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Abstract
Microfluidic System for Planar Patch-Clamp Electrode Arrays
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.
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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
by
Xiaohui Li
Dissertation Director: Mark A. Reed
Dec 2006
iii
Copyright © 2007 by Xiaohui Li
All rights reserved.
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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,
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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
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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.
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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
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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
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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
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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
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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
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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
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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
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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
1
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
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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
3
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
4
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.
5
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.
6
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.
7
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).
8
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
9
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
10
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).
11
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
12
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 micro-
pipette (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
13
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.
14
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).
15
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).
16
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
17
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
18
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).
19
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.,
20
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.
21
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;
22
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
23
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).
24
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).
25
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
26
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.
27
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
28
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.
29
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).
30
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).
31
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
32
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).
33
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)
34
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
35
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.
36
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.
37
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-
38
scale microfludic integration possible, and have potential for high density patch electrode
array application. The development of planar Ag/AgCl electrodes is also reviewed.
39
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.
40
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
41
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.
42
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
43
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).
44
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.
45
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
46
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
47
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
48
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.
49
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.
50
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,
51
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.
52
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.
53
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:
34)2/)(1()271.01(994.1
MaxDL
EtPν
ν−−
= ,
54
⎟⎠⎞
⎜⎝⎛
⎟⎠⎞
⎜⎝⎛=
Ly
LxDD Max
ππ coscos ,
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.
55
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
56
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.
57
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
58
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
59
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)
60
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
61
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.
62
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
63
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
64
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.
65
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.
66
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.
67
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
68
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.
69
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
70
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
71
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.
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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.
73
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.
74
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
2252
2 102.31
1Α×=⎟
⎠⎞
⎜⎝⎛ −∑
−= −
−
iiN nσ ,
where
75
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:
2261
0
22 1025.6)( Α×== −∫ dffSkHz
iσ ,
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 ( HzA / ).
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.
76
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.
77
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.
78
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
79
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
80
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.
81
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.
82
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