Hydrostatic Pressure

Fixation)

Figure 7. The AoC 1.0 design. Suction was applied through negative hydrostatic pressure, as previously described in Figure 6. Substances were delivered in the superfusion channels to the vessel, from a common reservoir, and washed out from the organ bath in the directions indicated by the arrows.

The AoC 1.0 chips (as shown in Figure 7) were built around a design which enabled substance

delivery via flow on the lateral surfaces of the vessel. The fixation mechanism was identical to

that used in SY 1.1 chips. Substances were premixed in the mixing cell (shown in Figure 8) and

delivered from a common source separately to the left and right sides of the resistance arteries, as indicated by the arrows in Figure 7. The two bottom channels, through which substances were removed from the chip, joined together and emptied into a common reservoir. The width of the organ bath was 500 microns, to accommodate any potential changes in vessel diameter

without impeding flow due to increased resistance. Chip depth was 150 microns. Vessels were

successfully subjected to pressures ranging from 20 to 100 mmHg, maintaining pressure and

remaining fixated for periods of over one hour (n=34).

3.c The Mixing Cell

In order to facilitate substance delivery to the vessel the AoC 1.0 design included an on‐chip mixing cell (Figure 8).

i ii

a b c

Figure 8. The on chip mixing cell used in AoC devices. The mixing cell shown in (i) had two connecting ports, (a) and (b) for infusion of two separate solutions, indicated by perfusion of Trypan blue in (a) and MOPS solution in (b). The two streams combined and began mixing at point (c) as they entered the main part of the mixing cell. The combined stream separated to the left and right walls of the artery, after mixing, at point (d).

In the AoC 1.0 design, substances were delivered individually to the right and left sides of the vessel (as shown in Figure 7), but from a common reservoir. The mixing cell in Figure 8i had two input ports, with Trypan blue and MOPS connected to ports (a) and (b) respectively. Both substances were delivered via syringe pump infusion. The substances met and mixing began at the feature labeled (c). As shown in Figure 8ii, mixing of the two substances occurred via diffusion, over a channel of a set length, depending on the molecular weight of the substances

being delivered to the vessel. After being mixed, the combined substance stream was separated at the feature labeled (d) in Figure 8i, and delivered separately to the right and left lateral

surfaces of the resistance arteries tested.

In order to properly define the length of the mixing cell, all substances were delivered at a

combined flow rate of 4 mL/h. By connecting PE at a concentration of 3 µM to port (a) and

MOPS to port (b), and by flowing both syringes at a flow rate of 2 mL/h (a combined rate of 4 mL/h) a final concentration of 1.5 µM PE was delivered to the vessel surface, due to mixing by

diffusion in the mixing cell. Various alterations to the flow rates of the PE and the MOPS syringe pumps allowed the delivery of any concentration within the 0 – 3 µM PE range. This enabled the delivery of solutions over wide concentration ranges without having to connect or disconnect the pumps connected to the chip. The dilution steps were only limited by the maximum and minimum flow rates of the pumps used, as the combined flow rate was kept constant at 4 mL/h for all experiments. This constant was necessary in order to ensure

adequate mixing by diffusion in the AoC mixing cell.

3.d Organ Bath Washout, and Separation of the Perfusion and Superfusion Streams

In order for diameter measurements on the chip to be accurate, and to replicate the effectiveness of the cannulation setup, an effective separation of the perfusion (intralumenal) and superfusion (extralumenal) fluid streams was obtained (Figure 9).

i ii

iii iv

Figure 9. Resistance artery loaded on‐chip and perfused with rhodamine dye (red) through the

lumen, and fluorescein dye (green) in the organ bath superfusion. Chip version AoC 1.0. The vessel is shown at (i) baseline, and at (ii) 4 seconds, (iii) 5 seconds, and (iv) 6 seconds after fluorescein infusion.

Rhodamine (red) was perfused through the lumen of the vessel, at pressures ranging from 20 to

100 mmHg. Fluorescein (green) was delivered from the mixing cell to the vessel surface, at a standard flow rate of 4 mL/h. From a baseline level (where only MOPS is delivered to the vessel surface), shown in Figure 9‐i, fluorescein bathes the vessel completely in 6 seconds, as shown in

Figure 9‐iv. Washout of the fluorescein also occurs over 6 seconds.

Figure 9 also indicates a separation of the fluids flowing in the perfusion line from those in the superfusion line. Although the baseline picture at time 0 does indicate yellow staining, this is

due to the experimental protocol, and will be further explained in the discussion. Within the

same experiment, in Figure 9 (iii) and (iv) a visible amount of fluorescein was lost to the bottom

suction channel. This amount was very limited however and only occurred due to the fact that

the vessel was not fully closed at the bottom end (thus, not fully pressurized) in order to permit

rhodamine to flow through the lumen. When the bottom channel was closed, and the vessel

was pressurized fully, substance wash‐in and wash‐out times were unaffected. Furthermore, no

visible amount of superfusate was lost to the bottom suction channels.

3.e Phenylephrine Dose Response Curves (DRC) on the AoC Device

In order to determine the viability of arteries loaded onto the AoC device, and to determine if the loading and AoC setup damages or changes the contractile properties of the resistance

artery being studied, PE DRCs were measured on‐chip and compared to DRCs obtained on a standard cannulation setup.

a b ) ) 90 100 max max Outer Diameter Inner Diameter 80 Outer Diameter 70 80 60 60 50 40 40 30 20 20 10

0 0 Tone (%constriction of Dia Tone (%constriction of Dia -7.5 -7 -6.5 -6 -5.5 -6.5 -6.0 -5.8 -5.7 -5.5 log [PE] (mol/L) log [PE] (mol/L)

Figure 10. Dose dependent responses to phenylephrine, measured in mesenteric arteries on the pipette cannulation setup (a) and the AoC 1.0 device (b).

Mesenteric arteries (average diameter ~150 µm), isolated and loaded as previously described from CD1 mice, exhibited a maximal outer diameter constriction of 44.5 ±2.5% at 3.0 μM PE

(n=5) on the cannulation setup (a). Dose dependent phenylephrine responses measured on the

AoC 1.0 setup (b) were comparable to the cannulation setup at 3.0 μM PE with a maximal outer diameter constriction of 42.2 ±3.8% (n=5).

The pumps used in the PE DRC experiments had a minimal flow rate of 0.1 mL/h. With a stock

PE concentration of 3.0 µM, and a flow rate of 0.1 mL/h, the lowest concentration of PE

possibly attainable by dilution on the chip mixing cell was 0.075 µM PE. During the heating

process, as the artery was brought to 37oC, it was necessary to maintain a constant flow in both

the PE and MOPS lines. The continuous flow rate was necessary to prevent bubble formation. A

channel with a stagnant flow would become blocked by air bubbles, which formed as they

evaporated out of the MOPS and PE solutions. This often caused the resistance artery being

studied to come in contact with air. As a result of this, the vessel was heated up in the presence of 0.075 µM PE. Once the vessel was properly heated, it was re‐washed in MOPS solution and given 30 minutes before PE DRCs started. It is also interesting to note that the concentration of

0.075 µM PE failed to elicit a constrictory response. Furthermore, this was not directly

comparable to any concentrations on the cannulation DRCs. As a result, the next smallest

concentration of PE which could be readily compared to the cannulation setup was 0.3 µM PE.

This was chosen as the starting point for the PE DRC. A concentration of 0.1 µM PE was not attainable due to the fact that the syringe pump only flowed in increments of 0.1 mL/h.

3.f Acetylcholine Dose Response Curves on the AoC Device

In order to test whether endothelial integrity was preserved in arteries loaded onto the AoC device, we stimulated the vessels using the endothelium‐dependent vasodilator Ach.

90 inner diameter 80

40 dilation 30 20

constriction) maximal of reversal (% 10

0 -8.0 -7.5 -7.0 -6.5 -6.0 -5.5 -5.0 log [Ach] (mol/L)

Figure 11. Ach dose response curve in mesenteric arteries loaded onto the AoC device (n=5).

Mesenteric arteries, isolated as previously described from CD1 mice, exhibited a dose‐ dependent dilation when exposed to Ach. At the highest Ach concentration, a preconstriction obtained using 1 µM PE was 77.8 ± 17.5% reversed by 10 µM Ach (n=5).

3.g Cultured Vessels on the AoC Device

In order to indicate that current cannulation setup protocols are transferable onto the chip, and to show that the AoC device is robust enough to preserve mesenteric artery smooth muscle and endothelial function over extended periods of time, 24 hour culture experiments were

conducted.

100 * diaouter * diainner

75 )

max 50 tone

(%dia 25

0 1 µmol/L PE 10 µmol/L ACh

Figure 12. PE and Ach responses, both inner and outer diameter, in vessels kept in culture for 24 hours on the AoC device. * denotes P < 0.05 for the inner and outer 10 µM Ach dilations compared to their respective 1 µM PE pre‐constrictions.

Vessels cultured for 24 hours on the AoC device exhibited constrictions indicated by

maintaining 63.2 ± 4.3% and 30.4 ± 5.8% of maximal diameter in the outer and inner diameter

respectively, in response to 1 µM PE (n=5). Dilatory experiments indicated a 94.7 ± 7.8% and

89.9± 5.9% reversal of the initial 1 µM PE preconstriction, in the outer and inner diameter

respectively, in response to 10 µM Ach (n=5). Outer diameter constriction to 1.0 µM PE was

30.4 ± 5.8%, which was stronger than the nearest correlating concentration measured acutely,

0.9 µM PE (as shown in Figure 10), which elicited a PE constriction of 17.4 ± 12%.

3.h One‐Sided PE Constrictions

An experiment unique to the AoC device, one‐sided PE constrictions, was conducted to demonstrate the versatility of the chip. In this experiment it was shown that different sections

of the resistance artery being studied can be simultaneously exposed to completely different substances, at different concentrations.

a b 100 100 left wall left wall 90 90 * right wall right wall 80 80 * m) 70 *  70 60 60

50 * 50 * 40 40 30 * 30 20 20

10 ( movement wall lateral 10 (mm) movement wall lateral 0 0 -10 -10 0.0 0.3 0.9 1.5 0.0 0.3 0.9 1.5 PE Concentration (M) PE Concentration (M) Figure 13. One‐sided constrictions obtained on‐chip by stimulating the left vessel wall with PE, while keeping the right wall bathed in MOPS, and measuring the changes in (a) outer diameter and (b) inner diameter. * denotes P < 0.05 when comparing left and right wall movement at individual corresponding PE concentrations.

A modified version of the AoC 1.0 device, AoC 1.1, was used to conduct experiments involving

localized PE delivery to the vessel surface. The AoC 1.1 chips had separate reservoirs connected

to the left and right superfusion channels, instead of a common reservoir which would separate into two channels to bathe both sides of the vessel. Left wall movement, the side from which

PE was delivered to the vessel, was significantly more pronounced than lateral movement in

the right wall, which was only exposed to MOPS solution, at all concentrations tested. Flow rates on both the left and the right vessel side were kept constant, at 4.0 mL/h.

3.i FURA‐2 On‐Chip Measurements

A widely employed technique used in vascular research is the use of the FURA‐2 ratiometric dye in order to study changes in intracellularCa2+. It was therefore important to demonstrate that

the optical properties of the AoC did not inhibit or interfere with FURA‐2 measurements.

Furthermore, as it is possible on the AoC to expose different part of a resistance artery to

different substances, successful FURA‐2 imaging could suggest a potential use for the AoC

device in novel studies of Ca2+ signal transduction. a b 3.0M PE washout 0.93 0.89 0.85 0.81

Ratio 0.77 0.73 0.69 0.65 0 40 80 120 160 200 Time Point (s) Figure 14. FURA‐2 measurements conducted on the AoC, showing smooth muscle cells loading (a) and ratiometric responses (b) to PE stimulation. 43

On‐chip imaging of vessels loaded with FURA‐2 indicated that PDMS and the composition of the chip did not interfere with the wavelengths necessary to conduct FURA‐2 ratiometric measurements, as the loaded smooth muscle cells were clearly visible (n=4). The loading and optical properties were permissive enough to enable the recording of a 3.0 µM PE‐stimulated

Ca2+ response.

4. Discussion

In order for the AoC device to properly mimic and improve upon the classic cannulation setup a

sequential series of experiments were performed. Resistance arteries were fixated in place on

the AoC, and were pressurized. They were then subjected to a wide range of intralumenal

pressure changes to indicate proper fixation. Substance delivery and removal from the vessel surface, in the organ bath, was then enabled and optimized. The last set of experiments focused on indicating that smooth muscle and endothelial cell function was maintained in the

AoC device, even after a 24‐hour culture process.

PDMS ‐ AoC Fabrication Material

As previously mentioned, PDMS chips were fabricated quickly, going from the design phase to

the quality control phase within 24 hours. They were relatively inexpensive compared to other

materials such as glass or silicone. Furthermore, PDMS is transparent, which made it ideal for

the measurement and imaging techniques employed in this study. The material is also non‐toxic

to cells and has a low thermal conductivity of 0.2 W/(m.K)43. While the low conductivity meant

that PDMS devices required 4‐5 minutes to reach body temperature when heated, this ensured

that the interior environment was kept at a fairly constant temperature, as little heat is

dissipated to the external environment37. The ease of bonding PDMS to glass slides was due to plasma treatment, which oxidized its surface and altered its chemistry. This allowed for bonding without having changed the properties or shape of the design39.

4.a Vessel Fixation and Pressurization

The first attempts to fixate the vessels using Vetbond Tissue Adhesive, in the V1 chip design, had several drawbacks. The 3M Vetbond used in this experiment is a modified version of N‐ butyl cyanoacrylate, mixed in an undisclosed fashion. Cyanoacrylate is generally an acrylic resin

which in the presence of hydroxide ions (such as in water) forms strong bonds44. As a result,

Vetbond polymerizes in seconds when it comes in contact with tissue and fluids42. Delivery to

the vessel surface proved to be very challenging. Due to the Vetbond viscosity of 4.3 centipoise

at 25O C, compared to that of water being only 1.002 centipoise, infusion had to take place via

syringe at a high flow rate of 4 mL/h in order to prevent polymerization in the delivery channels. As a result of this, it was not possible to deliver controlled amounts of adhesive around the vessel. This resulted in Vetbond spilling into and blocking other important channels

on the device, at the level of the organ bath.

Another challenge encountered in using tissue adhesive was that it heavily desiccated the resistance arteries that it came in contact with. Attempts to dilute the adhesive proved

unsuccessful.

In order to compensate for the inability to flow Vetbond over the vessel in a controlled fashion

the V2 chip design focused on applying Vetbond to the lateral sides of the vessel, with no flow

over the top and bottom. The smaller channels in the V2 design greatly reduced the amount of

tissue adhesive delivered to the vessel surface in order to reduce desiccation problems. Despite the numerous design changes the adhesive still dehydrated the vessel upon contact, compromising its structure.

Among several primary incentives in developing the AoC device was its ability to provide a more cost‐effective and time‐efficient method of studying resistance arteries. The Vetbond used in the V1 and V2 chip experiments bound irreversibly. While acetone is effective at dissolving cyanoacrylates it also causes notable swelling in PDMS, resulting in channel deformation and

occasional delamination from the glass slide surface45. As a result, a tissue adhesive chip could

only be used once, regardless of whether the experiment was successful or not. While chips can be made out of glass, to prevent deformation, this is a very time‐consuming and expensive process46.

The subsequent attempt to fixate arteries on the AoC device, using negative hydrostatic

pressure, proved to be successful. Vessels held in place in the S1 chip design remained bound

over a wide range of pressure changes, indicating the efficiency of the design. The pressure

used to suction and hold the vessel in place was set at around 45 mmHg in experiments where

intraluminal pressure was also kept at 45 mmHg. This avoided pressure gradients across the vessel walls at the suction points. Vessels loaded in the S1 design were unloaded and reloaded

in the AoC devices multiple times, as the chips were reusable.

For the purposes of this project, the major drawback of the negative hydrostatic pressure

fixation method was the limitation in the overall chip depth. The four pressure suction points in

the S1 devices (Figure 4) can be visualized as vertical strips, running parallel to the chip’s Z axis,

at the point of contact with the vessel. As a result, if the depth of the chip was greater than the

diameter of the vessel (as measured with Diamtrak) flow under and over the vessel would occur

at the suction points. This presented two problems. First, liquid being perfused through the

lumen of the vessel, from the top pressurization channel, would also flow over and under the

vessel and come in contact with the fluid in the organ bath. This would also expose the smooth

muscle cells to the substance delivered through the lumen to the endothelial cells, in a highly

uncontrolled fashion. Secondly, when the vessel was notably smaller than the depth of the chip

(more than 20 microns, as measured with Diamtrak), the vessel would snake in and collapse on itself in the Z axis, making it impossible to pressurize the lumen. Therefore, in order to ensure

proper fixation, and to separate the perfusion and superfusion streams, vessels at least greater

than 20 microns than the depth of the chip had to be used.

The average depth of the S1 chips was 150 microns. As a result the average outer diameter of

the vessels used in the S1 devices was around 170 microns. Vessels on the AoC held by suction

were thus in contact with the top and bottom layers of the device. While this could

mechanically restrain the vessel from dilating in the Z plane, viability experiments (which will be further discussed in this thesis) did not indicate any constraints on vessel behaviour as

compared to the cannulation setups or literature values. It is also important to note that when

using cannulation setups the vessel is also restrained in the Z axis, as it is brought in contact with the glass bottom of the organ bath in order to facilitate measurement. This is not a

notable drawback however, as both systems commonly used in our laboratory to measure

artery diameter (the Felix and the Living Systems) also measure diameter change only in the X

axis of the vessel.

To facilitate the use of negative hydrostatic pressure for fixation, the experimental protocol was

slightly changed. Due to biological variability in the diameter of resistance arteries each chip design tested was fabricated in three or four different depths. This was done in order to ensure that the diameter of each isolated artery was always greater than the diameter of the chip it was being loaded onto. Although this was a fabrication intensive alternative, the chips were reusable and cost‐effective to fabricate. Therefore, negative hydrostatic pressure became the preferred fixation method. This technique was used in final design, AoC 1.0, which was used to obtain experimental data on vessel functionality.

The hydrostatic pressure fixation method also presents potential advantages compared to the

cannulation setups. Vascular endothelium has been strongly associated with many roles in

resistance artery responses, being a key player in vasodilatory pathways. Endothelial

disruptions can have significant negative implications for vessel function. Localized cell damage

in the endothelium can propagate through gap junction and elicit downstream responses even

in non‐affected areas47. The cannulation setup presents several risks to endothelial structure due to the glass pipettes being inserted into the lumen and coming into direct contact with the vascular endothelial cells. Although this is not considered to be a significant problem in long vessels, and the damage is considered to propagate minimally and to be localized to the area

around the sutures and pipette tips, this may not be the case with shorter resistance arteries.

Due to its unique fixation mechanism the AoC device would also permit the study of intact

endothelium in shorter resistance arteries, which would otherwise get damaged on a

cannulation setup.

4.b The Organ Bath

The organ bath was designed with two primary functions: to (i) ensure controlled and efficient

substance delivery and removal to and from the extraluminal surface of the vessel, and (ii) to

ensure adequate separation between the luminal (perfusion) and extraluminal (superfusion)

streams.

The initial design for the organ bath was designed with the intent to flow substances under and

over the vessel. This was to be achieved via infusion from the right side of the vessel and

removal via an identical channel on the left side, in chip version BM 1.0 (Figure 5). The design

however had numerous corners and right angles. These jagged surfaces caused air bubbles to

form and to ‘bubble out’ of the MOPS solution when the chip was heated to 37oC. This made it

impossible to preserve a controlled microenvironment. Furthermore, due to the initial large size of the organ bath it also took a lengthier than optimal time to perform a complete washout of

the organ bath, where the vessel was located.

In order to address these issues the size of the organ bath was greatly reduced. All straight edges were eliminated in the subsequent design SY 1.1 (Figure 6). Although air bubble

formation was no longer a problem, this design proved to be incompatible with the system

used to fixate the vessel in place. Negative hydrostatic pressure fixation, previously described,

is dependent on the vessel being in contact with the top and bottom layers of the chip, or on

having no more than 20 microns of clearance. As a result, the vessel acted like a dam. It was not

possible to flow substances over and under the vessel at an acceptable flow rate, and without

causing artifact deformations on the surface of the vessel. Although numerous experiments

were performed with variations on chip depth, only two outcomes resulted: (i) the substance being infused in the superfusion line would bend the vessel, moving it out of place, collapsing it, and then exiting via the superfusion exit line, or (ii) the substance infused from the superfusion line would come in contact with the vessel, be redirected, and exit the chip via the loading channel or the perfusion channel.

In order to address the problem of substance delivery a new chip design, AoC 1.0 (subsequently

employed to obtain functional data), was used. Substances were delivered from two

superfusion channels, on the lateral sides of the vessel, rather than being flowed over and

under. The substance superfusion channels came from a common reservoir, so there was no

discrepancy between the concentrations at the left and right vessel surface. Due to the fact that all the features in the AoC 1.0 design (Figure 7) were rounded, air bubble formation in the

organ bath did not occur. Furthermore, as a result of the greatly reduced size of the organ bath, with a width of only 500 microns, length of 350 microns, and depth of 150 microns, the volume of the organ bath was only 0.003 mL. This greatly decreased the necessary time for substance

delivery and removal. Experiments conducted using fluorescent dyes indicated that substances

could be delivered to the vessel surface in six seconds, with a complete washout also taking six

seconds (Figure 9). This is a vast improvement over the cannulation setups, in which washouts

are more time consuming and often involve much larger volume amounts than 0.003 µL,

between 3.0 and 5.0 mL. Furthermore, in order to obtain a complete washout in the cannulation setup, several washes are required. Despite rigorous washes, remnants of the previous substance can persist even in that case, on the bottom glass edge of the organ bath.

Even though remnants might still occur in the AoC bath, they would be present in notably

smaller quantities than in a cannulation setup, due to the overall difference in the size of the

organ baths.

In order to validate that the AoC is a functional alternative, adequate separation of the

perfusion and superfusion streams had to be achieved. In cannulated vessels, the substances contacting the endothelial surface are separated from substances coming in contact with the

smooth muscle cells. Vascular permeability is the only factor that controls the interaction between the two substance pools. To illustrate proper separation, fluorescent dyes and

fluorescent beads, combined with quantum dots (fluorescent crystalline semiconductors which

are less prone to photobleaching48), were used. Short term (3h) and long term (24h) experiments respectively suggested a very high degree of separation of the perfusion and superfusion streams, over a wide range of flow rates.

It is important to note however that, unlike the cannulation setup, a small amount of the substance perfused through the vessel lumen is lost to the top suction channel, and does not

enter the vessel. Measurements of the volume accumulated in the top suction channel over a

long‐term 24h period indicated the amount lost to be equivalent to 3‐5% of the total volume

perfused through the vessel, making this a negligible loss. This was dependent on the quality of

the seals, which was primarily determined by artery diameter. A proportion of the fluid in the

superfusion stream was also lost to the bottom suction, as seen in Figure 9‐iv. However, it is

important to note that the fluid loss in the superfusion occurs after the substance has come in

contact with the vascular surface. The substance being ‘lost’ is waste substance that has to be

removed. Furthermore, the loss in the superfusion line is not visible when the artery becomes

fully pressurized. Overall, the net result is a small loss of fluid in the perfusion line and no

relevant loss in the superfusion line under experimental conditions.

It is important to mention that in Figure 9, at baseline time 0, there is slight yellow staining present around the vessel. This, however, is not due to the superfusion and perfusion fluids mixing. In order to obtain the proper camera exposure values, the vessel indicated in Figure 9 had been previously exposed to both rhodamine and fluorescein, alternating in the perfusion and superfusion channels. As such, a small quantity of the dyes had become embedded in the vascular connective tissue. The image in figure 9 serves as a visual example to indicate

perfusion and superfusion separation. However, its main purpose is to indicate that a complete

wash‐in/out can be performed within 6 seconds on the AoC. Further experiments, performed under closely controlled conditions, and which were subsequently published, indicate a nearly‐

complete separation of the perfusion and superfusion streams49.

The organ bath design presents the AoC device with a notable advantage over the cannulation

setup. Due to the open nature of the bath, the cannulation setup is prone to notable

evaporation. The AoC however has a low thermal conductivity, and an enclosed organ bath that

is ~60 000x smaller. This greatly reduces evaporation, and is especially useful in longer

experiments and during incubations of over 30‐45 minutes.

Small volumes and an enclosed bath also enable the use of solutions that are available in very

small amounts, either due to cost or bioavailability limitations. Substances that are delivered on

the AoC reach the vessel surface quickly, 6 seconds after the flow rate on the perfusing syringe

pump is changed. Furthermore, a complete washout of the bath takes less than 1 second. As a

result, biphasic resistance artery responses with a very rapid first component could be easily recorded on the AoC.

4.c The On‐Chip Mixing Cell

A further innovation brought about by the AoC project, and incorporated on the AoC 1.0 devices, is the on‐chip mixing cell (as shown in Figure 8). Multiple substances (PE, Ach, MOPS,

etc.) were connected into the mixing cell, via additional ports. This meant that it was possible to

run through the entire vascular protocols by simply changing the flow rates on the superfusion

syringe pumps. This was achieved by knowing the initial starting concentration of the

substances used, and by giving the substances enough length to mix by diffusion in the mixing

cell, at a flow rate of 4 mL/h. It was thus possible to expose the vessel to all the concentrations

necessary for the DRCs. The mixing cell presents three immediate advantages over the

cannulation setup. First, this method reduces the possibility of human error in pipetting.

Secondly, it reduces the probability that residues are leftover in experiments where several dilutions need to be performed. Third of all, as the measurement process is fully automated, it can be integrated into a computer interface that could remotely run vascular experiments.

There is however one factor to consider in using the mixing cell. The pumps used to deliver the substance must deliver fluid at a sufficiently low flow rate, and in small enough increments, when dealing with wide concentration ranges. For example, if a PE dose response curve ranges from 0.1 µM to 3.0 µM, then a ‘stock’ substance of 3.0 µM PE will be connected to the mixing

cell port A, with a MOPS solution in port B. In order to obtain a concentration of 0.1 µM at the

vessel surface, pump A will need to flow at a rate of 0.133 mL/h and pump B will need to flow

at a rate of 3.867 mL/h, as the combined flow rate is kept constant at 4 mL/h. Such flow rates,

however, are well within the ranges of modern syringe pumps and do not make this an

impediment when using the AoC device.

4.d Phenylephrine and Acetylcholine Dose Response Curves (DRC)

In order to demonstrate that the AoC device is a viable technology for studying resistance

arteries the smooth muscle and endothelial responses of on‐chip resistance arteries were

examined. Vessels studied on the AoC indicated a concentration dependent response to PE, as did the vessels studied on the cannulation setup. This data serves as proof‐of‐concept regarding the functional state of the smooth muscle cells, as they are able to mount a PE response. While the curve obtained on the cannulation setup (Figure 10a) covers a much wider range of concentrations, from 0.03 µM to 3.0 µM, the curve obtained on the AoC only covers concentrations from 0.3 µM to 3.0 µM PE. This occurred because at the time when these experiments were conducted syringe pumps which could flow at low rates were unavailable. As

previously described, due to the mechanics of the on‐chip mixing cell, the lowest concentration

attainable at the vessel surface is dictated by the lowest possible flow rate of the pump.

The highest PE concentration tested on both setups was 3.0 µM PE. It is interesting to note that on the cannulation setup the DRC does not yet reach a plateau phase at this concentration.

However, we felt that exceeding this concentration was unnecessary to demonstrate viability,

since the constriction obtained at this level is well over 30%, which is the accepted viability cut‐

off for vascular protocols in our laboratory. Constricting past this concentration may even cause endothelial damage, introducing an undesirable variable into the experiment. The curve

measured on the AoC device appears to reach a plateau phase. However, this observation can

be misleading due to the fact that the last two concentrations tested on the AoC are 2.7 µM PE and 3.0 µM PE respectively. The relatively small difference between the last two concentrations is because of the aforementioned syringe pump limitations. Therefore, it would be necessary to expand the curves past the level of 3.0 µM PE in order to properly consider whether a plateau

phase does indeed exist.

That, however, is beyond the purpose of this experiment, which primarily seeks a robust PE constriction at the level of 3.0 µM. At this level, the constriction observed is very similar between the two setups. The cannulation setup indicated a constriction of 44.5 ± 2.5%, and

42.4 ± 3.8% was recorded on the AoC setup. This suggests that the smooth muscle cells are responding as expected in vessels loaded and measured onto the AoC device. It is also

noteworthy that vessels on the AoC device also exhibited a robust inner diameter constriction

(78.3% at 3.0 µM PE). Direct comparison with the cannulation setup is not available however

due to the fact that, unlike the chip, the setup used to measure cannulated arteries cannot simultaneously record both inner and outer diameter.

Endothelial integrity measurements of the vessels studied on the AoC setup indicated a strong

concentration dependent Ach dilation in CD1 mice. Due to the equipment availability it was not

possible to conduct similar dilatory experiments with the CD 1 strain on the cannulation setup.

However, it is noteworthy to mention that cannulation data in C57BL/6 mice found in the

literature suggests a dilation of 56%50, notably less than that seen on the AoC. This is supported

by data obtained in our lab.

There are several reasons why the Ach responses were stronger on the AoC. As previously

mentioned, the AoC method does not involve pipettes coming into contact with and potentially

damaging the endothelium of the arteries being studies. Vessels are not collapsed and tied

inwards on the chip; they are rather pinched outwards at the fixation points.

Another possible explanation is that, unlike the organ bath in the cannulation setup, the Ach being delivered to the organ bath in the AoC is constantly replenished. As vessels in the

cannulation setup uptake the Ach present around the vessel it is possible that a gradient is

created in the organ bath. This gradient does not form in the AoC setup, as the Ach at the vessel surface is being constantly replenished via the superfusion line.

While genotypic differences between the CD1 and C57BL/6 strains51 do exist, they appear to be

minimal, and may not account for the difference in responses. Overall, the strong dilatory

responses seen on the AoC suggest that the endothelium is responsive and kept intact.

4.e Cultured Vessels on the AoC Device

Vessels cultured for a 24‐hour period on the AoC exhibited appropriate constrictor and dilatory

responses. Although the data sets measured acutely and after culture are in range of each

other, it is not surprising that cultured vessels are better responders. Vessel ‘recovery’ during culture was previously observed in hamster gracilis muscle arteries in our lab in unrelated

experiments. The culture process on the AoC device seems to maintain resistance arteries

intact over an extended period of time. On chip culture also presents three advantages

unattainable in the cannulation setups. First, due to the small volume of the organ bath, the

amount of culture media used can be greatly reduced. Secondly, there is less chance for

bacterial growth on the vessel surface to occur due to the fact that the fluid in the superfusion

line is in constant flow and removes any possible growth through shear forces. Third, nutrients

are quickly replenished and any metabolism byproducts produced by the vessel during the

culture process are quickly removed by the constant flow, enabling the possibility for very long

term resistance artery studies on the AoC device.

4.f Novel Experiments Unique to the AoC Device

Despite the importance of the microenvironment in resistance artery research, current setups do not allow for its manipulation at discrete locations along the vessel wall. In cannulation

myography the vessel is bathed uniformly in a given solution. However, the ability to establish

regional heterogeneities may prove to be an important step in further elucidating how cell‐cell

interactions control vascular response. In this regard, one‐sided PE constriction experiments

(Figure 13) suggested that it was possible to selectively and locally expose different areas on the surface of the vessel to different substances. Overall, these experiments suggested that the right and left superfusion channels can be separated, and different substances can be perfused

through them. When one side of the vessel was exposed to PE the constrictor response did not spread to the contralateral side. It is important to note that it cannot be determined whether

higher PE concentrations resulted in an increase in wall thickness. This is due to limitations of

the 2D imaging system used in this experimental set. However, determining whether

constriction occured by wall displacement, or by an increase in wall thickness, is beyond the scope of this project. What can be established is that these responses only occured unilaterally, on the side exposed to PE. This highlights an important advantage of the AoC platform: the ability to manipulate arterial microenvironments in a carefully controlled spatiotemporal

manner. This approach could present novel research applications and allow us to address new

biological questions.

An extension to the aforementioned idea was explored with vessels loaded with FURA‐2 (Figure

14). It is important to note that this experiment is not comparing FURA responses between the

AoC and cannulation setups, as this is beyond the scope of the current project. Its purpose is solely to indicate that the PDMS material can optically permit the measurement of a loaded vessel. The arteries were easily visible when loaded onto the AoC device, and a ratiometric response was present when stimulated with PE. PDMS does not appear to interact with the

wavelengths used for FURA measurements. It would thus be possible, for example, to use the

AoC device to study unilateral Ca2+ signal transduction and depolarization in intact arteries. Any

experiments performed along these lines would not be possible using the cannulation setup, as

the microenvironment cannot be separated and manipulated in the same way.

5.Conclusions

In this project, a microfluidic device was developed as an alternative to the standard cannulation pressure myographs used to study resistance arteries. Vessels held in place onto the AoC, using negative hydrostatic pressure, were successfully perfused over extended periods of time. Experiments were conducted to examine the viability of arteries loaded onto the AoC,

by testing smooth muscle and endothelial responses. The results indicated that the new

process did not seem to damage the vessels. Vascular responses were within comparable parameters to standard pressure myography systems. When stimulated with PE, arteries on the

AoC exhibited a maximal outer diameter constriction comparable to the cannulation setup.

Endothelium dependent vasodilation of 83.2% was observed on the AoC at 10.0 µM Ach, above

literature values of 56.0%. Fluorescent experiments, Ca2+ imaging methods, and long term culture experiments were successfully transferred and implemented onto the AoC device.

Furthermore, not only is the AoC an alternative to the cannulation setups but it also improves on a number of limitations present in current setups.

The experiments conducted with the AoC device also indicated improvements in general

resistance artery study protocols. The organ bath on the AoC device is, volume wise, between

60 000 and 100 000 x smaller than the bath in current cannulation setups52. Due to its enclosed

nature, and low thermal conductivity, the AoC is well suited for long‐duration experiments.

Taken together, these two factors also permit the use of substances that are available in very

small volumes, due to cost or availability. Furthermore, the small volume of the bath permits a

complete washout in less than one second, making the AoC ideal to study rapid responses.

Another key feature of the AoC device is that flow in the superfusion line can be continuous for

the duration of an experiment. The first benefit is that any waste products are likely to be

rapidly removed, and nutrients are quickly replenished. Secondly, vessels cultured onto the AoC

might be less prone to bacterial growth on the vessel surface.

The flexibility of the user‐friendly interface further adds to the advantages of the AoC device. A successful cannulation is defined as the artery being held in place, pressurized, and with no apparent damage. PE DRCs must also be comparable to literature values. Based on observations made in our lab, it often takes new users 3‐4 months using pressure myography to

cannulate larger resistance vessels, of about 200 µm, with a success rate of 70‐80%.

Furthermore, the glass pipettes are prone to snapping when the sutures are tied. The sutures

themselves can also slip off the pipette tips, leading to the loss of the vessel being measured.

Training duration and successful vessel fixation appear to be greatly improved upon with the

AoC. Vessel fixation on the device requires the user to activate a syringe pump and to set two

columns of negative hydrostatic pressure, according to a specific protocol. This ensures the

loading and pressurization of an artery onto the AoC device; no extensive manual skills are

involved in the process, other than the vessel isolation itself.

Alongside higher efficiency and the requirement for less training, the AoC device might also

better preserve the structure of the arteries being studied. Smooth muscle cell responses (as

shown by PE DRCs) appear to be comparable between the cannulation setup and the AoC

device. However, the data obtained in this project suggests that endothelial‐mediated Ach

responses are more pronounced on the AoC. A stronger preservation of endothelial integrity

could explain this difference. Further AoC studies in this area, with identical mouse strains,

could indicate a notable advantage over the cannulation setup.

One of the main advantages of the AoC technology is the low cost compared to pressure

myography cannulation setups. The PDMS chips themselves, from the design step to the

finished product, ended up costing on average $2.24 per chip. An AoC device that has

successfully passed quality control can be used, on average, 8 times. This brings the cost per

experiment to 28 cents. A standard cannulation setup has a minimal cost of around $5000. It would need to be employed 18 000 times without needing any maintenance or repairs to be as

cost‐effective as a PDMS AoC device. This experimental number might be virtually impossible to

attain. Based on observations in our lab and within our department, cannulation setups

undergo extensive wear and tear after much fewer experiments than that. More so, the AoC device has a compact design, the same size as a microscope slide. This means it can be used in a wide range of either upright or inverted microscopes without requiring any special adaptations.

The method of fabrication of the AoC device also means that one is able to quickly go through a

large number of iterations and design changes at minimal cost. This confers the AoC design with unique flexibility, as it can be fabricated to any desired size and specification within 48 hours.

The various developmental iterations of the AoC can be summarized in Figures 15 and 16:

cannulation cannulation bath

Vetbond ‐ version V2 bath ‐ version BM1.0 (page 22) (page 27)

NHP ‐ version S1 bath ‐ version SY 1.1 (page 25) (page 29)

AoC version 1.0 (page 30)

Figure 15. Flowchart of the AoC parallel development process, indicating the progression of the fixation method (NHP) on the left, and the progression of the organ bath design on the right. 65

1 cm

Figure 16. The completed AoC device is shown in the top left corner, with all microfluidic connections attached. The diagram on the right (courtesy of the Gunther lab) is a schematic showing the mixing cell and the organ bath. The bottom picture indicates the organ bath, blown up to size, with an artery fixated in place.

Future Directions

5.a Basic Science

The development of the AoC device has the potential to open many avenues in basic science resistance artery research. In the near term, the chip can be used to greatly automate and increase research productivity while studying intact resistance arteries. Due to the fact that measurement protocols are carried out by syringe pumps, which are connected to a computer

interface, it is possible to run hours‐long protocols with just the push of a button, and no other

user input. This can then lead to the development of a mass‐throughput chip, which can hold

and measure 3‐4 resistance arteries at the same time. This presents several advantages. First, it

would be possible to obtain an n=4 in the same time that it currently takes to run one vessel protocol. Secondly, this would be done without requiring the user to be present during the protocol, and without requiring extensive manual skills in vessel manipulation. Third of all, and

most importantly, the n number would be obtained under nearly identical experimental

conditions. The vessels on the chip would be exposed to the same substances in parallel, at the

same concentrations, and for the same durations. This will greatly reduce confounding variables associated with experimental protocol heterogeneity.

Another near‐term application would be the use of the AoC in studying cell‐cell signaling. As this project has shown, due to the flexibility of the AoC device, it is possible to expose different parts of the vessel surface to entirely different substances. Vascular Ca2+ responses were also successfully measured on the AoC device. Taken together, these results suggest that the AoC

could be used as an important tool to study signal transduction and signaling pathways in

isolated intact resistance arteries.

5.b Clinical Applications

At its culminating point, due to its portability, simplicity of use, and mass throughput potential,

the AoC device could be used in a clinical setting. Samples obtained from biopsies could be used

to study individual patient resistance artery responses to a particular drug, in an approach

towards personalized medicine. This would parallel and, in some cases, reduce the need for

pharmacogenomic‐based approaches. Genotyping for each patient can often be time and cost‐

intensive. Furthermore, although a gene can be associated with different drug responses, there

are often variations present in the population. As a result, the presence of a gene does not guarantee a resistance or susceptibility to a certain drug in a patient53. The AoC would be able

to overcome this by skipping ahead and looking directly at the tissue level, in each patient.

Individual responses to a particular drug could be observed and analyzed in real time, in the

vascular bed of interest, leading to a more efficient treatment approach.

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