Microfluidic System for Planar Patch-Clamp Electrode Arrays

Home , Automated patch clamp

Abstract

Xiaohui Li

Yale University

2006

The patch clamp has been widely accepted as a standard technique for fundamental

studies of ion channel proteins, and discovery of drugs that affect these proteins.

Traditional patch clamp has a very low throughput which has been proven to be a

bottleneck for the drug discovery process. Planar patch-clamp electrode array, which is

scaleable and easy to use, provides a potential way to solve this problem.

We present a microfluidic system integrated with disposable cell-interface partitions

for simultaneous patch clamp recordings. A disposable partition is made by bonding an

air-blown PDMS partition, which has a 2 µm air-blown aperture, to a small glass washer.

Then it is reversibly sealed to the fluidic system having fluid exchange channels with

isolation valves and Ag/AgCl electrodes. Fluid channels are molded from PDMS using

microlithographically defined molds. At the cross-over point, channels in different layers

formed a valve. Ag/AgCl electrodes are fabricated with standard microfabrication

techniques. The suitability of PDMS valves and microfabricated Ag/AgCl electrodes for

patch clamp measurement are examined in this report. Gigaseal patch recordings from

RBL-1 cells are obtained with a 24% success rate. Our system allows simultaneous

recordings from valve-isolated electrodes.

Microfluidic System for Planar Patch-Clamp Electrode Arrays

A Dissertation Presented to the Faculty of the Graduate School of Yale University in Candidacy for the Degree of Doctor of Philosophy

Xiaohui Li

Dissertation Director: Mark A. Reed

Dec 2006

iii Acknowledgements

This thesis was written based on five-year collaborating work. During the study period, I obtained tremendous help and support from people with various backgrounds.

Hereby, I would like to acknowledge it and extend my gratitude to them. Without their tireless and patient help, I would not expect to accomplish the thesis successfully.

I am deeply indebted to my supervisors: professor Mark A. Reed and professor Fred J.

Sigworth. Their constructive advising provided me great help through my research period and also in drafting this thesis. I have been under their supervision for 5 years. I owe them immense gratefulness for teaching me research skills as well as an attitude to both research and life. I greatly appreciate that I have the opportunity to work with them and learn from them.

I also thank the rest of my previous and current advisory committee members: professor Katepalli R. Sreenivasan, professor Marshall Long, professor

Alessandro Gomez , professor Juan de la Mora , professor Tso-Ping Ma and professor

James Duncan. They have provided very helpful guidance to make my PhD research progress move forward smoothly.

My colleagues in the two research groups under professor Mark A. Reed and professor Fred J. Sigworth have supported me a lot in my research work. I am especially grateful to Dr. Kathryn G. Klemic who directly supervised me on the research and is always ready to help. I also thank Dr. Youshang Yang and Ms. Yangyang Yan for their helpful advice on cell culture and patch clamp techniques. I thank Dr. James F. Klemic,

iv David Routenberg, Eric Stern, Aric Sanders, Ryan Munden, Stan Guthrie, Dr. Wenyong

Wang, Dr. Ilona Kretzschmar, Dr. Glenn Martin, Dr. Menno de Jong, Dr. Takhee Lee, Dr.

Guosheng Cheng and Dr. Nilay Pradhan for their cooperative support and for the

wonderful suggestions they have ever given during the cleanroom and general lab work.

I appreciate Dr. Liguo Wang, Dr. Shumin Bian, Dr. Qiuxing Jiang, Dr. David Chester and Puey Ounjai for their help in the biological laboratory.

I thank my friends who have helped me in my academic research and studies. Dr.

Zhengting Jiang helped me in using the scanning electron microscope at the Department of Geology and Geophysics. Dr. Rustom Bhiladvala is my first friend in US and introduced me to the technology of microfabrication. Many friends helped me out unselfishly: Linlin Wang, Chris Liu, Zhongping Bao, Biao Li, Yifan Chen, Beelee Chua,

Huiming Bu, Dechao Guo, Yanxiang Liu, Weiwei Deng, Yu Xiang, Jian Xu, Leidong

Mao and etc. It is my great pleasure to come to know them during the journey of my life.

I also have a church home in New Haven. I have met many brothers and sisters in the

Calvary Baptist Church. Their spiritual and physical support has given me enormous encouragement during the last three years especially whenever I came across difficulty.

I am especially grateful to Huiyuan Chen, Weihua Niu, Tiehong Wang, and pastor Roc

Wang. Without their vast enthusiasm, splendid planning, and unreserved efforts, our wedding would have never been like what I had last October.

I feel a deep sense of gratitude for my parents. They have always been supportive to me at their best in my life. The happy memory of my father constantly inspired me to

v overcome obstacles and keep moving forward during my Ph.D. period and it will keep

me on during the rest of my life.

I would like to give my special thanks to my wife Baohui whose patient love enabled the completion of my thesis.

This research has been supported and funded by NIH grant EB-002020 to F.J.S. I am very appreciative to Yale University for placing me in the world-top research

environment with preeminent faculty and researchers and for providing all kinds of equipments to meet the experiment needs.

Contents

Chapter 1. Introduction...... 1

1.1 Planar Patch-Clamp Electrode Array………………………………………….1

1.2 Outline of the Thesis…………………………………………………………..2

Chapter 2. Research Background……………………………………………5

2.1 Ion Channels…………………………………………………………………..5

2.2 Patch Clamp Technique……………………………………………………...10

2.2.1 Patch Clamp Configurations……………………………………….12

2.2.2 Whole-Cell Patch Clamp…………………………………………..15

2.2.3 Disadvantages of Traditional Patch Clamp………………………...15

2.3 Planar Patch Clamp…………………………………………………………..16

2.3.1 Cell Guidance in Planar Patch-Clamp……………………………..18

2.3.2 Planar Patch-Clamp Structure……………………………………...21

2.3.3 Materials for Planar Patch-Clamp………………………………….26

2.4 Microfluidics…………………………………………………………………30

2.5 Planar Ag/AgCl Electrodes…………………………………………………..34

vii Summary…………………………………………………………………………37

Chapter 3. Device Fabrication……………………………………………...39

3.1 A Disposable Planar PDMS Patch Partition…………………………………40

3.1.1 Partitions Molded with Microfabricated Silicon Master…………..40

3.1.2 Partitions Fabricated with the Air-Molding Technique……………44

3.2 PDMS Isolation Valves………………………………………………………47

3.2.1 Fabrication of PDMS Valves………………………………………47

3.2.2 Valve Isolation Resistance…………………………………………50

3.2.3 Valve Lifetime……………………………………………………..52

3.2.4 Simulation of the Valve Deformation……………………………...53

3.3 Fabrication of PDMS Microfluidics…………………………………………57

3.4 Planar Ag/AgCl Electrodes…………………………………………………..57

3.4.1 Fabrication of Ag/AgCl Electrodes………………………………..57

3.4.2 Lifetime of Planar Ag/AgCl Electrodes……………………………58

3.5 Assembly of the Microfluidic Device………………………………………..61

Summary…………………………………………………………………………63

Chapter 4. Results and Discussion…………………………………………65

viii 4.1 Cell Culture and Preparation…………………………………………………67

4.2 Recording Solutions………………………………………………………….67

4.3 Harvesting Cells……………………………………………………………...67

4.4 Recordings and Analysis……………………………………………………..67

4.5 Single Patch Electrode Measurement………………………………………..69

4.6 Compatibility with Commercial Planar Partitions…………………………...71

4.7 Simultaneous Measurement Isolated by Microfluidic Valves……………….71

4.8 Electrode Solution Exchange………………………………………………...73

4.9 Noise Comparison with Glass Pipette………………………………………..74

Summary…………………………………………………………………………77

Chapter 5. Conclusions and Future Direction…………………………..78

5.1 Summary of the Key Accomplishments……………………………………..78

5.1.1 Fabrication of Planar Partitions and the Microfluidic System……..78

5.1.2 Test of the Microfluidic System…………………………………...79

5.2 Suggestions for the Future Work…………………………………………….79

ix List of Figures

2.1 A cell membrane structure………………………………………………………...6

2.2 An ion channel protein…………………………………………………………….7

2.3 A gigaseal………………………………………………………………………...10

2.4 Cell-attached and whole-cell configurations…………………………………….11

2.5 Different configuration of conventional patch clamp……………………………14

2.6 A planar patch-clamp configuration……………………………………………..17

2.7 The CytoPatchTM chip……………………………………………………………19

2.8 Microfluidic chip for single cell patch-clamp measurement…………………….21

2.9 A smoothed DRIE etched aperture for planar patch clamp measurement……….23

2.10 A hollow SiO2 nozzle for planar patch-clamp measurement…………………...24

2.11 Side trapped patch-clamp array on a microfluidic platform……………………25

2.12 A Nanion glass chip…………………………………………………………….27

2.13 Planar PDMS patch-clamp recording system…………………………………..29

2.14 A planar silicon chip based patch-clamp system (QPatchTM)…………………..31

2.15 A pneumatically actuated valve………………………………………………...32

2.16 An elastomeric one-way diaphragm valve……………………………………...33

2.17 A very large scale microfluidic comparator chip……………………………….34

2.18 An exhaustible Ag/AgCl electrode……………………………………………..36

x 2.19 Structure of a thin-film Ag/AgCl electrode…………………………………….37

3.1 Schematic cross-section view of the microfluidic system……………………….39

3.2 Process of fabricating silicon master…………………………………………….41

3.3 SEM pictures of a microfabricated silicon master……………………………….42

3.4 Molding PDMS partition from the microfabricated silicon master……………...43

3.5 Micromolded PDMS partition from silicon master……………………………...44

3.6 Process of fabricating disposable planar PDMS patch partitions………………..45

3.7 Process of fabrication multilayer PDMS structure and valves…………………..49

3.8 16 valves of different dimensions………………………………………………..50

3.9 An optical picture of the microfluidic valve……………………………………..52

3.10 Electrical resistance degradation of fluidic valves……………………………...53

3.11 Simulation of a PDMS valve…………………………………………………...54

3.12 Microfluidic device fabrication procedure……………………………………...56

3.13 Set up to measure the lifetime of Ag/AgCl electrodes…………………………59

3.14 Potential drift of microfabricated Ag/AgCl electrodes…………………………60

3.15 A microfluidic device for parallel patch clamp measurements………………...61

3.16 A microfluidic device for simultaneous patch clamp measurement……………62

3.17 A microfluidic device for electrode solution exchange………………………...63

4.1 A Cartoon picture of the microfluidic system……………………………………65

xi 4.2 A simple setup for planar patch clamp measurement……………………………66

4.3 Recordings from RBL-1 cells with the microfluidic system…………………….69

4.4 Simultaneous recordings from two RBL-1 cells isolated by fluidic valves……...72

4.5 Cross-talk test of the patch-clamp electrodes……………………………………72

4.6 The cavity underneath the partition………………………………………………………74

4.7 The capacitance of the electrode…………………………………………………………76

xii List of Tables

3.1 isolation and threshold pressure of microfluidic valves...……………………….51

3.2 Comparison of simulated results and measured threshold pressure………..…………..55

4.1 Seal resistance from RBL-1 cells………………………………………………...70

xiii List of Abbreviations

CHO Chinese Hamster Ovary

DRIE Deep Reactive Ion Etching

HTS High Throughput Screening

LPCVD Low-Pressure Chemical Vapor Deposition

PDMS Polydimethylsiloxane

PECVD Plasma Enhanced Convention Vapor deposition

RBL-1 Rat Basophilic Leukemia

RIE Reactive Ion Etching

SEM Scanning Electron Microscope

SITE Single Ion Track Etching

TMCS Trimethylchlorosilane

xiv Chapter 1

Introduction

1.1 Planar Patch-Clamp Electrode Array

The patch clamp has been widely accepted as the standard technique for fundamental

studies of ion channel proteins, and for the discovery of drugs that affect these proteins

(Xu et al., 2001). The traditional patch clamp system consists of a fire-polished glass pipette with a 1-2 µm diameter tip, which is carefully pressed onto a cell membrane with a micromanipulator. The membrane patch is sealed to the pipette (sometimes by suction), and therefore is electrically isolated. Additional suction or a voltage pulse breaks the patch membrane, yielding the whole cell recording configuration. Thus, the

patch clamp technique measures the ionic current through the membrane patch or the

entire cell membrane area. However, this technique is very labor-intensive and requires

expensive equipment.

To meet high-throughput screening requirements, many efforts have been taken to

improve the patch clamp system. Planar patch-clamp electrodes, which are scaleable and

easy to use, have been developed using the following materials: silicon oxide coated

nitride membranes (Fertig et al., 2000), deep RIE etched silicon holes coated with

PECVD oxide(Pantoja et al., 2004), polyimide films (Kiss et al., 2003), track-etched

1 quartz (Fertig et al., 2002), silicon oxide nozzles (Lehnert et al., 2002), glass substrates

(Xu et al., 2003) and oxygen plasma treated PDMS poly (dimethylsiloxane) (Klemic et al., 2002; Klemic et al., 2005). Most of these electrodes have a proven reliability for

obtaining patch clamp recordings, as well as a high fabrication cost. As a material,

PDMS has the potential of much lower cost than any other materials. Fred Sigworth’s

research group (Yale University) has pioneered in fabricating disposable planar PDMS

cell-patch interfaces (“partitions”) for patch-clamp measurements since 2001 (Klemic et

al., 2002; Klemic et al., 2005).

One advantage of planar patch-clamp electrodes is the possibility of microfluidic

integration for low noise and solution exchange. Microfluidics have been integrated with

a single planar patch-clamp system to improve the noise level and to realize fast solution

exchange (Pantoja et al., 2004). However, dense arrays of electrodes also need

microfluidics to allow for common fluid lines. These lines, in turn, need isolation valves

to electrically isolate neighboring electrodes during measurement.

Our strategy is to develop a microfluidic system containing isolation valves and

planar Ag/AgCl electrodes. This reusable microfluidic system is to be integrated with

disposable PDMS patch partition array. This system allows simultaneous planar patch-

clamp measurements.

1.2 Key Contributions in the Thesis

The significant accomplishments of the present research consist of the fabrication of a

microfluidic system for planar patch-clamp electrode array, successful recordings from

2 RBL-1 cells, together with the first demonstration of simultaneous patch-clamp

recordings with the microfluidic system. The microfluidic system contains isolation

valves and planar Ag/AgCl electrodes. Disposable PDMS patch partitions are integrated

with this reusable microfluidic system for multiple patch-clamp measurements.

1.3 Outline of the Thesis

In chapter 2 we review the current research status of planar patch-clamp and the

necessity of fabricating a microfluidic system for high-throughput planar patch-clamp

measurements. A microfluidic system with isolation valves is needed to obtain

simultaneous patch-clamp recordings from a high density electrode array. Pneumatically actuated valves based on multilayer PDMS technology, which make large scale microfludic integration become possible, are quite useful for a high density patch electrode array. Development progress for planar Ag/AgCl electrodes is also reviewed.

In chapter 3 we go through the process of fabricating a disposable patch partition, a microfluidic device with isolation valves, and planar Ag/AgCl electrodes. We also examine the steps to assemble them together. A disposable electrode is made by bonding an air-blown PDMS partition to a small glass washer. Then it is placed onto the fluidic system, which has fluid exchange channels with isolation valves and Ag/AgCl electrodes.

Fluid channels are molded from PDMS using microlithographically defined molds.

Electrical resistance of the isolated valves seems to be higher than 10 GΩ, desirable for multi-electrodes recording. Ag/AgCl electrodes are fabricated with standard

3 microfabrication techniques. The lifetime of the isolation valves and Ag/AgCl electrodes

is measured.

In chapter 4 we test the microfluidic system. The gigaseal rate is 24 % for RBL-1 cells with our patch-clamp system. Simultaneous whole-cell recordings from RBL-1

cells are obtained with the microfluidic system. In addition, we also test the compatibility of the microfluidic system with commercial planar glass partitions. These results demonstrate the potential of a PDMS microfluidic system for high density arrays of planar patch clamp electrodes for high throughput measurement of ion channel activity.

Electrode solution exchange was also tested with our system.

In Chapter 5 we summarize the planar patch-clamp project and discuss its research direction in the future.

4 Chapter 2

Research Background

2.1 Ion Channels

One universal feature of all cells is that they have an outer limiting membrane, which is called the plasma membrane, crucial to maintain the essential differences between the intracellular and extracellular environment. The plasma membrane of cells normally

consists of a thin (about 5 nm) lipid bilayer and protein molecules, which are held

together mainly through noncovalent interactions (Figure 2.1). The lipid bilayer, inside

of which is primarily comprised of low dielectric hydrocarbon chains, is highly

impermeable to hydrophilic and charged molecules.

The transport of ions (e.g. Na+, K+, Ca2+ or Cl-) and small water-soluble organic

molecules across the membrane is accomplished by specialized transmemebrane proteins,

called ion channels (Figure 2.2). The opening and closing of ion channels are called gating. In the open state, ions can flow through a single ion channel pore at prodigious rates greater than 107 ions/second.

Figure 2.1 A cell membrane is composed of a lipid bilayer and proteins. Ion channels are specialized transmemebrane proteins which allow ions to pass through the cell membrane. From (Bullock and Henze, 1999).

Ion channels can be classified into six categories as follows, according to which chemical or physical modulator controls their gating activity:

- Extracellular ligand-gated channels;

- Intracellular ligand-gated channels;

- Voltage-gated channels;

- Inward rectifiers;

- ATP gated channels;

- Gap junction channels.

Figure 2.2 Ions transport through an ion channel. From http://campus.lakeforest.edu/~light/teaching.html.

Ion channels can also be classified according to which ion is transported. The most

prominent channels are sodium channels, potassium channels, calcium channels and chloride channels. Voltage gated sodium channels are crucial for the propagation of

action potentials in excitable membranes. They cause the cell membrane to depolarize by

allowing the influx of sodium ions into the cell (Denac et al., 2000). Potassium channels

are essential in excitable membranes. They are responsible for repolarizing the cell

membrane after an action potential has passed (Sansom et al., 2002). Voltage gated

calcium channels perform a number of important biological functions, such as stimulating

the contraction of skeletal and cardiac muscle (Benitah et al., 2002). Chloride channels

display a variety of important physiological and cellular roles including regulation of pH, volume homeostasis, organic solute transport, cell migration, cell proliferation and differentiation (Szewczyk, 1998).

7 Ion transporters can be classified as carriers or channels according to whether the

transport is active or not. For carriers, the active ion transport is carried out by a

conformational change that occurs within the protein forming an opening through which

specific molecules can pass. For channels, the passive ion transport is carried out by its

membrane-spanning hydrophilic structure which, when open, allows molecules to pass.

The malfunction of ion channels leads to diseases such as heart disease, neuropathic pain, diabetes, autoimmune diseases, cystic fibrosis and migraine. For example, the autonomic nervous system (ANS) regulates heart cells through receptors that modulate certain ion channels to influence ion movement. Ion channels play a vital role in neuronal signal transduction, neurotransmitter release, muscle contraction, and cell secretion, and even influence enzyme activation and gene transcription.

Ion channel activity can be recorded by measuring membrane potential, extracellular action potential, ion flux, and the patch-clamp technique (Sigworth and Klemic, 2005):

The cell membrane potential reflects ion channel activity and can be measured directly with a microelectrode, a saline-filled glass micropipette that impales the cell.

The membrane potential can also be monitored using voltage-sensitive optical probes with less precision.

Action potentials of neuron cells can be detected with extracellular microelectrodes.

The signals are very small, typically less than 1 mV in amplitude, and dependent on the extracellular current pathways. Although extracellular action potential measurements contain less information than membrane potential measurements, extracellular

8 microelectrodes are useful in monitoring the action potential activity from a population of

cells, for example in brain slices and in heart tissue (Melani et al., 2005).

Ion channel activity can also be monitored by measuring the ion flux across the cell

membrane. The ion flux generated by most types of ion channels is too small to be detected. However, there are two remarkable exceptions. In one case, rubidium ions can serve as an excellent tracer of most potassium channels, and fluorescent dyes sensitive to

rubidium report the total permeability of the channels (Terstappen, 1999; Terstappen,

2004). In the other case, Ca2+ fluxes can be measured with very high sensitivity. High-

affinity Ca2+-sensitive fluorescent dyes has been used to observe the opening and closing

of single neurotransmitter-receptor channels (Demuro and Parker, 2005).

The development of fluorescent voltage-sensitive dyes has improved the throughput

of membrane measurement from a few tests per day to tens of thousands of data points

per day, greatly enabling drug discovery for various types of ligand- and voltage-gated

ion channels (Falconer et al., 2002). However, these techniques are limited by their

inability to control the membrane potential and thus provide less information about

channel activity and conductance than patch-clamp techniques (Xu et al., 2001).

Patch clamp has been the central technique in electrophysiology since 1980s. It

directly measures the current or voltage drop through a small patch of cell membrane and

monitors the active channel function. Current pulses of a few pA/ms can be resolved

with the aid of state-of-art patch clamp amplifiers. Single channel recording yields

information about unitary conductance and kinetic behavior of ionic channels. Single

channel recording also leads to the exploration of new classes of ion channels, their ion

9 selectivity, conductance, voltage dependence, and ligand sensitivity. The patch-clamp technique also permits investigation of ion channels that are not electrically excitable

(Sakmann and Neher, 1995).

2.2 Patch Clamp Technique

The patch clamp technique was originally developed by Neher and Sakmann (Neher and Sakmann, 1976) to resolve currents through single acetylcholine-activated channels in cell-attached patches of membrane of frog skeletal muscle. Later work (Hamill et al.,

1981; Sigworth and Neher, 1980) significantly improved the noise level and patch stability. The development of the patch clamp method was honored with a Nobel Prize

(1991) and led to the foundation of molecular electrophysiology as a recognized science.

Figure 2.3 A gigaseal forms between the tip of glass pipette and the cell membrane. From (Neher and Sakmann, 1992).

10 The principle of the patch clamp technique is very simple, but based on many

ingenious innovations. By carefully heating and pulling a small glass or quartz capillary

tube, a very fine pipette can be formed. When pulled by machine, the opening of the

pipette tip may be only 1-2 µm in diameter. This glass pipette is pressed gently onto the cell membrane. The application of slight suction within a freshly prepared glass pipette

increases the seal resistance by several orders of magnitude by an unknown mechanism,

so that the seal resistance is larger than a giga ohm, now called Gigaseal (Figure 2.3).

Thus, the ion channels in the opening of the pipette tip are the only connection between

the inner side of the cell and the electrode fluid in the pipette. This is the basic

configuration of patch clamp and is known as the "cell-attached" configuration. Several

other patch-clamp configurations are derivatives of the “cell-attached” configuration.

Figure 2.4 Cell-attached and whole-cell variants of the patch-clamp technique. (A) Photograph of a patch pipette sealed to a cultured neuron. (B) Schematic of a cell-attached recording, where current is collected by the pipette from a small area of membrane. (C) Schematic of a whole-cell recording, in which the patch membrane is ruptured, giving the pipette access to the cell interior. From (Sigworth and Klemic, 2005).

After a gigaseal is formed (a small patch is electrically isolated from other part of the

cell membrane, Figure 2.4A), the pipette collects most of the current flowing through the

patch of membrane and transfers it to a current-measuring amplifier (Figure 2.4B). If the

patch membrane is ruptured, the pipette collects the current passing through the whole

11 cell membrane and carries it to a current-measuring amplifier (Figure 2.4C) (Sigworth and Klemic, 2005).

2.2.1 Patch Clamp Configurations

Several variations of the patch-clamp technique are applied according to different research aims. The "excised patch" configuration includes “inside-out” and “outside- out” configurations. “Cell-attached” and both “excised-patch” techniques are used to study the behavior of ion channels on the section of membrane attached to the micropipette (sometimes referred to single channel recording since the number of channels is in the order of one). “Whole-cell” configuration and the “perforated” configuration allow the researcher to study the electrical behavior of the entire cell membrane. Below are several variants of patch-clamp technique:

• “Cell-attached” configuration: This is the prototype configuration of patch-

clamp. It happens immediately after gigaseal forms. This permits the

recording of currents through single ion channels in that patch of membrane.

• "Inside-out" configuration: At the “cell-attached” configuration, the electrode

is quickly withdrawn from the cell, and the patch of membrane is ripped off

the cell. Therefore, the intracellular surface of the patch membrane is exposed

to the external media. This is useful when an experimenter wishes to

manipulate the environment affecting the inside of ion channels.

• “Whole-cell” configuration: At “cell-attached” configuration, a pulse of

vacuum suction or voltage is applied to rupture the patch membrane that is

12 inside the pipette opening. Thus the saline in the pipette is connected to the

interior space of the cell. In the whole-cell configuration, the soluble

intracellular contents will slowly be replaced by the saline of the pipette.

Therefore, any properties of the cell depending on soluble intracellular

contents will be altered. The whole-cell configuration measures currents

through membrane channels of the whole cell.

• "Outside-out" configuration: At the “whole-cell” configuration, the pipette is

slowly withdrawn from the cell so that the patch is torn off the cell. Then the

patch anneals and forms a ball of membrane at the end of the pipette, making

the outside of the membrane become the outside surface of the ball. “Outside-

out” patching provides opportunities to examine the properties of a single ion

channel when it is not in the cell environment.

• “Perforated” configuration: In the cell-attached configuration, a new solution

containing small amounts of an antibiotic, such as nystatin, is added into the

electrode solution to form small perforations on the patch membrane. The

“perforated” configuration has the advantage of keeping the intracellular

content, but also has two disadvantages. On the one hand, the access

resistance is higher because access resistance contains both the electrode

resistance and the resistance at the electrode-cell junction. This high resistance

will decrease current resolution, enhance recording noise, and magnify any

series resistance error. On the other hand, it may take a significant amount of

time (several minutes) for the antibiotic to perforate the membrane. Therefore,

the time length of experiment is limited.

Figure 2.5 Different configuration of conventional patch clamp. (A).A clean pipette approaches a cell. (B). A mild suction helps the gigaseal between cell membrane and pipette tip (cell-attached). (C). A strong pulse of suction breaks the patch and forms a whole-cell configuration. (D). From cell-attached mode, retracting the pipette would tear the membrane patch from the cell and generate an inside-out patch. (E). From whole-cell recording, retracting the pipette would tear a piece of membrane from the cell. (F). The torn membrane anneals and generate an outside-out patch. From (Purves, 2004).

Figure 2.5 shows the Cell-attached, inside-out, outside-out, and whole cell configurations of patch-clamp (Purves, 2004).

14 2.2.2 Whole-Cell Patch Clamp

This is the most powerful variant of patch clamp technique. Whole-cell patch clamping is used when we want to measure the average current across the entire surface area of one cell. Figure 2.4C illustrates the whole-cell configuration (Sigworth and

Klemic, 2005). The whole-cell configuration has two advantages. First, the current level is about two orders of magnitude higher than “cell-attached” configuration and “excised- patch” configurations. Therefore, the noise in whole cell patch measurement is more tolerable than these two configuration variants. Second, the pipette stays still after the seal forms, consequently, it is more reliable to obtain whole-cell recording than “excised- patch” recording. In ion channel drug discovery, whole-cell patch clamp is the predominant patch clamp application.

2.2.3 Disadvantages of Traditional Patch Clamp

The patch clamp technique has been widely accepted as a standard for monitoring the behavior of ion channels (Xu et al., 2001). Its high time resolution (microseconds) and precision (picoampere range) make it unparalleled in comparison with other ion channel assay techniques. The probing nature of traditional patch clamp pipette also renders this technique a lot of flexibility. For example, the pipette can penetrate the top layers of tissue and patch cells inside living tissue such as a living brain; patch clamp can also work at inside-out or outside-out mode so that single-channel recording becomes possible; the pipette can also be moved quickly between lines of solution flow for fast solution exchange. The last property has been integrated with microfluidics techniques and has been commercialized (DynaflowTM Technology, www.cellectricon.com).

15 The drawbacks of conventional patch clamp stem from the nature of probing cells

with micropipettes, which requires expensive equipment and is highly labor-intensive.

Although this probing feature endows conventional patch clamp with many versatilities,

it limits the throughput of electrophysiology and has proven to be a bottleneck for the

drug discovery process. Typically, with patch pipettes, an experienced technician can

only measure patch-clamp recordings from 10-20 individual cells per day. This level of

throughput is far below that required for primary (thousands to tens of thousands per day)

or secondary (hundreds to thousands per day) pharmaceutical drug screening. Therefore

conventional patch clamps are not practically applicable for high throughput screening

(HTS).

2.3 Planar Patch Clamp

Both screening of pharmaceutical compounds and functional analysis of ion-channel genes require improvement of the patch-clamp technique for high throughput and “ease of use”. In the late 1990s, several companies and university laboratories around the world started to develop the planar patch-clamp technology to meet the need of highly parallel automated voltage-clamp recordings from cultured mammalian cells.

Rather than manually probing pipette to the cell, planar patch clamp technology uses a stationary array of micromachined holes to record from cells. A cell suspension is dropped onto a planar device containing a microstructured aperture. The difference between the conventional patch clamp configuration and the planar patch clamp configuration is presented in Figure 2.6. This revolution turned the hard-to-automate

16 pipette micromanipulation process into an easy-to-automate cell positioning process

(Fertig et al., 2002).

Figure 2.6 Replacing the patch clamp pipette with a planar chip. (A) Whole cell configuration of the classical patch clamp technique. Using an x-y-z micromanipulator and a microscope, the tip (diameter 1–2 µm) of a glass pipette filled with electrolyte solution is positioned onto a cell. A scanning electron microscope (SEM) image of the tip of a typical borosilicate pipette is shown. (B) Whole cell recording using a planar chip device. The back cavity of the chip is filled with electrolyte. Extracellular solution is applied to the chip surface where a droplet is formed due to surface tension. Cells in suspension are positioned and sealed onto the aperture by brief suction. No microscope or micromanipulator is needed. From (Fertig et al., 2002).

In planar patch-clamp, the planar insulating partition separates two chambers which are filled with saline. The partition contains a 1–2 µm aperture which is topologically equivalent to the tip of a glass pipette. A single cell is then positioned on the hole by

17 suction and a gigaseal is formed. A pulse of vacuum or voltage breaks the patch

membrane, thus establishing the whole-cell recording configuration.

The planar geometry provides a variety of advantages compared to the classical geometry. First, the planar patch-clamp configuration does not need an expensive microscope and micromanipulator setup. Second, a highly skilled operator is not necessary. Third, the planar electrode array is fabricated with planar fabrication technology as is used in microelectronic chip fabrication. The economies of scale make it possible to reduce the cost per electrode exponentially. Fourth, planar electrodes can be integrated with microfluidic lines which enables automatic compound application in

HTS. Lastly, the amplifier electronics can be potentially integrated with the electrode to improve noise performance.

2.3.1 Cell Guidance in Planar Patch-Clamp

How to direct a cell to a patch recording site has become a challenging issue in planar patch-clamp applications. It is well known that once a cell (or debris in the solution) seals to the tip of a glass pipette or to the aperture of a planar patch-clamp device, a residue, which is very difficult to remove, will stay at the hole and prevent the subsequent formation of another gigaseal. Therefore, all patch-clamp systems use disposable patch

interface, and many planar patch clamp systems are carefully designed to maximize the

chance of a successful first contact of a cell with the patch recording site. Commonly used cell-positioning methods include negative pressure (Stett et al., 2003), dielectrophoresis (Schmidt et al., 2000), and directed fluidic flow (Pantoja et al., 2004).

Figure 2.7 The CytoPatchTM chip. (A). Cross section. A cell is trapped by suction applied to the large port 1 of the device. Subsequent suction on the central port 2 forms a seal, and currents are recorded through port 2. (B). Scanning electron microscope (SEM) image of the device from above. (C). Cross section diagram of the device, showing the fluidic compartments. (D). Corresponding SEM cross-sectional view. (E). Cross section diagram of the packaged chip with fluidic ports. From (van Stiphout et al., 2005).

Negative pressure has been extensively applied to attract cells to the patch recording site in traditional planar patch-clamp techniques. Stett and his colleagues (Stett et al.,

19 2003; van Stiphout et al., 2005) have developed an elegant planar microchip (Figure 2.7).

This microchip has two integrated channels: inner channel for current measurement and outer channel for cell guidance. Positive pressure is initially applied in the inner channel to prevent debris from approaching its surface. Suction in the outer channel directs a cell to the top of measurement site, which looks exactly like the tip of a pipette. When a cell is placed at the measurement site, suction is used in the inner channel to encourage seal formation.

Schmidt and his colleagues (Schmidt et al., 2000) have developed an interesting dielectric approach to guide cells. Unilamellar lipid vesicles position themselves at the micrometer-sized holes in a planar insulating diaphragm by dielectric focusing. A potential difference of less than 200 millivolts is imposed across a SiO2 coated silicon

nitride diaphragm about 100 nm thick. The resulting inhomogeneous electric field guides

non-conducting particles (small vesicles) to reach the aperture where the voltage gradient

is the highest. The authors have demonstrated that stable gigaseals to the micro-meter sized holes can be obtained within seconds. For larger structures such as mammalian cells no progress has been reported.

Pantoja et al. (Pantoja et al., 2004) integrate PDMS microfluidics with a planar microfabricated silicon patch interface. The microfluidics is simple and useful to direct a cell to the aperture with laminar flow. Matthews and Judy (Matthews and Judy, 2006) also use integrated PDMS microfluidics to direct cells to the measurement site. Their fluidic system is comprised of six solution exchange lines (Figure 2.8). It will be very

intriguing if they demonstrate solution exchange with their microfluidic system in the

future.

Figure 2.8 Microfluidic chip for single cell patch-clamp measurement. (A). Overview of the eight-port microfluidic system. (B). Photograph of the macroscopic test fixture with eight capillaries connecting to the micromachined planar patch-clamp system. (C). Schematic diagram of a planar patch-clamp dose-response measurement system. From (Matthews and Judy, 2006).

Another strategy is not to guide cells at all. Several groups (Fertig et al., 2002;

Klemic et al., 2005; Xu et al., 2003) have developed chips where the cell seals to a simple

round aperture in a planar surface. Without special cell guidance, these systems use a

very dense and debris-free suspension of cells to increase the likelihood of a cell docking to the partition aperture. Suction through the aperture helps sealing of the cell to the

round aperture.

2.3.2 Planar Patch-Clamp Structure

The basic structure for planar patch-clamp is a micrometer-sized hole in a suspended insulating layer. The insulating layer could be glass-coated silicon (Asmild et al., 2003;

21 Matthews and Judy, 2006; Pantoja et al., 2004), silicon nitride (Fertig et al., 2000), glass

(Fertig et al., 2003; Xu et al., 2003), polyimide (Kiss et al., 2003), or PDMS (Klemic et

al., 2002; Klemic et al., 2005). In addition to this basic structure, a double channel

structure (Stett et al., 2003), a hollow nozzle structure (Lehnert et al., 2002), and a side-

trapping microchannel structure (Ionescu-Zanetti et al., 2005; Seo et al., 2004) have been

developed to explore the possibility of high throughput electrophysiology.

A simple aperture in silicon diaphragm has been developed by etching silicon wafer with a double side deep reactive ion etch (DRIE) process, and depositing plasma enhanced chemical vapor deposition (PECVD) silicon oxide layers to provide insulation

(Pantoja et al., 2004). Gigaseals and whole cell recordings from CHO K1 cells have been

obtained with these chips. However, the quality of the recordings was limited by the large capacitance between the aqueous solutions and the bulk silicon due to the relatively thin insulating layer. The gigaseal rate for these devices is also low, presumably due to the roughness of the DRIE etched sidewall. To enhance the gigaseal rate, Matthews and

Judy (Matthews and Judy, 2006) have improved the sidewall smoothness by growing wet oxide, stripping the wet oxide, depositing amorphous silicon, and growing wet oxide again (Figure 2.9). The roughness of the microfabricated hole is greatly smoothed, and hence gigaseals are obtained with these silicon chips. However, no whole cell recording has yet been reported with the improved design.

A micrometer-sized hole in an insulating diaphragm has been developed by Fertig et al. (Fertig et al., 2000). The self-supporting Si3N4diaphragm was formed by anisotropic

etching of silicon. The formation of gigaseals with these devices was not observed,

presumably because the Si3N4 membrane (120 nm) did not provide sufficient sidewall

22 area for forming the membrane seal and because Si3N4 is not a good material for the cell interface. Such devices also show a large capacitance (hundreds of picofarads) across the partition, therefore, the noise performance will be impaired even if gigaseals are formed.

Figure 2.9 Improving the smoothness of DRIE etched aperture. (A). A hole in silicon etched by DRIE. (B). A layer of silicon oxide is grown. (C). Strip silicon oxide. (D). Deposit a layer of amorphous silicon. (E). Grown silicon oxide. Scanning electron microscope (SEM) images of the hole is shown at each step. From (Matthews and Judy, 2006).

A double-channel structure has been developed to automate patch clamp measurements (Stett et al., 2003). The microchip is manufactured by using processes such as PECVD of SiO2, chemical mechanical polishing, RIE, plating and sacrificial

layer etching. This device has two integrated channels: inner channel and outer channel

(Figure 2.7). The inner channel, which forms the 1-2 µm patch-clamp aperture as shown, is fabricated using the technique of focused ion beam milling. Gigaseals have been obtained with 92% success rate. This design has been commercialized a parallel automated patch-clamp system (CytoPatchTM, Cytocentrics AG, Reutlingen, Germany).

Figure 2.10 A hollow SiO2 nozzle for planar patch-clamp measurement. (A) SEM image of a micromachined SiO2 nozzle(diameter 10 µm, height 6 µm). The hollow oxide tube was broken open by cleavage of the Si substrate. (B) Nozzle (diameter 4 µm) embedded in a larger hole. From (Lehnert et al., 2002).

24 Lehnert et al. (Lehnert et al., 2002) replaced the pipette tip with a micronozzle by

creating microstructured hollow SiO2 nozzles on a silicon wafer (Figure 2.10). The

fabrication involves DRIE, oxidation and silicon etching. A maximum seal resistance of

240 MΩ has been obtained. The authors attribute the reason of no gigaseal to the roughness of nozzle sidewall.

Figure 2.11 Side trapped patch-clamp array on a microfluidic platform. (A) Cell trapping is achieved by applying negative pressure to recording capillaries, which open into a main chamber containing cells in suspension. Patch clamp recordings are obtained by placing AgCl electrodes in each of the capillaries, as well as in the main chamber. The device is bonded to a glass coverslip for optical monitoring. (B) Scanning electron micrograph of three recording capillary orifices as seen from the main chamber. (C) Darkfield optical microscope image of cells trapped at three capillary orifices. From (Ionescu-Zanetti et al., 2005).

A lateral microfluidic trapping structure for patch-clamp measurement has been developed by Lee and his colleagues (Ionescu-Zanetti et al., 2005; Seo et al., 2004). The

authors molded a PDMS microfluidic chip from microfabricated silicon/SU-8 master.

Cells are trapped at the micro-sized lateral channels and gigaseals have been obtained.

This geometry (Figure 2.11) allows inherent microfluidic integration, high density of

25 patch sites, and the ability to do fluorescence measurement during electrical recording.

The success rate to form gigaseals at the lateral-wall aperture is only about 5%. However,

in view of this system’s easy integration with microfluidics, it becomes promising for

situations where a low-resistance seal is acceptable.

2.3.3 Materials for Planar Patch-Clamp

Various materials have been developed to make the patch-clamp interface to cells.

They include silicon nitride (Fertig et al., 2000), thermal oxide (Matthews and Judy,

2006), PECVD oxide (Pantoja et al., 2004), quartz glass (Fertig et al., 2003), borosilicate glass (Fertig et al., 2003), glass with proprietary coating (Xu et al., 2003), polyimide

(Kiss et al., 2003), treated PDMS (Klemic et al., 2002; Klemic et al., 2005), and untreated

PDMS (Ionescu-Zanetti et al., 2005).

The fabrication of silicon nitride, thermal oxide and PECVD oxide is compatible with standard microfabrication techniques. Successful gigaseals measured with silicon nitride partitions have never been reported. Nevertheless, gigaseals have been obtained from thermal oxide (Matthews and Judy, 2006) and PECVD oxide (Pantoja et al., 2004). The gigaseal rates measured with both oxides were low and not suitable for high throughput patch-clamp measurement.

As the only successful pipette material in conventional patch clamp, glass (including quartz glass, borosilicate glass etc.) is also the most successfully commercialized planar patch interface. However, the fabrication of glass is not compatible with traditional microfabrication techniques, therefore, the costly glass patch interface is not ready for high throughput screening.

Figure 2.12 Nanion chip processed with the single ion track etching (SITE) method. A planar glass substrate is locally thinned and exposed to a highly accelerated ion. During its passage, the ion damages the glass structure and leaves a latent track. This track exhibits an increased etch rate compared to the bulk substrate. Upon etching, a conical-shaped cavity originates along the track, which enables the fabrication of round, micron-sized apertures. From (Fertig et al., 2003).

The fabrication processes of commercial glass chips are proprietary. Figure 2.12 demonstrates the only published fabrication process to make the commercial glass chips

(Fertig et al., 2003). A 200 µm fused-quartz or borosilicate glass wafer is thinned locally to 20 µm thick by wet etching. The center of the thin part is then exposed to a

27 highly accelerated ion (Au+18). As it passes, the ion damages the glass structure and

leaves a latent track. This track exhibits an increased etch rate compared to the bulk

substrate. Etchant removes the damaged track quickly and generates a micron-sized

conical-shaped aperture with a round and smooth opening. A gigaseal rate of 50% has

been reported with these glass chips. Glass planar chips based on this general design are

made by Nanion Technologies and are used in single-well and 16-well recording systems

(Fertig et al., 2003).

A proprietary coating has been used to glass planar chips (SealChipTM) and greatly

enhances the ability to form gigaseals (Aviva Biosciences Corp., San Diego, CA). With

this coating, a gigaseal rate of 75% has been obtained (Xu et al., 2003). SealChipTM is

being used in the PatchXpress instrument (Molecular Devices Corp., Sunnyvale, CA),

which is a highly automated device that measures 16 whole-cell recordings simultaneously.

A different material to make the planar patch-clamp is polyimide (Kiss et al., 2003).

A two-dimensional 8 × 48 (2.25 mm spacing) well array (PatchPlate™) was developed for HTS by Molecular Devices Corp., Sunnyvale, CA. There is a tiny aperture in the polyimide bottom of each well. A cell seals to the polyimide aperture with a sealing resistance in the order of 100 MΩ (loose patch clamp). The system works in the perforated configuration. The leakage currents are substantial. Therefore this technology is used in primary screening when the resolution is not quite critical. A success rate of about 70% in practical drug screening is obtained.

28 Another polymer that has been utilized for planar patch clamp is PDMS

(polydimethylsiloxane) (Klemic et al., 2002; Klemic et al., 2005). PDMS is also well known as Sylgard 184 in the electrophysiology field. Its dielectric constant and loss factor are similar to that of glass. Once treated by oxygen plasma, the surface of PDMS will have a composition very similar to glass (Langowski and Uhrich, 2005). Fred J.

Sigworth’s group (Yale University) has taken advantage of these similar properties to replace the glass pipette with a planar patch clamp electrode (Klemic et al., 2002; Klemic et al., 2005). The currently used fabrication method applies a stream of air to define a 2

µm hole in a PDMS sheet. Subsequent oxygen plasma treatment of the cured PDMS forms a thin silica surface layer that is suitable for forming gigaseals with cells. Gigaseal and whole-cell recordings have been obtained with the micromolded aperture. The probability of successful recordings is relatively low, about 10% (Klemic et al., 2005).

However, because of the simplicity of the fabrication method and the low material cost,

PDMS patch electrode provides a promising way to reduce the cost for high throughput ion channel drug screening. Figure 2.13 presents the planar patch-clamp recording system.

Figure 2.13 Planar PDMS patch- clamp recording system. The PDMS electrode is filled with electrode solution, and mounted onto a 1.6 mm silver tube. The surface of the silver tube was converted to AgCl by soaking in Clorox. The top chamber (A) is screwed into place atop the electrode. A 10 nl volume of cells is dropped onto the aperture, and suction is applied via tygon tubing on the silver tube connected to a 10-ml syringe. From (Klemic et al., 2005).

29 Untreated PDMS is also a potential material for high throughput electrophysiology.

Ionescu-Zanetti et al. (Ionescu-Zanetti et al., 2005) have used untreated PDMS

microfluidic lateral channels as patch interface and have successfully obtained a 5% gigaseal rate.

2.4 Microfluidics

One advantage of using the planar patch-clamp configuration is that microfluidic

integration can minimize the capacitance and noise level of electrodes. Several research groups have tried to integrate microfluidics with single planar patch-clamp interfaces

(Asmild et al., 2003; Matthews and Judy, 2006; Pantoja et al., 2004). In their systems, microfluidics is used to fill the chambers separated by a planar patch partition and to transport cells to the patch-clamp recording sites.

Pantoja et al. (Pantoja et al., 2004) bonded PDMS microfluidic layers to each side of

a silicon planar interface which has a micrometer-sized pore. The microfluidics are

simple and useful to direct the flow to the hole with laminar flow. The authors

demonstrated whole cell recordings but without an extracellular electrode solution

exchange. Matthews and Judy (Matthews and Judy, 2006) integrated PDMS

microfluidics with a silicon based planar patch interface. Their fluidic system contains 6

solution lines, one cell inlet and one cell outlet (Figure 2.8). They have not reported any

whole cell recording results. Asmild et al. (Asmild et al., 2003) placed a planar silicon

chip (QPatchTM) into an assembly (QPlateTM). The assembly contains recording

electrodes, electroosmotic flow pumps, and flow channels allowing for application of

cells and also the rapid exchange of solutions (Figure 2.14).

Figure 2.14 A planar silicon chip based patch-clamp system (QPatchTM) in an assembly (QPlateTM). The assembly contains recording electrodes, electroosmotic flow pumps, and flow channels allowing the application of cells and also the rapid exchange of solutions. From (Asmild et al., 2003).

These devices have made fast fluid exchange for single electrodes possible. However, dense arrays of electrodes also need microfluidics to allow for common fluid lines.

These lines need, in turn, isolation valves to electrically isolate neighboring electrodes during measurement. The requirements of a patch clamp measurement demands that the

DC electrical isolation between neighboring electrodes be greater than about 10 GΩ.

PDMS is a kind of silicone rubber which consists of repeating -OSi(CH3)2- units. For chemical and biochemical applications PDMS has significant advantages such as easy fabrication, transparency, low cost and biocompatibility. Also, the use of PDMS elastomer for microfluidics has numerous advantages over silicon and glass. First, as a material, PDMS is optically transparent and compatible with many optical methods for detection. Second, it is compatible with biological studies because it is impermeable to water, nontoxic to cells, and permeable to gases. Finally, the major advantage of PDMS over glass and silicon is the ease of fabrication and its integration to PDMS, glass, silicon and other surfaces. This feature makes it very simple to fabricate multilayer microfluidic

31 channel structures. There are three ways to bond PDMS layers together. First, two partly

cured PDMS layers with different base-to-catalyst ratios can be placed together and bonded tightly after fully curing (Unger et al., 2000). Second, fully cured PDMS layers

can be bonded together after a brief oxygen plasma treatment (Xia and Whitesides, 1998).

Lastly, PDMS layers can be bonded together after stamping a curing agent or silicone

glue onto the surfaces (Satyanarayana et al., 2005).

As a fundamental element in microfluidcs, valves have been developed to realize

microfluidic controls in a Lab-on-chip. Valves can be actuated by pH-sensitive

hydrogels (Beebe et al., 2000), electrochemically generated microbubbles (Suzuki and

Yoneyama, 2003), thermally induced expandable microspheres (Griss et al., 2002) and most importantly, pressure (Unger et al., 2000).

Figure 2.15 A pneumatically actuated valve. (A). A simple mechanical valve fabricated using multilayer soft lithography, in which a fluid channel is separated from a control channel by a thin membrane. (B). When the control line is pressurized, the membrane is pushed up to close the fluid channel. From (Studer et al., 2004).

32 The elastomeric property of PDMS has been exploited to fabricate the pneumatically

actuated PDMS valve by Quake’s research group at Stanford University (Thorsen et al.,

2002; Unger et al., 2000). Unger et al.(Unger et al., 2000) fabricated one type of valve using a crossed-channel architecture. When pressure is applied to the bottom channel, the membrane at the crossover point deflects upward. Sufficient pressure closes the top channel (the flow channel). The shape of the flow channel is important for proper actuation of the valve. Flow channels with a round cross-section can close completely.

The valve opening can be precisely controlled by changing the pressure applied to the control line. Figure 2.15 shows how this pneumatically actuated valve works.

Whiteside’s research group at Harvard University has developed an elastomeric one- way diaphragm valve by assembling several pre-fabricated PDMS sheets together (Jeon et al., 2002). In their design, the membrane can only deflect in one direction. Flow in the opposite direction is blocked by the presence of a barrier. Figure 2.16 illustrates how the one-way valve works.

Figure 2.16 An elastomeric one- way diaphragm valve by assembling several pre- fabricated PDMS sheets together. (A). Flow from top channel to bottom channel is blocked by the presence of a barrier. (B). Flow from bottom channel passes through the valve as the membrane deflects. From (Jeon et al., 2002)

33 Pneumatically actuated valves have several advantages such as ease of fabrication, rapid response time, voidance of air bubbles, as well as scalability. Thorsen et al.

(Thorsen et al., 2002) have fabricated a device containing 2056 microvalves (Figure 2.17).

The technology has been commercialized by Fluidigm Corporation (San Francisco, CA).

Figure 2.17 Optical micrograph of the microfluidic comparator chip, containing 2056 microvalves, which is capable of performing more complex fluidic manipulations. In this case, two different reagents can be separately loaded, mixed pairwise, and selectively recovered, making it possible to perform distinct assays in 256 subnanoliter reaction chambers and then recover a particularly interesting reagent. From (Thorsen et al., 2002).

2.5 Planar Ag/AgCl Electrodes

The patch clamp measurement requires that metal electrodes interface the ionic solution and transform current smoothly from a flow of electrons in a metal wire to a

34 flow of ions in solution. Several types of electrodes are used in electrophysiological

measurements; the most commonly used is a silver/silver chloride (Ag/AgCl) interface

(Axon Instruments, 1993), which is a silver wire coated with silver chloride (Figure 2.18).

If electrons flow from the metal wire to the Ag/AgCl electrode, Cl- ions become hydrated

and enter the solution. If electrons flow in the reverse solution, Ag atoms in the electrode

give up their electrons (one electron per atom) and combine with aqueous Cl- ions to

make insoluble AgCl. Therefore, current can flow in both directions in Ag/AgCl electrodes. The operation of Ag/AgCl electrodes requires a solution containing chloride ions. A pair of Ag/AgCl electrodes is needed for the current to flow in a complete circuit.

Because current flow always induces the reduction of either silver or silver chloride, the

Ag/AgCl electrode is exhaustible. The exhausted electrodes may poison proteins and

have unpredictable junction potentials. Used properly, Ag/AgCl electrodes provide

predictable junction potentials and safety to the patch clamp measurement.

A simple way to fabricate Ag/AgCl electrodes in the laboratory is to place a clean Ag

wire in a saturated KCl solution and to pass positive current through this wire.

Electrochemically, a layer of silver chloride is generated on the surface of the silver wire.

Ag/AgCl electrodes are available commercially in different forms.

Experimental and theoretical studies of Ag/AgCl electrodes have been conducted by

various groups (Cranny and Atkinson, 1998; Jin et al., 2003; Temsamani and Cheng,

2001). It has been found that the potential drift of Ag/AgCl electrodes is dependent on

the Cl- concentration, pH value, the mixing/diffusion of the ionic solution, and the way

the AgCl coating is generated.

Figure 2.18 The Ag/AgCl electrode is reversible and exhaustible. From (Axon Instruments, 1993).

A planar patch-clamp electrode array needs planar Ag/AgCl electrodes.

Microfabrication techniques have been used to fabricate planar Ag/AgCl electrodes

(Suzuki and Taura, 2001). A 0.3 µm film of Ag is deposited on a U-shaped gold backbone pattern (Figure 2.19). The silver layer is passivated with polyimide that has four 10 × 10 µm open pinholes. The silver chloride starts to grow from the pinholes. A lifetime of 300 h is achieved using an electrode current of 10 nA. The AgCl layer can be regenerated and the exhibited potential and the lifetime of the element are reproducible.

Figure 2.19 Structure of the thin-film Ag/AgCl electrode. (a) conventional thin-film Ag/AgCl element (cross section), (b) top view of the novel thin-film Ag/AgCl element, (c) cross section along the line X-X’. From (Suzuki and Taura, 2001).

Summary

This chapter discusses current research status and the necessity to make a

microfluidic system for high-throughput planar patch-clamp measurement. Ion channels

are transmembrane proteins that allow ions to pass through cell membrane. Patch clamp

is the central technique to study ion channels. Planar patch clamp is a promising technology for high throughput screening. A microfluidic system with isolation valves is

needed to obtain simultaneous patch-clamp recordings from a high-density electrode

array. Pneumatically actuated valves based on multilayer PDMS technology make large-

37 scale microfludic integration possible, and have potential for high density patch electrode array application. The development of planar Ag/AgCl electrodes is also reviewed.

38 Chapter3:

Device Fabrication

In this chapter, we describe the fabrication of the microfluidic system integrated with disposable planar PDMS patch partitions, and planar Ag/AgCl electrodes. Figure 3.1

shows a schematic cross-section view of such a system.

Figure 3.1 Schematic cross-section view of the planar patch-clamp system. A dense suspension of cells is dropped onto the PDMS partition, which readily seals to the reusable microfluidic system.

39 3.1 A Disposable Planar PDMS Patch Partition

Micromolded PDMS proved to be a suitable material for planar patch clamp

partitions (Klemic et al., 2002; Klemic et al., 2005). PDMS has a similar dielectric

constant and loss factor to that of glass. Once treated by oxygen plasma, the surface of

PDMS will have a composition very similar to glass (Langowski and Uhrich, 2005). The

molding process makes it promising to fabricate PDMS partitions at a low cost. Here we describe the process of making these disposable patch partitions.

3.1.1 Partitions Molded with Microfabricated Silicon Master

In the early stage of this project, we molded the PDMS partitions with a microfabricated silicon master.

The microfabrication process flow to make the silicon master is shown in Figure 3.2.

After a 0.5 µm layer of silicon oxide was wet-grown on a silicon wafer with (100)

orientation, silicon nitride was deposited through the LPCVD (Low Pressure Chemical

Vapor Deposition) process and patterned to make squares with corner compensation

structures as a KOH etch mask. A timed etch of silicon in KOH (potassium hydroxide in

water, 30%) at 60 °C generated a mesa structure. The silicon nitride mask was then

stripped through RIE (Reactive Ion Etch). A timed silicon etch with oxide as mask in

DRIE (Deep Reactive Ion Etch) yielded 10 µm high pillars on top of a silicon mesa.

Those pillars had 2 µm or 4 µm diameter. The silicon oxide in the front side of wafer

was then stripped with hydrofluoric acid. Finally a 0.2 µm aluminum film was sputter coated on the front side of the wafer. This film promotes release of PDMS after curing.

The photomask for lithography was designed by Dr. James F. Klemic. The

40 microfabrication was accomplished at the Cornell NanoScale Science & Technology

Facility (CNF). Figure 3.3 shows SEM pictures of the microfabricated silicon master.

Figure 3.2 Process of fabricating silicon master. A 0.5 µm layer of silicon oxide is wet-grown on silicon wafer. A 0.2 µm layer of LPCVD nitride is deposited and patterned. A timed etch of silicon with nitride as mask generates a mesa structure. The silicon nitride mask is then stripped. A timed silicon etch with oxide as mask in DRIE yields pillars on top of the silicon mesa. The silicon oxide in the front side of wafer is then stripped with hydrofluoric acid. Finally a 0.2 µm aluminum film is sputter coated.

Figure 3.3 SEM pictures of a microfabricated silicon master. A pillar-on-mesa silicon structure is shown with different magnifications. The pillar shown has a diameter of 4 µm and a height of 10 µm.

PDMS partitions with with 2 µm or 4 µm apertures were then molded from the micromachined pillar-on-mesa master. A small quantity of liquid PDMS prepolymer was dropped onto the master and covered with a plastic coverslip or transparency film.

The surface tension of the PDMS prepolymer generates a uniform low vacuum pressure inside the liquid, which draws the coverslip to silicon masters and squeezes the prepolymer away where the silicon pillar contacts the plastic coverslip. After the PDMS was cured for 30 minutes in an 85 °C oven, the cover-slip was peeled away at 90 °C

42 ambient and a partition with 2 µm or 4 µm apertures was obtained (Figure 3.4). Figure

3.5 shows the optical microscope and SEM (Scanning Electron Microscope) pictures of

the molded PDMS partitions.

Figure 3.4 Molding PDMS partition from the microfabricated silicon master. PDMS prepolymer is placed on silicon master and covered with a plastic coverslip. After curing, the coverslip is removed and PDMS partition with holes is peeled off.

These PDMS partitions have the desired geometry (a tiny hole in a thin membrane followed by a pyramidal cavity), which is necessary to reduce the access resistance and capacitance for low noise patch clamp measurement. However, the bottom surface of the partition is not smooth because of the unevenness of the timed-etched silicon master. It was difficult to seal the partitions with the bottom measurement chamber. Therefore, we were not able to successfully test those partitions. Also the molding masters had a limited time of use due to the fragile nature of the silicon pillars. To overcome these problems, Dr. Kathryn G. Klemic came up with a novel and simple idea to mold PDMS partitions repeatedly (Klemic et al., 2005).

Figure 3.5 Micromolded PDMS partition from silicon master. (A) Top view of a sliced PDMS partition. (B), (C) Side views of a sliced PDMS partition. (D) A SEM picture of a molded aperture.

3.1.2 Partitions Fabricated with the Air-Molding Technique

A disposable planar PDMS patch electrode was micromolded using a micron-sized stream of nitrogen to define an aperture in the silicone elastomer, by the method of

Klemic et al (Klemic et al., 2005). However, instead of using a PDMS secondary support for the partition, here we use a glass washer as secondary support, which provides a robust support to the elastomeric partition and a surface that seals well to PDMS microfluidics.

Figure 3.6 Process of fabricating disposable planar PDMS patch partitions. A metal plate with an array of 2 µm holes is mounted in a chamber with a resistive heater and compressed air supply (A). The primary PDMS support washer is prepared by punching 13-gauge and 20-gauge needles into a 450 µm PDMS sheet. After being painted with PDMS prepolymer, the primary support is placed onto the metal plate and aligned carefully to the hole. The jet flow of compressed air defines a tiny hole in the PDMS as it is cured by heating (B). The glass washer is prepared by drilling a 1 mm hole in 0.15 mm thick glass disc. After curing, the PDMS partition is peeled off, flipped over and bonded onto the glass washer with fresh PDMS prepolymer. An SEM picture (C) shows that the air-blown hole has a smooth surface. Picture (D) shows the cross-section of the air-blown partition. Picture (E) shows two variants of patch partitions.

Figure 3.6 shows the procedure to fabricate an air-blown partition. Different base to catalyst ratio (10:1, 7:1, 5:1) PDMS partitions were made. Here a metal plate fabricated

45 with eight tiny holes was ordered from Dynamic Research (Wilmington, MA). The hole was defined by a high-aspect-ratio electroplating process and had a conical structure in the metal plate. The opening of the hole in the front side of the metal plate was 2 µm.

This metal plate was mounted in a chamber with six resistive (20 Ω) heaters and a compressed gas (air or nitrogen) supply. The resistive heaters were connected in parallel to a 15 VDC power supply. The gas supply was maintained at 20 Psi and there was a continuous gas flow through the hole. A PDMS primary support was cut from a 450 µm thick sheet. The inside hole of diameter 500 µm was punched by a gauge-20 needle, and outside diameter of 2.2 mm was cut by a gauge-13 needle. After the PDMS primary support was painted with liquid PDMS prepolymer, it was placed on the metal plate, with the inner ridge carefully aligned to the hole in metal plate under a stereo microscope.

The gas flow in the PDMS prepolymer formed a 2 µm diameter hole and a series of bubbles, of which the largest was visible under the stereo microscope. The thickness of the PDMS membrane was controlled by adding or reducing prepolymer with a single-hair brush so that the largest air bubble was about 100µm. The PDMS cured after a 5-minute heating and a 12-minute naturally cooling. Peeling off the PDMS partition and flipping it over yielded a partition with a 2 µm opening in the front side.

The glass washer secondary support was prepared by drilling a hole in a 0.15 mm thick glass piece. A glass piece could be a round (5 mm diameter) glass cover slip

(Electron Microscopy Science, Hatfield, PA) or a manually cut square (4 × 4 mm) glass slide. The glass piece was bonded to a 2” × 3” PDMS-coated (20 µm) glass slide before being drilled. Water was used as coolant in the drilling. A 0.75 mm Electroplated diamond solid drill bit (UKAM Industrial, Valencia, CA) was mounted on a drilling

46 machine (DELUXE Drill 212, DREMEL, Racine, WI) and was carefully pressed into the

center of glass piece to drill a 1 mm diameter through hole. After the hole was drilled, the glass washer was removed and rinsed with methanol and isopropanol. The glass washer was then ready for being bonded with the PDMS partition.

The cured PDMS partition was peeled off, flipped over, placed onto the glass washer support and aligned to the drilled hole with tweezers (Figure 3.6). PDMS prepolymer was painted to the edge of the partition with a single-hair brush carefully to seal the

PDMS partition to the glass washer support. The fully mounted electrode was then heat- cured in an 85 °C oven for 1 hour. An SEM picture (Figure 3.6C) shows a smooth air- blown aperture. The cross-section of the hole (Figure 3.6D) shows a series of bubbles in

PDMS reflecting the shape of air flow in the PDMS prepolymer. Figure 3.6E shows two variants of patch partitions.

This is a simple way to fabricate glass supported patch partition in a laboratory.

Using our current approach, one can fabricate ~60 PDMS partitions per day.

3.2 PDMS Isolation Valves

Dense arrays of electrodes need microfluidics to allow for common fluid lines. These

lines need, in turn, isolation valves to electrically isolate neighboring electrodes during

measurement. The requirements of a patch clamp measurement demands that the DC

electrical isolation between neighboring electrodes be greater than about 10 GΩ. In this

section we discuss the fabrication of microfluidic valves and how our valves fulfill the

dense array requirement.

3.2.1 Fabrication of PDMS Valves

47 The method that we used is similar to Unger (Unger et al., 2000). The fabrication of

a mold master was in a class 100 cleanroom environment (Yale University). A HTG

mask aligner (64-5X, Hybrid Technology Group, Inc. San Jose, CA) with 365 and 405

nm radiation lines and in constant power mode at 8 mW/cm2 was used. Customer-

designed transparency film masks were ordered from Silverline studio (Madison, WI)

with 3600 dpi resolution. The positive photoresist Shipley SPR 220-7i (Microchem,

Newton, MA) was patterned and reflowed at 110 °C to form a 15 µm high, smooth mountain structure on 3” silicon wafer. The wafer was then treated by

Trimethylchlorosilane (TMCS) for 1 minute, presumably forming a monolayer which promoted releasing PDMS. Liquid PDMS was poured onto the master and fully cured in a 65 °C oven before peeling off. The resultant PDMS layer is composed of a microchannel structure which is a high fidelity negative reflection of the photoresist pattern (Figure 3.7A).

Figure 3.7B shows the process of fabricating microfluidic valves. Three layers of

PDMS with different thicknesses of 3 mm, 30 µm, and 30 µm were molded from reflowed photoresist pattern. They were irreversibly bonded together after a brief (less than 0.5 minutes) oxygen plasma treatment (100 W, 700 mTorr, Anatech SP100 plasma system, Anatech, Springfield, VA). The device was then placed in a 65 °C oven overnight and the surfaces of channels recovered hydrophobicity as the small molecules

in PDMS migrate to the surface. Figure 3.7C shows a cross-section of a PDMS

microfluidic channel.

Figure 3.7 (A) PDMS channels molded from reflowed photoresist pattern. (B) Three layers of PDMS bonded together after oxygen plasma. Valves forms at the cross-over point of channels. (C) SEM picture shows the microchannel has a smooth cross-section, preferable for complete closure of valves.

49 At the cross-over point, channels in different layers form a valve. Figure 3.8 shows a

microfluidic device with 16 valves. A positive pressure in one channel presses the thin

elastomer film and pinches off the ionic solution in the other channel.

Figure 3.8 Sixteen valves of different dimensions are formed by combining channels with four different widths (50 µm, 100 µm, 200 µm, 300 µm).

3.2.2 Valve Isolation Resistance

Table 3.1 shows tests of isolation and threshold pressure in valves having 16 different combinations of dimensions. Channels were 50 µm, 100 µm, 200 µm, and 300 µm while the depth was always 15 µm. The thickness of the second PDMS layer was 30 µm. The maximum electrical resistance of the closed channel was measured from the current response to a 5 mV voltage step. The maximum resistance that can be measured by our set up is 10 GΩ. The threshold pressure to achieve “successful pinch-off” (resistance >

10 GΩ) was also recorded. The normal response time to pinch-off the valve was within

0.5 minutes at threshold pressure. When the pressure is higher than threshold pressure,

50 the valve can be actuated within seconds. Both “press-down” (top channels as control channels) and “push-up” (bottom channels as control channels) were tested and the same results were obtained. For channel widths of 200 µm and 300 µm, the success rate was

100%.

Table 3.1 Testing results of isolation resistance from 16 different dimensioned valves. The results are from measurement of 10 devices. Channels are 50 µm, 100 µm, 200 µm, and 300 µm. Both “press-down” (top channels as control channels) and “push-up” (bottom channels as control channels) were tested and the same results were obtained. The success rate (electrical resistance higher than 10 GΩ) is shown as well as the average threshold pressure to successfully close valves.

From Table 3.1 we see the elastomeric valves of 200 µm and 300 µm meet the requirement for high-throughput screening. However, it was easier to go back to the open state if both channels were 200 µm and if the top channel was used as the control channel. Therefore, in the final microfluidic system, we used valves composed with 200

µm channels and used the top channel as the control channel.

Figure 3.9 shows how the pressure change in one channel isolated the liquid containing blue dye in the other channel.

Figure 3.9 (A). Two micromolded channels form a valve at the cross-over point. Channel widths are 200 µm and channel depths are 15 µm. (B). The 35 mm long flow channel is filled with ionic solution (135 mM NaCl, 5 mM KCl, 2 mM CaCl2, 2 mM MgCl2, 10 mM HEPES, 5 mM Glucose, pH 7.4) containing blue dye and has a resistance of 3MΩ when the pressure in the control channel is zero. (C). The flow channel is closed when there is a positive pressure (10 Psi) in the control channel (filled with air in this example). The measured electrical resistance in this case is higher than 10 GΩ.

3.2.3 Valve Lifetime

The microfluidic system is intended to be reusable to minimize the cost per patch

recording. Hence it is necessary to find out the lifetime of the microfluidic valves and

how to prolong it.

Through electrical measurements of the fluidic valves, we observed that the resistance

decreased with time, presumably due to contamination from the ionic solution. After every six hours of continuous use, the valves needed to be cleaned. 36 valves were cleaned with six different protocols and lifetimes were measured (Figure 3.10). The longest valve lifetime occurred when the valve was cleaned with deionized water and solvent (acetone, methanol, or isopropanol) and then left in air overnight. In that case, the lifetime of our valve (when the isolation resistance became less than 10 GΩ) was one week.

Figure 3.10 Electrical resistance degradation of fluidic valves. 36 valves were tested to measure valve lifetime. The resistance was measured twice per day at about six hour intervals. At the end of the day, the channels were rinsed with one of six protocols listed.

3.2.4 Simulation of the Valve Deformation

The fluidic valve closes or opens as the thin elastomeric PDMS membrane deforms due to the pressure change in the control channel. It would be very interesting to simulate the valve deformation and get an idea about roughly how much pressure is needed to actuate the valve and to compare the simulation results with measurement results.

We used a model of a four-edge clamped membrane (Maierschneider et al., 1995) with uniform load to simulate the PDMS membrane deformation:

1.994(1− 0.271ν )Et P = D 3 , (1−ν )(L / 2) 4 Max

53 ⎛ πx ⎞ ⎛ πy ⎞ D = DMax cos⎜ ⎟cos⎜ ⎟ , ⎝ L ⎠ ⎝ L ⎠ where

P: Load pressure (Pa)

L: Membrane dimension (50, 100, 200, 300 µm in this case)

E: Young’s modulus (7.5MPa, from product data sheet) t: Membrane thickness (assumed to be of uniform thickness 15 µm)

ν: Poisson ratio (assumed to be 0.5)

D: Displacement (µm)

DMax: Maximum displacement (µm)

Figure 3.11 Normalized deformation of PDMS membrane with 4-edge clamped model.

The simulated deformation of the thin membrane is shown in Figure 3.11.

54 The threshold pressure calculated for valve actuation (DMax = 15 µm) and the measured pressure to close the valve (resistance > 10 GΩ) are shown in table 3.2.

The bonding between PDMS layers can not hold more than 25 Psi pressure. With less than 25 Psi pressure, we were not able to close the valves of 50 × 50 µm and 100 ×

100 µm. As we can see from table 3.2, the simulated result is about the same order of magnitude as the measured threshold pressure. The difference between them could be attributed by the simplicity of the model: (1). The model use a uniform membrane thickness; (2). The simulation uses a 4 - edge clamped model, and the actual situation is more like a 2 – edge clamped model; (3). For electrical resistance to be greater than 10

GΩ, the maximum deformation has to be more than 15 µm.

Table 3.2 Comparison of simulated results and measured threshold pressure.

A rather complicated simulation was made with the finite element method in order to represent the three-dimensional geometry of the microvalves and to solve the equations governing their deformation and closure (Studer et al., 2004). Their simulation results were consistent with their experimental results. However, the actuation pressure in both cases was only about half as the threshold pressure measured in our experiment. That is

55 due to two reasons: first, the PDMS we used (Sylgard 184) is tougher than the PDMS they used (RTV 615); second, a valve needs larger pressure to achieve complete electrical

isolation than just to be actuated mechanically.

Figure 3.12 Microfluidic device fabrication procedure. Four layers of PDMS with different thicknesses of 4 mm, 0.5 mm, 30 µm, and 60 µm are molded from patterns in reflowed photosensitive resist. Then they are cut, treated briefly with oxygen plasma, and bonded together to generate a monolithic fluidic device. Metal electrodes are formed by shadow evaporation of 50 Å thick nickel and 0.5 µm thick silver onto a cleaned glass slide. A coating of Spin-on-Glass is applied and cured. After the SOG is patterned with standard lithography, small droplets of bleach are applied onto the open windows to chemically react with exposed silver and generate a thin layer of silver chloride coating. After opening holes are punched, the PDMS monolithic piece is bonded onto the microfabricated Ag/AgCl electrodes with the help of UV ozone treatment.

56 3.3 Fabrication of PDMS Microfluidics

The process flow of making PDMS microfluidics is shown in Figure 3.12. Four layers of PDMS were molded individually from silicon wafers. The top layer of PDMS,

4 mm thick without microchannels, was cut to the desired shape for handling purposes.

The second layer, 0.5 mm thick, was molded with flow-control channels and vacuum- control channels. The 30 µm thick third layer, spin-coated at 2000 rpm for 1 minute, has electrode solution flow channels and vacuum suction channels. The 60 µm bottom layer, spin-coated at 1000 rpm for 1 minute, was molded from a plain silicon wafer. The top handle layer was bonded onto the second layer after half a minute oxygen plasma treatment (100 W, 700 mTorr, Anatech SP100 plasma system, Anatech, Springfield, VA).

After the two layers were irreversibly bonded together overnight, the monolithic PDMS was peeled off and holes were punched for fluid connection with a sharpened gauge 19 needle. The same process was repeated so that all four layers were bonded together and a monolithic PDMS microfluidic part was generated. A through hole was punched with a sharpened gauge 19 needle to make electrical connection between electrode solution and metal electrodes.

3.4 Planar Ag/AgCl Electrodes

A planar patch clamp system requires a planar Ag/AgCl electrode array to access the electrode solution electrically. Microfabrication is well-developed for patterning planar electrode structures. Integration with PDMS fluidics requires a layer of glass on top of the Ag/AgCl electrodes.

3.4.1 Fabrication of Ag/AgCl Electrodes

57 Figure 3.12 shows the process of fabricating planar Ag/AgCl electrodes with standard microfabrication technology. A 2” × 3” glass slide was cleaned by Piranha (H2SO4 (98% in water) : H2O2 (30% in water) = 3:1) for 10 minutes, blown dry under nitrogen flow, and baked at 200 °C for 10 minutes before being loaded into an e-beam evaporator. A shadow mask custom-designed from Fotofab Corp (Chicago, IL) was used to selectively deposit nickel (50 Å) and silver (0.5 µm) onto the cleaned glass slide. Spin-on-glass polymer (SOG 500F, Filmtronics, Butler, PA) was spin-coated (0.5 µm) on the glass slide and then cured in a tube furnace at 450 °C with a continuous nitrogen flow. Windows

(500×500 µm) were then etched open in the SOG layer with 10:1Buffered Oxide Etch

(BOE) with lithographically patterned Shipley 1813 resist (Microchem, Newton, MA). A clorox bleach droplet was used to react with the exposed silver for 20 minutes and generate a thin coating of AgCl on silver electrodes.

3.4.2 Lifetime of Planar Ag/AgCl Electrodes

Integrated with the microfluidic system, the planar Ag/AgCl electrodes should be reusable. It is also necessary for the electrodes not to be exhausted in the middle of patch clamp measurement. In this section we measured the liquid-metal junction potential of the electrodes over time and demonstrate that the planar electrodes have a long enough lifetime for patch clamp measurement.

Figure 3.13 shows the set-up to measure the lifetime of Ag/AgCl electrodes. The electrode solution is 135 mM NaCl, 5 mM KCl, 2 mM CaCl2, 2 mM MgCl2, 10 mM

HEPES, 5 mM Glucose, pH adjusted to 7.4 with NaOH. The bath is connected with the ground through a silver wire coated with AgCl. Two microfabricated Ag/AgCl

58 electrodes are connected to two amplifiers. The exposed Ag/AgCl window is 500 × 500

µm. A thin layer of AgCl is formed after treating silver metalization in clorox for 20 min; the Ag/AgCl total thickness is about 0.5µm. The potential of each electrode was measured while a DC current was passing through the electrodes.

Figure 3.13 Set up to measure the lifetime of Ag/AgCl electrodes. Two planar Ag/AgCl electrodes are connected with two amplifier signal inputs. The bath is grounded through a Ag/AgCl wire. The solution is sealed to minimize the evaporation. Constant current passes through the electrodes while the potential is monitored.

The reaction at the surface of the electrode is:

e- + AgCl (s) ⇔ Ag(s) + Cl- (aq)

Figure 3.14 Potential drift of microfabricated Ag/AgCl electrodes with time. (A) Potential drift measurement shows that Ag/AgCl electrode potential is stable for several days when current is 5 nA (to deplete Ag). (B) Potential drift measurement shows that Ag/AgCl electrode potential is stable for less than 0.5 hours when current is -5 nA (to deplete AgCl).

Figure 3.14 shows the potential vs time relation of four electrodes. (A) Potential drift measurement shows that Ag/AgCl electrode potential is stable for 100 hours when current is 5nA (in the direction to deplete Ag). Once silver is completely depleted, an oxygen bubble is generated through electrolysis and a potential jump is observed. (B)

When the current is -5nA (to deplete AgCl), the Ag/AgCl electrode has an effective lifetime <0.5 hr. This means that the clorox bleach only generates a very tiny amount of

AgCl on the silver surface. However, the electrode can be rechlorided electrochemically

60 between two measurements. Therefore, the microfabricated planar Ag/AgCl is suitable for the patch clamp measurement.

3.5 Assembly of the Microfluidic Device

The PDMS microfluidics and the planar Ag/AgCl electrodes were carefully aligned and bonded together after both being treated with UV ozone (T10X10/OES, UVOCS Inc.,

Montgomeryville, PA) for 2 minutes. Tin coated copper wire was glued onto the silver leads with silver conductive paste (Alfa Aesar, Ward Hill, MA). PE50 polyester tubing was plugged into the connection holes in PDMS. 5 minute two-component epoxy glue was used to promote the seal between tubing and PDMS. After curing, the epoxy glue was encompassed with PDMS prepolymer to ensure there was no leakage at the tubing connections. The fully assembled device was placed in 65 °C for one hour and was then ready to use.

Figure 3.15 A fully assembled microfluidic device for parallel planar patch clamp measurements. For each electrode there is a fluid inlet and a vacuum line. There is no connection between different electrodes. Eight patch partitions are mounted onto the PDMS chamber through reversible bonding.

Figure 3.16 A microfluidic device for simultaneous planar patch clamp measurement. A common fluid line is used for all electrodes isolated by valves. Eight patch partitions are mounted onto the PDMS chamber through reversible bonding.

Three kinds of devices were developed for different purposes. Figure 3.15 shows a fully assembled microfluidic device for parallel planar patch clamp measurement. For each electrode there is a fluid inlet and a vacuum line. There is no solution connection between neighboring electrodes. Single patch clamp measurement can be performed at each electrode without contaminating other electrodes. However, this kind of device is only suitable for a low-density electrode array because of the large number of fluid tubing connections. Figure 3.16 shows another microfluidic device in which a common flow line is used for eight electrodes. Valves are used between neighboring electrodes for electrical isolation. This device has a reduced number of fluid connection lines in

62 comparison with the previous mentioned device. Therefore, this device has potential application in a high density electrode array. Figure 3.17 shows a microfluidic device for electrode solution exchange. There are two independent electrodes on this device. For each electrode, there are two solution flow-in channels and one solution flow-out channel which works as a vacuum suction line as well. These channels form a “Y” shape. Each of these channels is controlled independently through a PDMS valve.

Figure 3.17 A microfluidic device for electrode solution exchange. “Y” shaped fluidic channels are used for solution exchange purpose. Two partitions are mounted on two parallel electrodes.

Summary

In this chapter we describe the process to fabricate a disposable patch partition, to fabricate a microfluidic device with isolation valves, to fabricate planar Ag/AgCl electrodes, and the process of assembly. A disposable patch-clamp partition is made by

63 bonding an air-blown PDMS partition to a small glass washer. It is placed onto the fluidic system having fluid exchange channels with isolation valves and Ag/AgCl electrodes. Fluid channels are molded from PDMS using microlithographically defined molds. Ag/AgCl electrodes are fabricated with standard microfabrication techniques. At the cross-over point, channels in different layers formed a valve. The suitability of

PDMS valves and microfabricated Ag/AgCl electrodes for patch clamp measurement are also studied.

64 Chapter 4:

Results and Discussion

In this chapter we discuss the patch-clamp measurement results with the microfluidic system. We used RBL-1 cells to test our patch-clamp system. The electrode solution exchange was also tested. We also tested the compatibility of our microfluidic system with commercial planar glass partitions.

Figure 4.1 A cartoon picture of the microfluidic system, not drawn in scale. Two electrodes are filled with ionic solution and isolated with valves. Two cells with ion channel proteins seal to the holes with the help of vacuum. The interior of one cell connects to amplifier through the ionic solution and metallization. The exterior solution (not shown) is grounded. The ionic current through the whole membrane is recorded.

In chapter 3 we discuss the process of fabricating a microfluidic system for planar patch clamp measurements. We have fabricated a microfluidic device for single patch measurement (Figure 3.15), a microfluidic device for simultaneous patch measurement isolated with fluidic valves (Figure 3.16), and a microfluidic device for electrode solution exchange (Figure 3.17). Figure 4.1 is a cartoon showing two simultaneous measurements isolated with a fluidic valve. Figure 4.2 shows the set up for patch clamp measurement with our microfluidic device. Similar to traditional patch clamp, the set up is on an air table in a Faraday Cage. However, the inverted microscope is replaced with an inexpensive stereo microscope; and the setup can be operated by less skilled people since no micromanipulator is used.

Figure 4.2 A simple setup for planar patch clamp measurement. No expensive inverted microscopes or micromanipulators are required.

66 4.1 Cell Culture and Preparation

RBL-1 cells were used to investigate the system’s suitability for patch clamp measurements. The cell line was maintained at 37 °C, 5% CO2 in 75 ml culture bottles containing Minimum Essential Medium (MEM), 1% MEM Sodium Pyruvate Solution,

1% MEM Non-Essential Amino Acids Solution, 1% Penicillin Streptomycin, and 15%

Fetal Bovine Serum.

4.2 Recording Solutions

The bath (extracellular) solution contained 130 mM KCl, 4.4 mM NaCl, 2 mM CaCl2,

2 mM MgCl2, 10 mM HEPES, 5 mM Dextrose, adjusted to pH 7.4 with NaOH; the electrode (intracellular) solution contained 130 mM KCl, 10 mM NaCl, 4 mM CaCl2, 2 mM MgCl2, 10 mM HEPES, 10 mM EGTA, adjusted to pH 7.4 with NaOH.

4.3 Harvesting Cells

Once RBL-1 cells reached 3.0×105 /ml in the culture bottle, they were transferred to a

15 ml centrifuge tube. The cells were spun down in an Allegra X-12 centrifuge

(Beckman Coulter, Fullerton, CA) at 500 rpm for 3 minutes and the supernatant was discarded. The cells were then re-suspended in 10 ml recording solution. After being spun down again at 500 rpm for 3 minutes, the cells were re-suspended gently in 100 µl recording solution and were ready to be deposited onto the planar patch electrode. For each recording, 5 µl of cell suspension was dropped onto the patch partition.

4.4 Recordings and Analysis

Because PDMS is permeable to air, all channels were filled with liquid before measurement. However, flow-control channels and vacuum-control channels were filled

67 with deionized water; flow channels and vacuum suction channels were filled with ionic electrode solution.

Planar PDMS patch partitions, treated in oxygen plasma, were mounted onto the

PDMS microfluidic device. The supporting glass of the PDMS patch electrode readily sealed with the PDMS microfluidic layer in a reversible way. This seal also formed an electrical barrier between the bath solution and electrode solution. Positive pressure (20 mmHg) on the flow channel was used to force the electrode solution to fill the cavity below the partition and the hole in partition as well. Then a droplet of bath solution was introduced with a syringe onto the top of the partition. The bath solution was grounded with an AgCl coated silver wire. The planar Ag/AgCl electrode was connected to the signal input of the amplifier (Multiclamp 700A patch clamp amplifier, Molecular Devices,

Sunnyvale, CA). The signal was recorded with pClamp8.1 acquisition software using the

Digidata 1322A interface (both from Molecular Devices).

Unlike the situation in a conventional patch clamp set up, cells could not be resolved in our stereo microscope. After 5 µl cell suspension was dropped onto the patch electrode, we waited 30 seconds and then applied suction. A very gentle suction was used (< 50 mm Hg) to draw a cell to the hole. The seal resistance was monitored from the pClamp8.1 acquisition software. If a gigaseal was observed, the suction was released and the recording protocol (voltage steps from -120 mV to 120 mV) started. The voltage steps in the protocol easily broke the patch and whole cell configuration was achieved.

68 Data were collected with a 1 kHz cut-off frequency and an output gain of 500 MΩ.

Neither leak subtraction nor series resistance compensation was implemented in the protocol.

4.5 Single Patch Electrode Measurement

A total of 278 PDMS patch partitions were tested on the PDMS microfluidics. To explore process parameters to optimize the gigaseal results (discussed later), the air- blown PDMS partitions were prepared with different ratios of resin to catalyst (10:1, 7:1, and 5:1); the patch partitions were baked for various times (0 – 7 days) in a 180 °C oven; and the patch partitions were oxidized for various times (2 – 30 hours). Because of the irreversible nature of patch seal, the patch electrodes were never reused.

Figure 4.3 Inward rectifier currents from RBL-1 cells recorded with the microfluidic system. Traces are voltage steps -120 mV to +120 mV. The holding potential is 0 mV. (A). Whole- cell current recording. Seal resistance was 1.8 GΩ before breaking the patch membrane to allow whole cell recording. (B). The voltage steps for recording. (C). The current-voltage relation of the same whole cell recording.

A whole cell recording from an RBL-1 cell is shown in Figure 4.3(A). The recording is the ionic current response to repeated voltage steps from -120 mV to 120 mV (Figure

4.3(B)). The reversal potential was zero since the intracellular and extracellular solution

69 had the same potassium concentration. Figure 4.3(C) shows the I-V curve of the recording, reflecting the overall behavior of thousands of endogenous inward rectifier potassium channels in the cell membrane. These channels do not pass current at positive voltage (Lindau and Fernandez, 1986), where the recording just shows linear leakage current. At negative voltage (intracellular potential relative to extracellular potential), the potassium channels are open and potassium conductance predominates.

Seal resistance Percentage (%)

> 5.0 GΩ 6.9

1.0 ~ 5.0 GΩ 16.8

100 ~ 999 MΩ 5.3

< 99 MΩ 45.8

Bad partitions (hole blocked etc.) 25.2

Table 4.1 Seal resistance of 131 measurements from RBL-1 cells. Partitions were made from 7:1 PDMS mixing ratio, baked for 48 hours at a 180 °C oven, and treated longer than 4 hours with oxygen plasma.

In order to obtain gigaseals, the PDMS partitions needed to be treated with oxygen plasma for at least 4 hours. After this treatment, the PDMS partitions were used within half an hour. Baking at 180 °C seemed to improve the gigaseal rate (for 7:1 and 10:1

PDMS ratios) and the ease of obtaining whole cell access. However, the tradeoff was that the 2 µm PDMS hole also tended to be blocked after high temperature baking, possibly due to thermal degradation of PDMS (Camino et al., 2002). Initial tests showed that the best yield of planar PDMS patch partitions occurred when the PDMS was mixed at a 7:1 ratio, baked 2 days in a 180 °C oven, and treated longer than 4 hours with oxygen

70 plasma. The gigaseal yield obtained was 24 % out of 131 partitions in this case. Table

4.1 shows the distribution of seal resistance of the 131 measurements.

4.6 Compatibility with Commercial Planar Partitions

To test the compatibility of our microfluidic system with commercial planar partitions, we made a small number of recordings with unpackaged glass planar partitions as used in the Nanion Port-A-Patch (courtesy of Nanion Technologies, Munich, Germany). These glass chips are similar to the glass support for our PDMS partition and therefore formed tight, reversible seals when placed onto the PDMS microfluidic system. To ensure the performance of these glass partitions, they were stored in the air-tight shipping package until used (~ 6 months) and they were tested within 3 days after the package was opened.

With the same cell suspension and experimental condition, we obtained three gigaseals out of 19 attempts, showing that our microfluidic system is also compatible with glass partitions.

4.7 Simultaneous Measurement Isolated by Microfluidic Valves

The advantage of using microfluidics is the ability to make simultaneous recordings.

This is made possible by the isolation of individual electrodes by PDMS pinch valves.

Figure 3.16 shows such a system with 8 PDMS partitions, having a common electrode solution inlet and outlet, but separate suction ports and Ag/AgCl electrodes. Eight simultaneous patch-clamp measurements should be possible in such a system. Two simultaneous measurements were done in our system since our existing current measurement system only has two parallel amplifiers. Figure 4.4(A) and 4.4(B) show simultaneous recordings from two RBL-1 cells isolated with microfluidic valves.

Figure 4.4 Simultaneous recordings from two RBL-1 cells isolated by fluidic valves.

To verify that the fluidic valve completely isolates two neighboring electrodes, a cross-talk test was performed between two neighboring electrodes isolated by a valve.

Figure 4.5(A) shows a whole cell recording of an RBL cell measured with one electrode.

Figure 4.5(B) shows the current response of the same cell when the voltage protocol was applied to the neighboring electrode. The capacitive transients show the start and stop time of the voltage steps in the neighboring electrode. From the current response we calculated the electrical isolation of the valve to be 10 GΩ.

Figure 4.5 (A). Whole cell recording of a cell in an electrode array. (B). The current response of the same cell when the voltage steps are applied only to the neighboring electrode.

4.8 Electrode Solution Exchange

A microfluidic system was fabricated for intracellular solution exchange (Figure

3.17). However, we found it very difficult to realize fluid exchange with such a system.

There are two reasons for this:

(1). Patch clamp seals lasted only a short time. For the RBL-1 cells we tested, the normal time was about 3 to 5 minutes if the pressure underneath the seal was zero. The seal broke immediately if the pressure underneath the seal was larger than +15 mm Hg.

The seal lasted less than 3 minutes if the pressure underneath the seal was less than -15 mmHg (vacuum). This means that we would have to exchange the solution in a few minutes with a small driving pressure.

(2). The smallest cavity under the partition (shown in Figure 4.6) is 1 mm in diameter and 0.5 mm depth with our current fabrication approach. Based on our observation that flow rate in the microfluidic channel was about 1 mm/s when the driving pressure was 15 mm Hg, it takes more than 8.3 minutes to exchange the solution in the cavity, which is much longer than a seal would last.

Figure 4.6 The cavity underneath the partition. It is difficult to exchange solution in the cavity in a brief time with a small driving force.

In order to solve this problem, the structure and hence the fabrication of the microfluidic system would have to be changed. A potential way is to fabricate the micron-sized aperture and the fluidic channel together since both can be molded together.

However, the fluidic channel can not be treated by oxygen plasma. Therefore, a new coating material or treating method has to be developed to promote the gigaseal rate of cell to the aperture.

4.9 Noise Comparison with Glass Pipette

The noise of the measurement signal from our patch-clamp system connected with

Axon 200B amplifier (Molecular Devices, Sunnyvale, CA) was calculated to be

− 2 2 1 ⎛ ⎞ −25 2 σ = ∑⎜i − i ⎟ = 3.2×10 Α , N −1 ⎝ n ⎠

where

74 σ: the standard deviation of current signal (A),

N: number of data points (900 in this case),

in: the measured current (A),

− i : the mean of current signal (A).

The noise from a glass pipette connected to an Axon 200B amplifier (Molecular

Devices, Sunnyvale, CA) was calculated from the power spectrum (measured by Jie

Zheng, unpublished data) to be:

1kHz σ 2 = S 2 ( f )df = 6.25×10−26 Α 2 , ∫ i 0

for Kimax glass without Sylgard coating, where

σ: the standard deviation of current signal (A),

f: frequency (Hz),

Si: the spectral density function of frequency ( A/ Hz ).

Hence, our patch clamp system is more noisy than a sophisticated traditional patch clamp. There might be two reasons for this. First, we used a long metal insulated wire between the electrode and headstage; this could introduce thermal noise. Second, above the silver line we have a suction channel which is filled with conducting solution; therefore, thermal noise from the conducting solution could be coupled capacitively into the silver line.

Figure 4.7 The capacitance of the electrode. Two cross-sections of one electrode are shown. There are three sources of capacitance: from the cavity to the bath (C1); from the cavity to the control channel (C2 + C3 + C4); from the silver metal line to the vacuum channel filled with ionic solutions (C5).

The capacitance of our planar electrode is also measured to be around 1 pF. This is about the same as that of a glass pipette. A theoretical estimation of the capacitance

(Figure 4.7) shows that there are three contributions: the capacitance from cavity to the bath solution (C1 ≈ 0.1 pF); the capacitance from the flow channel to the control channel through the closed valve (C2 + C3 + C4 ≈ 0.1 pF); and the capacitance from the silver pad to the solution-filled vacuum channel (C5 ≈ 0.95 pF). Therefore, metal line should be placed away from any fluidic channels so that the noise can be reduced.

76 Summary

In this chapter we discuss the patch-clamp measurement results with the microfluidic system. Gigaseal patch recordings from RBL-1 cells were obtained with a 24% success rate. Simultaneous recordings from valve-isolated electrodes were obtained. Our microfluidic system is also compatible with other cell interface partitions; we demonstrate success with glass partitions used in the Nanion Port-A-Patch system. These results demonstrate the potential of a PDMS microfluidic system for high density arrays of planar patch clamp electrodes for high throughput measurement of ion channel activity.

Electrode solution exchange was also tested with our system.

77 Chapter 5

Conclusions and Future Directions

5.1 Summary of the Key Accomplishments

A microfluidic system for planar patch-clamp electrode arrays has been developed.

The need for this microfluidic system grew out of the requirements for a dense electrode array for high-throughput screening of pharmaceutical compounds and functional analysis of ion-channel genes. The design and fabrication of the microfluidic system are described in this thesis. The microfluidic system is integrated with disposable planar patch partitions and planar Ag/AgCl electrodes. This microfluidic system allows simultaneous patch-clamp measurements.

5.1.1 Fabrication of Planar Patch Partitions and the Microfluidic System

A disposable planar patch partition is made by bonding an air-blown PDMS partition to a small glass washer. The PDMS partition has a 1-2 µm hole molded from a micron- sized air stream. The microfluidic system consists of fluid channels with isolation valves and planar Ag/AgCl electrodes. Fluidic channels are molded from PDMS using microlithographically defined molds. Fluidic channels of different PDMS layers form a valve at the cross-over point. The electrical resistance of the isolated valves is measured

78 to be higher than 10 GΩ, desirable for multi-electrode recording. The Ag/AgCl electrodes are fabricated with standard microfabrication techniques. The lifetime of the isolation valves and Ag/AgCl electrodes was measured.

5.1.2 Test of the Microfluidic System

RBL-1 cells were used to investigate the system’s suitability for patch clamp measurements. The gigaseal rate was 24 % for RBL-1 cells with our patch-clamp system. Simultaneous whole-cell recordings from RBL-1 cells have been obtained with the microfluidic system. The microfluidic system is also compatible with other cell interface partitions; we demonstrate success with glass partitions used in the Nanion Port-

A-Patch system. These results demonstrate the potential of a PDMS microfluidic system for high density arrays of planar patch clamp electrodes for high throughput measurement of ion channel activity.

5.2 Suggestions for the Future Work

Thus far, the high density arrays of planar patch clamp electrodes for high-throughput measurement of ion channel activity is still in the stage of experimental exploration.

There exist several issues need to be solved before the commercialization of the high density array.

The first issue is about the lifetime of the planar PDMS patch interface. In the current patch-clamp measurement system, the disposable PDMS patch interface needs to be used within half an hour after oxygen plasma treatment. This is due to the molecular

79 reorganization in PDMS. After half an hour, the surface of PDMS returns to a hydrophobic state and the chance of getting gigaseal is rare. Dr. Kathryn G. Klemic

(Yale University) has been using several kinds of chemicals to extract mobile small molecules from the PDMS partition surfaces so that the reorganization may happen more slowly. This idea would be a great breakthrough for the PDMS high-throughput patch- clamp screening if proven to work.

The second issue is how to reduce the complexity of the microfluidic system. The current microfluidic system uses a common fluid line to address several electrode chambers sequentially while valves are used to isolate electrodes afterwards. Even though this is a great breakthrough for multi-electrode measurements, the microfluidic system is still complex since each chamber is connected with a vacuum line. This is acceptable if there are only several electrodes on one device. However, this design is not suitable for high-throughput measurement which requires simultaneous measurement from hundreds of electrodes. It would be very intriguing to design and test a microfluidic system which uses a common vacuum line to actuate multiple electrodes simultaneously.

The third issue is to reduce air bubbles in the fluidic channel. PDMS is hydrophobic and permeable to air. Therefore it is very easy to trap air inside the fluidic channels.

However, any air bubble inside may induce undesired surface tension effects inside the channels. It would be simpler and easier to handle fluid and pull suction if the channel is hydrophilic. It is necessary to test whether the isolation valve still works when the channel inner surface is hydrophilic.

80 The fourth issue is the how to reduce the noise of the patch-clamp recording. The rms noise of the present system at 1 KHz bandwidth is 1.5 times larger than that measured with a conventional glass pipette (refer to section 4.8). There might be two reasons for this. First, we used a long metal insulated wire between the electrode and headstage; this could introduce thermal noise. Second, above the silver line we have a suction channel which is filled with conducting solution; therefore, thermal noise from the conducting solution could be coupled capacitively into the silver line. First reason could be reduced if we integrate the microfluidics with the amplifier array together in future. Dr. Kathryn Klemic and Fara Laiwalla in professor Sigworth’s group have been developing a digital amplifier array based on Silicon-On-Sapphire technology. As for the second reason, the microfluidics can be carefully designed so that capacitance between silver line and fluidic line may be reduced to minimum.

The last issue is to make an array of patch partitions. The current method for making patch partition is a simple one for laboratory usage. It is necessary to demonstrate that the fabrication of patch partition is scaleable to high volume production.

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