UNIVERSITY OF CINCINNATI
Date: 8/01/2006
I, Adam Bange , hereby submit this work as part of the requirements for the degree of: Doctor of Philosophy , in: Chemistry ,
It is entitled: Development and Characterization of Miniaturized Electrochemical Immunosensors
This work and its defense approved by:
Chair: H. Brian Halsall ______Chair: William R. Heineman ___ Pearl Tsang ______Carl J. Seliskar _
PhD Dissertation Development and Characterization of
Miniaturized Electrochemical Immunosensors
Adam Bange
A dissertation submitted to the
Division of Research and Advanced Studies
of the University of Cincinnati
in partial fulfillment of the
requirements for the degree of
DOCTORATE OF PHILOSOPHY (Ph.D.)
in the Department of Chemistry
of the Colleges of Arts and Sciences
2007
Committee Chairs: Dr. H. Brian Halsall,
Dr. William R. Heineman
Abstract
The objective of this research was to improve the capabilities of immunosensing devices through miniaturization and using nanoscaled materials. Collaboration between the Chemical
Sensors Group and the UC Department of Engineering was established to develop sensors and sensing systems using modern microfabrication technology.
One facet of this collaboration included work to miniaturize a microbead-based sandwich immunoassay that was developed by previous group members. A procedure to fabricate poly(dimethylsiloxane) microfluidic devices was developed to control the movement of small amounts of immunoassay reagents and microbeads. The devices were then used to evaluate bead and fluid manipulation, and detection. While the electrochemical and fluorescence measurements required for an immunoassay were demonstrated, we were not able to efficiently manipulate microbeads due to their adsorption to the microchannel walls.
Research was then done to improve the detection step of a miniaturized immunoassay. A microbead based sandwich immunoassay was developed using an interdigitated array (IDA) electrode with nanoscale dimensions (220 nm electrode width, 620 nm gap). The IDA was fabricated using an electron beam lithographic lift-off technique. After an antibody-assisted capture of antigen using paramagnetic microbeads, a β-galactosidase labeled secondary antibody was used to convert p-aminophenyl galactopyranoside (PAPG) into the redox active p- aminophenol (PAP). Amperometric detection of PAP with IDA electrodes at +300 and -200 mV vs. a Ag/AgCl reference electrode was used to measure the result, detecting MS2 bacteriophage concentrations as low as 10 ng/mL.
iii The third research project focused on making a label-free immunosensor using an array of carbon nanotubes as an electrode and electrochemical impedance spectroscopy (EIS) for detection. Highly aligned multi-walled carbon nanotubes were grown by chemical vapor deposition using a metallic catalyst, Fe/Al2O3/SiO2, on Si wafers. The nanotube towers were removed from the silicon and cast in epoxy, then polished so that one end was exposed for electrical connection and the other used as the electrode array surface. The nanotubes were functionalized electrochemically to form carboxyl groups and then chemically conjugated to antibodies. EIS was used to directly monitor the antibody-antigen binding.
iv v Acknowledgements
Throughout my graduate career, I have benefited immeasurably from collaborations both within and outside of my research group. From the UC department of engineering, I would particularly like to thank Erik Peterson, Dr. Xiaoshan Zhu, and Dr. Yeoheung Yun for their contributions. Erik and his advisor, Dr. Ian Papautsky, taught me the basics of microfabrication while we were directly working together, then he continued to give me technical, scientific, and professional advice, computer help, and fantastic lunch conversations for my entire graduate career. My collaboration with Xiaoshan and his advisor, Dr. Chong Ahn, was a great learning experience for me, and I sincerely appreciate the long nights of hard work he put in to fabricate the IDA electrodes. Yun and his advisors, Dr. Mark Schulz and Dr. Vesselin Shanov, made the nanotube based biosensor project possible, with their innovative ideas, fast paced work schedule, and patience to pursue very hard problems, even when our experiments weren’t working out as well as we had hoped.
I greatly appreciate the support of my research group, and the efforts that everyone made to build a community atmosphere through parties, pot luck lunches, birthday lunches, etc. Also, several group members were important collaborators on my research projects. Irena Nikcevic played a critical role in the microfluidics project, particularly by handling the fluorescence measurements used to characterize microchannels and monitor bead adsorption. Dr. Jian Tu taught me how to efficiently do immunoassays, and was always available to help make measurements or troubleshoot when something was going wrong. I owe a special thanks to Dr.
Kevin Schlueter for sharing his insights concerning my research as well as chemistry in general, jobs, real estate, the environment, cars, computers, meteorology, rocket science, pharmaceuticals, the criminal justice system, geopolitics, firearms, botany, nuclear fusion, the energy industry,
vi natural disasters, and a few hundred other topics that he knows enough about to make encyclopedias obsolete.
None of my graduate school success would have been possible without the support of my research advisors, Dr. William Heineman and Dr. Brian Halsall. While their teaching, advising, and writing styles are very different, I knew I could always count on them for the finest advice concerning all aspects of my life as a graduate student. As a new graduate student, it can be very easy to take exceptional research direction for granted. I admit I didn’t realize how fortunate I was until I began to talk to other graduate students about the way that their labs were run, and saw their jealousy and disbelief when I described how accessible and helpful my advisors were, and how they made their students’ development a priority.
I will definitely miss Dr. Heineman’s patience, optimistic attitude, and commitment to his students now that I am leaving UC. He always made sure that I had plenty of opportunities to succeed, by doing things such as introducing me to new collaborators and giving me the opportunity to write review articles, and for this I am very grateful. I am equally grateful for Dr.
Halsall’s resolve to make sure that I make the most of my talents and resist settling for mediocrity. He taught me, by example, to constantly strive for improvement and to be critical of myself. I would also like to thank Dr. Carl Seliskar for his generous assistance and advice, as he was like a third advisor to me for much of my graduate career.
Finally I would like to thank my parents for teaching me the value of education, and supporting and believing in me for more than twenty years of school. I sincerely appreciate the efforts and sacrifices that they made so that I could have the education and experience to thrive at the graduate level. They, along with my brothers and sisters, cousins, aunts and uncles, friends, and especially my girlfriend Valerie, have been there for me when I was stressed, worn
vii out, and needing encouragement. I would especially like to thank my grandmother for her kindness and generosity, particularly during my final year of graduate school. This past year, despite being my busiest and most stressful, has been a pleasure.
viii Approval Form
Title Page
Abstract
Copyright Notice
Acknowledgements
Table of Contents
Chapter 1 Introduction to Electrochemical Immunoassay 1 1.1 Chemical Sensors/Biosensors 2 1.2 Immunoassay 4 1.2.1 Antibodies 5 1.2.2 Immunoassay Procedure 7 1.2.3 Labels 9 1.2.4 Bead-based Sandwich Assay 9 1.2.5 Electrochemical Detection 10 1.2.6 Label-Free Detection 14 1.3 Miniaturization 14
Chapter 2 Microfluidic Device for Bead-Based Immunoassay 19 2.1. Introduction to Microfluidics 20 2.1.1 Microfluidic Immunoassay 20 2.1.2 Microfluidic Materials 22 2.1.3 Microfluidics using Microbeads 25 2.1.4 Surface Modification 26 2.2. Fabrication of PDMS Microfluidic Device 28 2.2.1 Creation of the SU-8 master 28 2.2.2 PDMS casting 31 2.2.3 Bonding 32 2.2.4 Connection to Syringe Pumps 35 2.2.5 Proposed On-Chip Immunoassay 37 2.3. Compatibility of Standard Detection Methods with Microfluidic Device 38 2.3.1 Fluorescence Detection 38 2.3.2 Electrochemical Detection 39 2.4. Bead adsorption Study 41 2.4.1 Reagents 42 2.4.2 Equipment 42
ix 2.4.3 Bead Labeling Procedure 43 2.4.4 PDMS Sample Preparation 43 2.4.5 Surface Treatments 43 2.4.6 Data Analysis Procedure 44 2.4.7 Results and Discussion 44 2.5. Conclusions 47
Chapter 3 Electrochemical Immunoassays using Interdigitated Array 54 Microelectrodes 3.1 Introduction 55 3.1.1 Electrochemical Immunoassay Background 55 3.1.2 Miniaturized Electrochemical Immunoassay 56 3.1.3 Analytical advantages of Microelectrodes 57 3.1.4 IDA Electrodes 57 3.2 Characterization of Commercial IDAs 62 3.2.1 Materials 62 3.2.2 Characterization of IDAs 62 3.3 Immunoassay with Nanometer Scaled IDA 68 3.3.1 IDA Fabrication 68 3.3.2 Physical Characteristics of Nanoelectrode 69 3.4 E. coli Immunoassay 73 3.4.1 Materials 73 3.4.2 Buffer preparation 74 3.4.3 Preparation of Biotinylated Antibodies 74 3.4.4 Paramagnetic Bead-Based Immunoassay for E. coli O157:H7 75 3.4.5 Nano IDA Detection 76 3.5 MS2 Immunoassay 77 3.5.1 Materials 77 3.5.2 Preparation of the Immunoassay Buffers 78 3.5.3 Paramagnetic Bead-Based Immunoassay for MS2 78 3.5.4 Nano IDA detection 79 3.5.5 Immunoassay results 79 3.6 Conclusions 83 3.6.1 Problems Associated with the Microscale 83 3.6.2 E. coli Assay 85 3.6.3 MS2 Assay 85 3.6.4 Future outlook 86
Chapter 4 Carbon Nanotube-Based EIS Biosensor 91 4.1 Introduction 92 4.1.1 Chemical Structure of Carbon Nanotubes 92 4.1.2 Physical Properties of Nanotubes 94 4.1.3 Electrical Properties 94 4.1.4 Sensing Applications 95 4.1.5 Electrochemical Impedance Spectroscopy 95 4.1.6 Goals of This Work 98
x 4.2 Materials/Methods 100 4.2.1 Synthesis of Nanotubes 101 4.2.2 Biosensor Fabrication 101 4.2.3 Immobilization of Antibody 103 4.2.4 Electrochemical Analysis 106 4.3 Results/Discussion 106 4.3.1 Nanotube Characterization 107 4.3.2 Electrochemical Analysis 110 4.4 Conclusions/Future Directions 115
Appendix A 123
xi
Figure and Table Legends
Chapter 1 Figure 1.1 Elements of a chemical sensor. Figure 1.2 Diagram of antibody molecule Figure 1.3 Typical dose-response curves for competitive and noncompetitive immunoassays. Figure 1.4 Alkaline phosphatase catalyzed conversion of PAPP to PAP. Figure 1.5 Oxidation of PAP to form PQI.
Chapter 2 Figure 2.1 Diagram of SU-8 master on which PDMS was cast. Figure 2.2 Steps of PDMS device fabrication. Figure 2.3 Photographs of PDMS microfluidic device. Figure 2.4 Proposed immunoassay setup for PDMS microfluidic. Figure 2.5 Calibration curve for fluorescein in PDMS microchannel. Figure 2.6 Log-Log Calibration Plot for PAP in PBS pH 7, 50 µL/min flow rate in Kapton fluidic. Figure 2.7 Effect of flow rate on the measured current of 1mM ferricyanide in 0.1 M KCl Figure 2.8 Comparison of the ability of different surface treatments to resist the adsorption of microbeads. Figure 2.9 The water contact angle is a measure of surface hydrophobicity/hydrophilicity. Table 1 Summary of PDMS bonding Table 2 Contact angle measurements on treated PDMS surface
Chapter 3 Figure 3.1 Redox cycling of p-aminophenol (PAP) and p-quinone imine (PQI). The PAP is produced by the enzyme β-Galactosidase on the substrate p-aminophenyl galactopyranoside (PAPG) Figure 3.2 Light microscopy images of 5 µm gap IDAs. Figure 3.3. CV with Ferrocyanide on 5µm IDA electrode, 100 mV/sec. Figure 3.4. Close up of 100 mV/sec CV on 5 µm IDA with 10 mM concentration traces removed to show detail. Figure 3.5. Calibration plot for PAP on 5 micron IDA using anodic current. Figure 3.6 Calibration plot for MS2 bacteriophage using 5 µm IDA for detection. Anodic current was used for the measurement. Figure 3.7 SEM images of nanoIDA. Figure 3.8 SEM image of IDA showing the slight misalignment of electrode fingers. Figure 3.9 E. coli detection with nanoIDA. Figure 3.10. Diagram showing how immunoassay measurements were made on the IDA. Figure 3.11. Plot of the current response as a function of time for immunoassay samples containing 0(a), 10(b) 50 (c) and 100 (d) ng/mL of MS2 phage. Figure 3.12. Calibration curve for MS2 using nanoIDA. Figure 3.13A. Damaged IDA electrode. Figure 3.13B Close-up of damaged region of IDA electrode. Table 1. Comparison of nanoIDA results to previous work with the same MS2 assay.
xii
Chapter 4 Figure 4.1 Diagram showing different ways of “rolling” graphene to form nanotubes, showing a) armchair, b) chiral, and c) zig-zigzag conformations. Adapted from Saito et al. Figure 4.2 Typical Nyquist plot for sensing applications. Figure 4.3 Schematic of the nanotube immunosensor (not to scale) Figure 4.4 Advantage of using nanotube composite for electrode material over bulk surface. Figure 4.5. Steps of biosensor fabrication. Figure 4.6. Coupling strategy used to attach antibody to carboxyl groups on exposed nanotube surface. a) EDC is used to activate carboxyl group, b) unstable acyl-isourea intermediate is formed, c) Sulfo-NHS displaces the EDC to form stable intermediate, and d) stable amide bond is formed to amine group on antibody. Figure 4.7. ESEM images of (A) aligned high density MWCNT patterned arrays on Si substrate, (B) side view of patterned nanotube array, (C) high resolution view of individual nanotubes in high density array. This array was fabricated with the growth conditions of: 200 SCCM of H2 flow, 200 SCCM of C2H4 flow, 100 SCCM bubbler flow, 750º C growth temperature. Figure 4.8. Wired nanotube tower-epoxy electrode. Figure 4.9. Cyclic voltammetry of 6 mM K3Fe(CN)6 in 1.0 M KNO3 with nanotube tower electrode with 100 mV/s scan rate: after (a) functionalization of nanotube array, (b) immobilization of antibody and (c) antigen binding Figure 4.10. Electrochemical impedance spectra of immunosensor response with the addition of different concentrations of antigen (inner to outer curve); (a) without antigen, with (b) 500 ng/mL, (c) 1µg/mL, (d) 5 µg/mL, (e) 10 µg/mL, and (f) 100 µg/mL of antigen. Figure 4.11. Calibration plot for mouse IgG on the impedance sensor. The Z values used for this plot were obtained by taking the square root of the sum of the squares of the real and imaginary components of the impedance at a frequency of 0.1 Hz (the lowest frequency measured). Figure 4.12. Electrochemical impedance spectra for a nanotube immunosensor with (a) donkey anti-mouse IgG immobilization, and mouse IgG binding after the incubation for half (b), one (c), two (d) and four (e) hours incubation (inner to outer curve) EIS is done at DC potential 0 V with frequencies between 0.1Hz and 300KHz. Sinusoidal potential magnitude is ±20 mV in 5 mM K4[Fe(CN)6], K3[Fe(CN)6] with PBS (pH 7.0). Table 4.1. Comparison of single walled and multi walled nanotubes. Table 4.2 Rejected host materials
xiii
Chapter 1:
Overview of Chemical Sensors and Electrochemical Immunoassay
1.1 Chemical Sensors/Biosensors
Chemical sensors are devices that are used to detect or measure the amount of a particular analyte, and are ubiquitous in the modern technological age. Common
examples include the oxygen sensor that controls the fuel-air mixture in modern internal combustion engines, as well as the monitoring kits used to control pH, chlorine, and
water quality in swimming pools. The medical field depends on chemical sensing to
diagnose and treat disease, and the closely related pharmaceutical industry makes
extensive use of chemical sensors to design, synthesize, and test therapeutics. Other
important applications of chemical sensors include, but are not limited to, environmental monitoring, military/homeland security, industrial quality control, agriculture, food production, and national defense. This reliance on sensors and sensing procedures drives the research to come up with faster, more reliable, cheaper, more accurate, longer lasting, easier to use devices and procedures.
The function of a chemical sensor can be three main steps, as illustrated in Figure
1.1. The first step is a molecular interaction that is somewhat specific to the analyte.
This interaction is typically a binding interaction with a probe molecule, but can also be another type of specific molecular action such as selective uptake into a film or diffusion through a membrane. The second step is signal transduction: the conversion of the molecular interaction into a measurable quantity such as an electrical signal or a color change. Typical signal transduction procedures include optical, electrochemical, acoustic, piezo-resistive, and radioactive decay measurements. Development and
2 optimization of this step of the sensing procedure is the primary focus of this dissertation.
The final step of the sensing procedure is converting the measured signal into a readable output. In recent years, this step has been somewhat trivialized by the advanced hardware and software instrumentation that converts the electronic signal generated by instruments into a digital format that can be manipulated by a computer.
Molecular Recognition Signal Transduction Readable Output
Figure 1.1 Elements of a chemical sensor.
There are many different types of chemical sensors of varying complexities.
Some sensors can be used for continuous monitoring, and give real-time information about the concentration of their target analyte. Examples include pH and ion selective electrodes, gas sensors, and some optical flow cells. These sensors, while easy to use, potentially inexpensive, and fairly sensitive, are generally only available for a limited number of target analytes, typically simple molecules that are found in relatively large
3 amounts in sample matrices with few interfering species. For more difficult analytes and
samples, separations, preconcentration, labeling, and other techniques are required to
achieve useful analytical results. As a result, a laboratory and skilled scientists may be
required to make measurements, making the analysis slower and more expensive. The challenge addressed in this work is to make the detection of more complex samples more like the pH electrodes and oxygen sensors, while still retaining the sensitivity and selectivity of the laboratory procedures.
Perhaps the most difficult types of chemical sensors to develop are biosensors.
Specific biomolecules typically are difficult to detect because they are generally found in a matrix surrounded by other biomolecules with similar chemical and physical properties.
To make matters more difficult, the detection levels required to give relevant results are frequently nanomolar or below. This limits direct detection techniques such as fluorescence, electrochemistry and absorbance, because there are so many potential interferences that can completely overwhelm the small signal generated due to the analyte. In order to make sensitive measurements in such complex samples, molecular recognition must be used. Examples of molecular recognition are the interaction of a receptor with its ligand, a nucleic acid chain with its complementary strand, and an antibody with its target antigen. A biosensor is a transducer that takes the molecular interaction between biological molecules and turns it into a detectable or quantitative signal.
1.2 Immunoassay
4 Immunoassay is a physical or chemical sensing procedure that uses the affinity and selectivity of an antibody to detect and quantitate the amount of antigen in a sample.
All the work discussed in this dissertation will be directly or indirectly related to the improvement of some facet of immunosensing. In particular, strategies to improve the sensitivity, speed, cost, and reliability of existing immunoassay procedures through miniaturization are the primary focus.
1.2.1 Antibodies
The remarkable analytical performance of immunoassay procedures is made possible by the highly specialized structure of antibodies. Antibodies, also called immunoglobulins, are large glycoproteins that are produced by the immune system for the purpose of recognizing foreign materials such as bacteria and viruses. In mammals there are five main types of immunoglobulins, designated as IgA, IgD, IgE, IgG, and
IgM. These different forms differ in structure, function, and location in the body.
Immunoassay procedures generally make use of IgG type immunoglobulins, as they are the easiest to collect and purify.
The structure of an IgG molecule is shown In Fig 1.2. The Y-shaped structure consists of four polypeptide chains: two heavy chains and two light chains joined together by disulfide bonds. In mammals, the heavy chains are roughly 500 amino acid residues long, and the light chains are slightly more than 200 residues long. The stem of the “Y” shape is known as the Fc region, for crystallizable fragment. This part of the molecule is constant in structure. The two Fab regions of the antibody, or antigen binding fragment, consist of a structurally conserved region and a hypervariable region at the very
5 tip of each arm of the “Y”. The amazingly diverse number of possible structures of this
region is what allows antibodies to be selective for a particular structural element on the
analyte.
Antigen binding sites
Light Chain
Disulfide bonds Heavy Chain
Figure 1.2 Diagram of antibody molecule
Antibodies that are used in immunoassay are usually generated in one of two ways, and the type of antibody produced can be important for different applications. The first type of antibody, polyclonal, is created by immunizing an animal, typically a mammal, and then collecting the antibodies from the serum produced against the specific antigen. Antibodies produced in this manner, while specific for a target antigen, do not
6 necessarily recognize the same epitope, as their binding sites may bind to different
regions of the target molecule or organism.
Monoclonal antibodies, on the other hand, are cloned from a single cell line of B-
cells fused to tumor myeloma cells. Being structurally identical, monoclonal antibodies
will bind to the same epitope of the target antigen.
1.2.2 Immunoassay Procedure
Immunoassay procedures can be divided into several different categories based on the way that the antibody-antigen binding event is transduced into a detectable signal.
Competitive assays are based on the competition for binding sites by a known amount of
labeled antigen and the sample. Because the number of antigen binding sites is roughly
constant, the concentration of sample analyte bound will be inversely proportional to the
amount of labeled analyte. Quantitation of the labeled analyte can thus determine the
concentration in the sample. The other class of assays is noncompetitive. The relationship
of signal to analyte concentration for competitive and noncompetitive can be seen in
Figure 1.3. Nearly all noncompetitive assays are sandwich assays. In sandwich assays,
two antibodies are used, a capture antibody and a detection antibody. The capture
antibody is attached to a solid support, so that when the target antigen is bound, a rinse
step can be done to remove molecules that are not recognized by the antibody. In order
to detect the bound antigen, a stoichiometric excess of labeled secondary antibody is
added. This antibody binds to the captured antigen, and the amount bound is directly
proportional to the amount of analyte captured. The amount of antigen is thus directly
related to the amount of label detected.
7
Figure 1.3. Typical dose-response curves for competitive and noncompetitive immunoassays.
Competitive and noncompetitive assays have their strengths and limitations.
Competitive assays are often simpler and faster, as they require fewer steps. Competitive assays are very common in applications such as clinical diagnostics and high throughput screening, where speed and efficiency are essential. The two-step, non-competitive, sandwich immunoassay is often considered to be the most sensitive and selective procedure, as the critical rinsing steps create a very low background, and multiple molecular recognition steps maximize specificity.
Other ways that immunoassay types can be classified are homogeneous and heterogeneous. Homogeneous assays do not contain a separation step to isolate the antibody-antigen complex, and are typically easier and faster to do. Heterogeneous assays use a solid support to separate the bound antigen from the sample, and are therefore more sensitive [1].
8 1.2.3 Labels
A good label for immunoassay is a molecule that can be detected in miniscule amounts with minimal interference from the background of the sample or environmental effects such as photobleaching. The first labels developed for immunoassay were radioisotopes, which satisfy these criteria very well [1, 2]. However, radioisotopes have significant shortcomings, principally the hazards of working with radioactive material, and the cost of its disposal. Alternative labels were therefore developed such as metals, nanoparticles, enzymes, and optically active and electroactive labels [3]. Enzyme labels are very useful because they can amplify the signal by converting thousands or even millions of substrate molecules into detectable product.
1.2.4 Bead-based Sandwich Assay
One type of heterogeneous immunoassay uses microbeads as the solid support for the capture antibody [4]. Beads are useful because they can dramatically increase the capture surface area in a small fluid volume, and when dispersed, reduce incubation times by shortening the diffusional distances. Very rapid turnover times can be obtained if beads are pre-loaded with antibody, and replaced in the immunoassay vessel with a fresh aliquot of beads between assays. This strategy also provides a highly reproducible control of the antibody load being used [5]. Often, polystyrene microbeads are made with an iron core so that they are paramagnetic, and can be manipulated with magnetic fields, allowing the beads to be immobilized and released at different stages of the assay. This is potentially advantageous in sampling from a large volume. The beads can be dispersed in the large sample for the capture step (in a sandwich assay, for example), collected by a
9 magnet, and transferred to a small detection volume thus providing increased sensitivity from the preconcentration. The beads are commonly available with reactive functional groups, such as amines, on the surface so that antibodies or other molecules can be attached. Beads are also available with a streptavidin coating, allowing a biotinylated molecule to be immobilized easily.
1.2.5 Electrochemical Detection
Of the various detection methods associated with different modes of signal transduction that have been developed in immunoassay techniques to recognize an antibody/antigen binding event, electrochemistry has advantages that have made it one of the most commonly used methods . In addition to high sensitivity nearing that of fluorescence in many situations, electrochemistry is not affected by the turbidity or light absorption characteristics of solutions in which the analyte resides, which would otherwise interfere with optical detection. In addition, electrochemical instrumentation can be easily miniaturized, giving rise to a system requiring low operating power. The synergy of all these features thus makes electrochemistry suitable for miniaturized detection, particularly when compactness and portability are important.
Unless the analyte is electrochemically active in a reasonable potential range, a redox-active label must be used for detection. A good redox label is a molecule that is easily oxidized or reduced at a potential where interfering species are not likely to be affected. Enzyme labels can also be used for electrochemical detection, because they can convert a substrate that is inert in the desired potential range into a product that is electrochemically active. Depending on the velocity of the enzyme catalyzed reaction,
10 thousands or even millions of molecules can be converted per minute, greatly amplifying
the signal [1]. Some examples of enzyme labels that have been used for electrochemical
immunoassay are: horseradish peroxidase, glucose oxidase, alkaline phosphatase, β-D-
galactosidase, glucoamylase, catalase, urease, glucose-6-phosphate dehydrogenase,
malate dehydrogenase [6]. A common enzyme label used in our laboratory is alkaline
phosphatase. In the presence of the substrate, para-aminophenyl phosphate (PAPP), the
enzyme reaction seen in Figure 1.4 yields the product para-aminophenol (PAP).
PAPP PAP
O- Alkaline Phosphatase
- O P O NH2 HO NH2
O
Figure 1.4. Alkaline phosphatase calalyzed conversion of PAPP to PAP.
When an oxidizing potential is applied, typically around +300 mV vs Ag/AgCl, PAP is
oxidized to form para-quinone imine (PQI), as seen in Figure 1.5. The resulting
oxidation current can then be quantitatively related to the amount of antigen analyte
bound to the secondary antibody-enzyme conjugate.
11 P A P P Q I
-2e-, -2H+
H O N H2 O N H
Figure 1.5. Oxidation of PAP to form PQI.
Most electrochemical measurements are either potentiometric or voltammetric.
Potentiometric measurements are made by measuring the potential with no current flow,
while voltammetry measures the current in response to an applied voltage. Voltammetry
is typically the electrochemical technique used for detection in immunoassays. In
particular, amperometry, a form of voltammetry where the steady state faradaic current resulting from a fixed potential is measured, is most commonly used to detect the
electrochemically active enzyme product in electrochemical immunoassays.
The signal generated from amperometric detection depends on the number of
redox active molecules that are transported to the electrode surface. Such molecules
dissolved in solution can be brought into contact with the electrode surface principally by
diffusion, migration, and hydrodynamic flow.
Diffusion is a phenomenon based on the random movement of inert particles that
causes a system to decay to a state of maximum uniformity, and can be represented
mathematically by Fick’s second law:
12 where is the concentration of the diffusing molecule, t is time, D is the diffusion
coefficient, and is the gradient operator. The observable effect of diffusion is that
particles move from areas of high concentration to areas of low concentration. In the
context of amperometry, an electron transfer occurs at the electrode surface, creating a
concentration gradient. In the case of an oxidation, the oxidized species diffuses away
from the surface, while more of the oxidizable species diffuses to the electrode surface.
The current that results from this system, as long as the bulk concentration does not
change significantly, is known as the limiting current.
Migration is the movement of charged particles (ions) due to the influence of an
electric field. Positively charged particles are attracted to a negatively charged electrode,
and vice versa. While this effect is very important in some situations, it is generally undesired in most electrochemical measurements, as it introduces an additional source of
signal variation. The effects of migration are often reduced by adding an inert
electrolyte, often called a supporting electrolyte, in high concentration relative to the
analyte to decrease the electric field strength near the electrode.
Hydrodynamic mass transport is caused by the movement of solution, typically
by stirring, flowing solution over the electrode, or rotating the electrode within the
solution. Electrochemical detection is very sensitive to movement of the solution over the
electrode because convection can be orders of magnitude faster than diffusion. While
this is a limitation in some situations, controlled flow and diffusive mixing can be used to
greatly enhance the detectable current by increasing the flux of redox active molecules to
the electrode surface. Electrochemical assays typically use stirring, flow injection
13 analysis [7], or rotating disk electrodes [4] to increase the observed limiting current over
that of plain diffusion.
1.2.6 Label-free Detection
Another class of immunoassay does not involve a label at all, but directly
monitors the binding of antibody to target antigen. Some examples of these label-free
techniques are micro- and nano-cantilevers [8], surface plasmon resonance [9], quartz
crystal microbalances [10], piezoelectric crystals [11], and electrochemical impedance spectroscopy [12]. Label-free techniques have the advantage of monitoring binding
directly, thus eliminating many time-consuming and potentially artifact inducing steps.
The development of a label-free immunosensor that makes use of electrochemical
impedance spectroscopy is discussed in Chapter 4.
1.3 Miniaturization
Like many mature technologies, traditional immunoassay procedures have been
developed to the point where substantial increases in analytical performance are not
obtained by simply altering procedures. Instead, a tradeoff is observed, where an
increase in one quality is offset by a decrease in another. Miniaturization promises to
break free of this and increase the performance of the sensing technique with very limited
tradeoffs.
Miniaturization is effective because it preserves the essential steps of an analytical
procedure, but increases efficiency by reducing the scale of the elements involved,
namely time, materials, power, and consequently, cost. The big caveat is that
14 miniaturized systems can be difficult to fabricate, characterize, and interface with
existing macroscale systems because humans have been traditionally relegated to
manipulating materials on a scale limited by optical acuity and manual dexterity. Modern tools, particularly those developed by the semiconductor industry for fabricating
integrated circuits, have greatly decreased the scale in which sensor materials can be
manipulated. In contrast to this “top down” approach to miniaturization, small structures
can also be created by the self-assembly of molecules based on chemical properties and
molecular shape, similar to the way in which biological systems are formed.
While the possibilities are limitless, there are many practical limitations that have prevented miniaturized sensors from becoming more widespread. One serious limitation is fragility. Most micro- or nano- scale devices are extremely fragile, and with few exceptions, cannot be repaired if damaged. Even a microscopic dust particle can ruin electrodes, clog microchannels, or damage microscale optics. This fragility has limited the complexity of microsensors, as it is not cost effective to invest a great deal of time and effort into making something that is likely to be ruined soon after it is exposed to air.
A second limitation is cost. While the final manufactured miniaturized product may seem more cost effective than a macroscale one, microfabricated systems are very expensive to develop. Typical microfabrication requires a clean room facility, highly trained technicians, high purity materials such as gold, platinum, and crystalline silicon, and expensive instruments to do processes such as electron-beam writing, scanning electron microscopy, molecular-beam epitaxy, and atomic force microscopy. At the present time, only a limited number of sensing applications, such as monitoring glucose and detecting explosives, have a large enough market to support the multi-million dollar
15 investment required to commercially develop and produce microscale sensing devices. In
the future, more efficient and reliable fabrication techniques, cheaper materials, and a more experienced pool of scientists and engineers should increase the frequency of sensor miniaturization.
In addition to practical concerns, there are fundamental limitations to microscale
sensing based on scaling and sampling. The concept of “concentration” breaks down when dealing with very small volumes. Macroscale sensing relies on the statistical average of numerous molecular interactions, while in some cases micro- and nano- volume measurements do not measure enough discrete molecular recognition events to give useful data. For example, consider a one liter volume with 1,000,000 bacteria. A sensor that only samples a 10 nL volume, even if able to detect a single organism, cannot accurately measure the concentration of the entire liter with a single measurement as the sample volume only has a one in a hundred chance of containing a bacterium. Still, miniaturization has a long way to go before fundamental limitations such as this are applicable to the sensing of molecules (as opposed to organisms).
This dissertation describes the work my collaborators and I have done to improve immunosensing through miniaturization. In Chapter 2, the fabrication of prototype microfluidics, as well as the manipulation of fluids and particles in microchannels, are described. Specifically, experiments were done to transfer the laboratory steps of a microbead-based sandwich immunoassay to a microchip. Chapter 3 discusses my work with interdigitated array microelectrodes, and examines their utility as the detection element for a bead-based assay. Chapter 4 moves away from full immunoassay and focuses on label-free detection using electrochemical impedance spectroscopy with an
16 antibody probe. The small-scale aspect of this research is its use of carbon nanotube arrays and their unique nanometer scaled geometry.
In addition to the core research that is presented in this dissertation, I spent a great deal of time working on ancillary projects that are closely related. Information about some of these projects can be found in the publications listed in Appendix A.
17 References
[1] C. P. Price, and D. J. Newman, Principles and Practice of Immunoassay, ed., Macmillan Publishers, ltd., New York 1991. [2] R. Schall, Jr, and H. Tenoso, Alternatives to radioimmunoassay: labels and methods, Clin Chem 27 (1981) 1157-1164. [3] A. Ronald, and W. H. Stimson, The evolution of immunoassay technology, Parasitology 117 (1999) 13-27. [4] C. A. Wijayawardhana, H. B. Halsall, and W. R. Heineman, Micro volume rotating disk electrode (RDE) amperometric detection for a bead-based immunoassay, Analytica Chimica Acta 399 (1999) 3-11. [5] J.-W. Choi, K. W. Oh, J. H. Thomas, W. R. Heineman, H. B. Halsall, J. H. Nevin, A. J. Helmicki, H. T. Henderson, and C. H. Ahn, An integrated microfluidic biochemical detection system for protein analysis with magnetic bead-based sampling capabilities, Lab on a Chip 2 (2002) 27-30. [6] M. J. Green, Electrochemical Immunoassays, Philosophical Transactions of the Royal Society of London. Series B, Biological Sciences 316 (1987) 135-142. [7] H. Gao, T. Jiang, W. R. Heineman, H. B. Halsall, and J. L. Caruso, Capillary enzyme immunoassay with electrochemical detection for determining indole-3-acetic acid in tomato embryos, Fresenius' Journal of Analytical Chemistry 364 (1999) 170 - 174. [8] J. H. Lee, K. S. Hwang, J. Park, K. H. Yoon, D. S. Yoon, and T. S. Kim, Immunoassay of prostate-specific antigen (PSA) using resonant frequency shift of piezoelectric nanomechanical microcantilever, Biosensors and Bioelectronics 20 (2005) 2157-2162. [9] D. R. Shankaran, K. V. Gobi, T. Sakai, K. Matsumoto, K. Toko, and N. Miura, Surface plasmon resonance immunosensor for highly sensitive detection of 2,4,6- trinitrotoluene, Biosensors and Bioelectronics 20 (2005) 1750-1756. [10] S. Kurosawa, J.-W. Park, H. Aizawa, S.-I. Wakida, H. Tao, and K. Ishihara, Quartz crystal microbalance immunosensors for environmental monitoring, Biosensors and Bioelectronics 22 (2006) 473-481. [11] H.-C. Lin, and W.-C. Tsai, Piezoelectric crystal immunosensor for the detection of staphylococcal enterotoxin B, Biosensors and Bioelectronics 18 (2003) 1479 - 1483. [12] F. Darain, D.-S. Park, J.-S. Park, and Y.-B. Shim, Development of an immunosensor for the detection of vitellogenin using impedance spectroscopy, Biosensors and Bioelectronics 19 (2004) 1245-1252.
18
Chapter 2:
Microfluidic Immunoassay
19
2.1 Introduction to Microfluidics
2.1.1 Microfluidic Immunoassay
In the past two decades microfluidic technology has evolved from an intriguing concept to a vigorous growth industry that produces numerous publications and patents per year and yet can still be considered in its infancy. This process is being driven from multiple directions: technological advances in fabrication that push the limits of what types of devices can be created, and the demand for the accurate, efficient, fast, automated analytical systems that can be achieved through miniaturization. An enormous amount of interest in microfluidics is due to the recent explosion in biotechnology, with high throughput, small volume, combinatorial screening representing the new paradigm of proteomics and drug discovery. In particular, microfluidics promise to improve immunosensing systems by scaling down the physical dimensions of the sensing device, leading to the cost and analytical benefits that are discussed in this chapter.
Microfluidic immunosensor systems typically consist of microchannels for transporting fluids, with part or all of the necessary components of an immunoassay procedure integrated with each other on the chip. Relevant information on microfluidic technologies related to immunoassays can be found in several reviews [1-4]. Information
on the closely related field of immunoassay microarrays has also been reviewed [5, 6].
Microfluidic technology, in general, seeks to improve analytical performance by
reducing the consumption of reagents, decreasing the analysis time, increasing reliability
and sensitivity through automation, and integrating multiple processes in a single device.
20 These features are particularly suitable for immunoassay applications for a variety of
reasons:
1) Reagent and sample amounts. Many of the reagents integral to immunoassays
are expensive, often hundreds of dollars per milligram, and therefore miniaturization can reduce reagent costs drastically. In trace analysis applications, particularly those of a
medical, and high throughput combinatorial nature, for example, it is the analyte that is
the precious commodity, such as blood from a neonate, or spinal fluid
2) Decreased analysis time. One of the shortcomings of the heterogeneous
immunoassay procedure for some diagnostic applications is that it is a batch analysis
rather than a real-time sensor. By reducing diffusional distances and limiting analysis
time to seconds or minutes rather than hours, immunoassay sensor systems can
potentially be fast enough to monitor continuously without sacrificing sensitivity or
reliability, and thereby closely mimic a true sensor.
3) Automation. A major source of the loss of precision in many bench-top
immunoassay procedures is simple human error. Unlike larger scale robotic automated
equipment, microfluidic devices can be constructed to function without any moving parts
that wear and break, so they can be relatively inexpensive to produce and operate. The
movement of fluids can be precisely controlled by software linked to pumps, valves,
mixers etc. so that each sample is essentially run identically. These off-chip accessories
do contain moving parts and will wear out eventually, but are typically much easier to
replace than the robotic equipment used for larger scaled analyses. The end result is that
accurate measurements that typically require the skill of an experienced technician can be
obtained by the simple execution of a program.
21 4) Integration and portability. There is potentially more benefit to miniaturization than a
simple scaling down of the dimensions and volumes used [7]. The surface area to
volume ratio can be much higher when operating on the microscale, making material
transfer from bulk solution to microchannel surface more efficient. Therefore chemical
interactions between the surface and solution phase are much more pronounced because the diffusional distances are shorter.[8]. In addition, the necessary fluid volumes are so
small that very little energy is needed to manipulate them. This makes possible passive devices that require no external power input, but instead rely on diffusion and/or capillary action to control the assay [9], and portable devices that capitalize on a reduced power consumption [10].
Microfluidics can make existing biochemical assay applications more practical.
For example, a typical clinical procedure can involve the collection of a sample at one location and then a delay while the sample is sent to the lab and processed. Small, self- contained, integrated microfluidic devices promise to provide rapid, point-of-care diagnostics that greatly enhance the speed at which results can be obtained[11]. Another important practical capability provided by microfluidics is portability. The fluid volumes that need to be moved are so small that little power is consumed, making passive or battery powered analyses feasible. When full laboratory access is not practical, such as when on a boat, engaged on a battlefield, or even in space, microfluidics may represent a good option for sensitive detection.
2.1.2 Microfluidic Materials
22 The materials used to construct microfluidic immunoassay devices vary
depending on the application, and the vast majority are constructed of glass, silicon, or
polymers. Each of these materials has its own advantages and limitations. Much of the
early microfluidics work was done using silicon as a substrate, as the technology for
patterning, etching, and bonding silicon wafers had already been developed in the
electronics industry, and for microchips and other microelectromechanical systems
(MEMS). Silicon is structurally very strong, and can be etched isotropically or
anisotropically to create precise features on a nanometer scale. In addition, the surface
chemistry of silicon, or rather the microscopic layer of silicon dioxide that forms upon
contact with air, has been studied extensively and many techniques exist to attach
molecules and coatings covalently to the reactive silanol groups. Despite these
advantages, silicon has been replaced by glass and polymers as the preferred material for
most immunoassay applications. This is because silicon is not optically transparent at
the wavelengths typically used for optical detection, and the surface properties of silicon make it unsuitable for applications that rely on electrokinetic flow to transport reagents.
Proteins and other biomolecules also tend to adhere to silicon surface groups, and while
the adsorption can be reduced by a surface treatment such as silanization, the sensitivity
of a sensing application may still be reduced. Many of the problems encountered with
silicon were overcome with glass substrates. Although more fragile than silicon, glass
can be made with outstanding optical properties throughout the visible spectrum, and can be used to transport fluids electrokinetically. In addition, glass is highly resistant to solvents. For many immunoassay applications, particularly those involving optical detection, glass is the standard material. Glass and silicon devices do possess a few
23 potential shortcomings, however. For example, the wet etching process typically used in
fabrication, while automatable and highly reproducible, does not produce high aspect
ratio features, and the microchannels generally have a curved bottom that can complicate
optical detection.
Although, polymers were viewed initially as an inferior yet far cheaper alternative
to glass, they can be mass produced easily, as the expensive patterning procedures are
only required for constructing the mold or tool Recently there has been increased
interest in polymer and plastic microfluidics mostly because the sheer variety of
materials, each with specific optical, surface, and mechanical properties, and solvent
resistance offers the possibility of tailoring a material to a specific application [2]. In the
future, the bulk of immunoassay applications will probably be done on a polymer
substrate, as the versatility, reduced cost, and simpler fabrication methods outweigh the
benefits of glass and silicon, and the solvent resistance properties of glass would not be needed[12].
One particular polymer that has recently been used extensively is poly(dimethylsiloxane), or PDMS. PDMS is a transparent, elastomeric polymer that can be rapidly fabricated with features having dimensions as small as 10 nm [3]. The
elastomeric nature of PDMS makes it seal well, and often the adhesion due to conformal
surface contact with a smooth, flat surface can be enough to seal microchannels for low
pressure applications. Alternatively, PDMS can be bonded irreversibly to itself, glass, or
silicon by treating both bonding surfaces with an oxygen plasma[13]. Another significant
advantage of PDMS over glass, silicon, and many other polymers is that the PDMS
fabrication process can be done under normal laboratory conditions, as opposed to the
24 confines of a clean room that can be expensive to maintain or difficult to access.
Furthermore, the recent surge in interest for PDMS as a material for microfluidics has
resulted in an extensive body of literature about modifying and functionalizing PDMS
surfaces [14, 15]. The principal limitations of PDMS for immunoassay applications are due
to its hydrophobic surface properties. Thus, proteins and other molecules tend to adsorb
to an untreated PDMS surface, and the surface is poorly wetted and prone to bubble formation in aqueous systems. These limitations can be reduced or eliminated through chemical or plasma treatment of the surface[13, 16].
2.1.3 Microfluidics Using Microbeads
Immunoassay, as discussed in Chapter 1, is a sensitive analytical technique that uses the affinity and selectivity of an antibody to sense for a specific antigen. The most sensitive type, the heterogeneous sandwich immunoassay uses a solid support loaded with antibody to capture the intended analyte. In order to reduce the signal from the background of interfering substances, a rinsing step is used to wash away the interferences while retaining the captured antigen. The solid support has typically been
the walls of the vessel that conbtains the immunoassay solutions (wells, cuvettes,
capillaries, microchannels), but this can be inefficient because the analyte can have a long
distance to diffuse from the middle of the bulk solution to the surface. In addition,
devices using their surfaces as the solid support for the immunoassay must be thoroughly
cleaned and reloaded with antibody between measurements, or they can be single use.
This is not a problem if the solid support is a a microcentrifuge tube or microtiter plate
well, but microfluidics, particularly when in the prototyping stage, can be too expensive
25 and time consuming to fabricate to be used but a single time. Immunoassays have been
developed that use microbeads as the solid support chemically or physically linked to the
antibody.[17] As discussed in Chapter 1, microbeads provide more surface area for a given volume than a microchannel, and when dispersed, reduce incubation times by shortening the diffusional distances. Also, beads can lower analysis time if they are pre- loaded with antibody, and replaced in the channel with a fresh aliquot of beads between
assays. This strategy also provides a highly reproducible control of the amount of
antibody being delivered to the channel. Beads can be made with an iron core so that they
are paramagnetic and can be manipulated with a magnetic field. This allows selective
immobilization and dispersion at different points in the assay procedure. Commercially
available beads usually contain reactive surface groups that can be attached to antibodies,
labels, or other molecules. Beads are also available with a streptavidin coating, allowing a biotinylated antibody to be immobilized easily.
2.1.4 Surface Modification
In the microscale environment, the high surface area/volume ratio of surface to
solution magnifies the effects of non-specific binding. This effect is especially important
in immunoassays where key reagents such as antibodies and enzyme labels can adsorb to
hydrophobic surfaces, seriously degrading the assay performance. To address this
problem, different strategies have been used to block or modify the surface of the
microchannel, the most common of which has been to use a blocking agent such as
bovine serum albumin (BSA) to pre-coat the surface. While sufficient for many
applications, the surface modification is not permanent, and the resulting surface is
26 somewhat heterogeneous. Recently there has been work done to simultaneously reduce
non-specific binding and add functionality to microchannel surfaces. Lahann and co-
workers presented a method to create a well-defined reactive surface coating for PDMS
using chemical vapor deposition (CVD) [18]. This method provides control over the
surface chemistry of the microchannel, while simultaneously presenting reactive amine groups to covalently attach antibodies. Another work described an immunoassay
procedure where the microchannel surfaces were covered by lipid bilayers[19].
Recognizing that antibodies directly adsorbed on a silica surface may be partially or completely denatured due to the conformational change associated with the adsorption process[20], strategies were developed to control the nature and orientation of antibody
attachment. The techniques included: (a) direct covalent attachment via glutaraldehyde-
activated APTES, (b) covalent attachment to adsorbed BPEI (branched
poly(ethyleneimine)), (c) covalent attachment to covalently bonded BPEI, (d) covalent
attachment to adsorbed LPEI (linear poly(ethyleneimine)), and (e) adsorption to a layer
of adsorbed LPEI. Their results indicated that the best stability and sensitivity were
achieved when antibodies were covalently attached to a polymer.
The work discussed in this dissertation specifically addresses the use of
paramagnetic microbeads as a solid support for immunoassay in a PDMS microfluidic
device, although much of the data acquired can also be applied to more general cases of
protein adsorption on other surfaces. In fact, the PDMS prototypes discussed in this
chapter are merely model systems for future microfluidic sensing platforms. Specifically
addressed are the procedures for fabricating PDMS devices, and analyses that show the
feasibility of the different steps of immunoassay.
27
2.2 Fabrication of PDMS Microfluidic Device
Devices were fabricated in collaboration with Erik Peterson, a graduate student of
Ian Papautsky from the Department of Electrical and Computer Engineering, University of Cincinnati. The procedures outlined here describe the techniques with which we had the most success.
2.2.1 Creation of the SU-8 Master
The patterns for the PDMS microfluidic devices were fabricated by first depositing
SU-8 negative photoresist to 100 microns thick, (2075, MicroChem Corp.) on 76mm silicon wafers. SU-8 was patterned using UV lithography and a laser-fabricated chromium mask to produce the negative mold for casting the PDMS. The pattern for the mask was generated using AUTOCAD. The lithographic procedure used to create the devices was as follows:
1. 3” (76mm) Si wafers are cleaned:
a. acetone rinse (30 s)
b. methanol rinse (30 s)
c. DI rinse (30 s)
d. piranha clean – 7 parts concentrated (18.4 M)H2SO4: 3 H2O2 (70ml :
30ml) in Petri dish for 10-15 min.
e. DI rinse (3 min)
f. BOE Dip 15-30 s (BOE is Buffered oxide etchant- 3 parts DI H2O : 3
parts NH4F : 1 part concentrated HF;
28 g. DI Rinse (30 s)
h. N2 blow dry
2. Dehydrate on 150°C hotplate for 10 – 15 min then cool to room temp.
3. Spin coat resist (for approx 100 micron thickness)
a. spread: Ramp 100 rpm/sec to 500 rpm and hold for 10 sec
b. spin: Ramp 300 rpm/sec to 1700 rpm and hold for 30 sec
c. place wafer on level surface for 10- 20 min
4. Soft bake (Pre Bake) in programmable convection oven on level surface
a. –Oven Program #1--
b. ramp from room temp at 1°C/min to 65°C and hold for 10 min
c. ramp from 65°C to 95°C at 1°C/min and hold for 1 hour 45 min.
d. turn off heat and cool naturally to room temp
(Soft bake takes approx 6-7 hrs)
5. Expose using the I-line (365nm) high pass filter (Deep UV will mess up the
exposure)
a. exposure energy: 550 mJ (typically 6.5mW/cm2 for 85sec)
b. may use layer of glycerin for reduced diffraction effects (Glycerin must
then be rinsed with DI and then wafer must be blow dried before going
back into oven)
6. Hard Bake (Post Exposure Bake) --same oven
a. –Oven Program #2--
b. ramp from room temp at 1°C/min to 65°C and hold for 10 min
29 c. ramp from 65°C to 95°C at 1°C/min (might be 0.5°C/min?) and hold for
20 min.
d. turn off heat and cool naturally to room temp (Hard Bake takes approx 5
hrs)
7. Develop- use SU-8 developer
a. place wafer on dipper in 500ml or 1L beaker full to 1-2cm above top of
wafer, stir at 200-300rpm for 25-50min until complete (periodic inspection
with microscope)
b. rinse with isopropyl alcohol
c. N2 Blow dry
8. Plasma Clean
a. O2 Plasma for 30 s – 5 min depending on amount of residue left on wafer
after development.
b. O2 at 20 sccm, 130-150 mTorr, 300W, 30kHz
c. allow wafers to stand for 1-2 days before casting PDMS.
The mold used is illustrated in Figure 2.1. All lithography procedures were done in a clean room facility. SU-8 masters were inspected with light microscopy and a profilometer.
30 400µm wide 200µm wide 4 cm long 4 cm long
Figure 1. Diagram of SU-8 master on which PDMS was cast.
2.2.2 PDMS Casting
PDMS was then prepared by mixing the base and curing agent at a 10:1 ratio
(m/m) vigorously for at least 2-3 minutes. Insufficient mixing was shown to create a
material that would not cure completely, and thus destroy microscale patterning when
removed from the mold. After this the PDMS was degassed in a vacuum, poured into
Petri dishes with the SU-8 master, degassed again, and finally cured on a level hotplate at
80°C for 1 h. The degassing step is necessary to remove the bubbles that form when the
precursors are vigorously mixed. If PDMS of optical quality is not required, the second
degassing step can be omitted. To facilitate mold release, the SU-8 masters were dip
coated with Sigmacote and air-dried prior to pouring the PDMS. This procedure was
31 effective when casting PDMS between 500 µm and several millimeters in thickness.
Thinner layers tend to tear when removed from the SU-8 master, and thicker castings
require more time to fully cure.
2.2.3 Bonding
The cast PDMS, when released from the master, contains three of the four walls
of the microfluidic channel. In order to seal the trench and create a true microchannel, it
must be bonded to another substrate. We experimented with both glass and polymer substrates to make the fourth wall. If we needed to do electrochemistry in the channel, it is much easier to deposit microelectrodes on a glass substrate than polymer, so procedures were developed to bond to glass. Procedures to bond to PDMS were also explored, because, in addition to being less prone to breakage than glass, it created a more uniform microchannel surface to use for experiments that characterize surface properties.
Several strategies were used to seal the microchannels. The first and simplest strategy was to simply use mechanical pressure to hold the two pieces together. Several fixtures were constructed, out of stainless steel and Plexiglas, that allowed screws to be tightened to clamp the two halves together. In addition to frequently breaking the glass substrates, these methods were only moderately successful. Invariably, small leaks would develop, necessitating a tightening of the screws. Eventually, the pressure would become too great and the PDMS would buckle slightly, causing a serious leak.
A second method, that was used to bond PDMS to PDMS to seal the microchannels, was to remove the PDMS from the mold before it was finished curing, but after it had solidified enough to preserve the pattern. Then, the pieces were placed
32 together and cured, creating a bond between the two. This method was generally very
effective, although it produced a lower quality of microchannels. The edges of the
partially cured PDMS were less sharp and more rounded than usual, creating slightly
misshapen microchannels.
[21, 22] The third method used to bond the PDMS was brief exposure to O2 plasma .
In order to seal the microchannels, the molded PDMS substrates were cleaned and bonded to smooth PDMS using a reactive ion etch (RIE) O2 plasma (20 s, 70W, 13.56
MHz, 20 sccm)[23]. The two halves of PDMS were slightly wetted with ethanol to prevent
an instant bond so that they could be aligned. When the ethanol dried an irreversible bond was formed. This procedure, while effective, required the use of a clean room and an expensive piece of equipment. An alternative bonding procedure was developed using the Harrick PDC-32G plasma cleaner. In place of carefully controlled oxygen flow, oxygen from the air was used in the plasma. An evaluation of the elements of the bonding procedure can be seen in Table 1. In summary, wash steps and oven drying were essential to achieving a bond, as was a short plasma duration. Based on these parameters, a double washing step (HCl and ethanol) followed by an oven dry and 15 s plasma exposure was used to bond devices.
33
Table 2.1 Summary of PDMS bonding
Sample HCl Wash Ethanol 30 min Plasma Bond
Sumber Wash Oven Dry Duration(s)
1 yes yes yes 15 yes
2 yes yes yes 60 no
3 yes no yes 15 yes
4 yes no yes 60 no
5 yes yes yes 15 yes
6 yes yes no 60 no
7 yes yes yes 15 yes
8 no yes yes 60 no
9 no no yes 15 no
10 no no yes 60 no
11 yes yes yes 10 no
12 yes yes yes 20 yes
13 yes yes yes 30 yes
14 yes yes yes 40 yes
15 yes yes yes 50 no
16 yes yes yes 60 no
34
2.2.4 Connection to Syringe Pumps
Chromatography tubing and fittings were used to connect the device to a syringe pump for testing. A Plexiglas fixture was constructed to immobilize the tubing at its point of connection to the device. This was necessary because the fragile bond between the tubing and the fluidic could be easily broken with the slightest torque to the tubing.
In order to add additional strength to the tubing/fluidic junction, additional cylinders of
PDMS were cast in the Plexiglas mold, greatly increasing the device’s durability.
Devices were fabricated with channels 4 cm long and 200 µm, 400 µm, and 1 mm in width. The fabrication procedure can be seen in Figure 2.2 Figure 2.3 shows one of the
PDMS microfluidic devices used in the experiments.
Figure 2. Steps of PDMS device fabrication.
35
Figure 3. Photographs of PDMS microfluidic device.
36
2.2.5 Proposed On-Chip Immunoassay
A diagram of the proposed on-chip immunoassay can be seen in Figure 2.4. In order for a bead-based immunosensing procedure to be effectively carried out on a microchip, some important questions must be answered. The ability to introduce and manipulate microbeads must be demonstrated, and the microchip must be shown to be compatible with currently used detection strategies. Our research group has focused on both electrochemical and fluorescence detection methodologies for the bead-based assay, so microchannel measurents evaluating both were compared. Bead manipulation is a much more difficult objective to master. Preliminary testing with fluid samples went very smoothly, but when paramagnetic microbeads became involved problems began to arise.
Beads tended to settle in the space below the tubing connection, the walls and corners of the microchannel, and especially at the air/solution interface if a bubble was introduced.
No matter what flow rate, tubing or channel diameter, bead adsorption was consistently observed. To address this, a bead adsorption study was devised to identify the reason why the beads stick, and then to reduce or eliminate the adsorption through rinsing or surface treatment.
37 Figure 4. Proposed immunoassay setup for PDMS microfluidic.
2.3 Compatibility of Standard Detection Methods with Microfluidic Device
2.3.1 Fluorescence detection
In order to demonstrate the feasibility of fluorescence detection in microchannels, a calibration curve was constructed for fluorescein injected into the PDMS microfluidic channels. This was to confirm that the microscope intensity readings would directly
correlate to the concentration of fluorophore in the channel. The curve for an assay
relevant range of fluorescein concentrations measured by Irena Nikcevic, the collaborator
working on the aspects of the project involving fluorescence detection, showed an
acceptable correlation coefficient, indicating that this detection method could be used for
measuring microbeads within channels (Figure 2.5).
38
2.60E+09
2.10E+09
1.60E+09 y = 2.27·107x + 1.81·108 R2 = 0.99996 1.10E+09
Fluorescence Intensity (a.u.) Intensity Fluorescence 6.00E+08
1.00E+08 0 20406080100
Fluorescein Concentration (nM)
Figure 5. Calibration curve for fluorescein in PDMS microchannel.
2.3.2 Electrochemical detection
The use of vapor deposited microelectrodes in PDMS microchannels has been
reported in the literature[24, 25], but it would add considerable cost to an otherwise inexpensive fabrication procedure. Instead, laser-cut Kapton fluidics that were readily available were used to make on-chip electrochemical measurements. A calibration plot for an electrochemically active enzyme product, para-aminophenol (PAP) can be seen in
Figure 2.6. A more thorough discussion of PAP and its role as an enzyme product can be found in chapter 3. The microelectrode showed sensitivity over a large dynamic range, from nanomolar concentrations to micromolar. In addition, the effect of flow rate was investigated, and can be seen in Figure 2.7. As expected, the higher flow rates produced
39 higher currents due to the increased efficiency of redox molecule transport to the electrode surface. Although expected, this analysis was necessary to show that sufficient sensitivity could be achieved even at very low flow rates that could be limited by the durability of the fluidic.
Log Log Calibration Plot
-4.000
-4.500 y = 1.1154x - 0.3627 -5.000 R2 = 0.9996
-5.500
-6.000
-6.500
-7.000
log current (A) log current
-7.500
-8.000
-8.500
-9.000 -8.000 -7.500 -7.000 -6.500 -6.000 -5.500 -5.000 -4.500 -4.000 -3.500 -3.000 Log PAP concentration (M)
Figure 6. Log-Log Calibration Plot for PAP in PBS pH 7, 50 µL/min flow rate in Kapton fluidic.
40
1800
1600
1400
1200
1000
800 current (nA) current
600
400
200
0 0 50 100 150 200 250 300 350 400 450 flow rate (µL/min)
Figure 7. Effect of flow rate on the measured current of 1mM ferricyanide in 0.1 M KCl
2.4 Bead Adsorption Study
When microbeads were injected into the PDMS prototype fluidics, a significant amount of bead adsorption was observed on the microchannel walls, particularly in corners, turns, and other places where lower flow velocity allowed them to settle.
Because bead manipulation is such an integral part of the immunoassay procedure, a bead adsorption study was done to identify the nature of the observed bead adsorption, and to identify strategies to reduce or eliminate it. This work was done in collaboration with
Erik Peterson and Irena Nikcevic.
41
2.4.1 Reagents
Polystyrene coated magnetic beads with streptavidin covalently attached to the
bead surface, M-280 Dynabeads Steptavidin, were obtained from Dynal Biotech. The
beads were obtained as a monodisperse suspension (2.8 µm diameter, 6.7·108 beads/mL).
The fluorescent dye, 5(6)-FAM, SE (5-(and-6)-carboxyfluorescein, succinimidyl ester), was obtained from Molecular Probes. Tris buffer (pH 9) and phosphate buffer, PBS (pH
7.2) were used as solvent. Bovine serum albumin (BSA) and Tween 20 were obtained from Fisher Scientific. Sigmacote was obtained from Sigma-Aldrich. Triton X 100 was obtained from Amersham Biosciences. Teflon AF 1600 was from DuPont.
Polydimethylsiloxane (PDMS) was purchased as a two component kit (Sylgard 184) from
Dow Corning (Midland, MI).
2.4.2 Equipment
Beads were observed using a Nikon Eclipse TE-2000 inverted epifluorescence microscope. Images were obtained with a 16 bit CCD camera (Roper Scientific-
Photometrics (Tucson, AZ) at 40x, 100x and 400x magnifications. A xenon arc lamp
(Sutter Instrument Company LB-ls/17) was used as the fluorescence excitation source.
The appropriate excitation and emission wavelengths were selected with a FITC filter cube (Chroma Technology Corp.). This signal was spatially filtered and then detected with a CCD camera. Images were acquired using MetaMorph software (ver. 6) from
Universal Imaging Corp. Environmental scanning electronic microscope (ESEM) images were acquired using a XL 30 (FEI Company).
42 Contact angle measurements were made using the sessile drop technique with a
Tantec CAM-Micro contact angle meter. In this technique, a small drop of liquid was dispensed onto the surface of a sample using a microliter syringe, and the contact angle
was measured through the liquid phase using the half-angle method. Measurements were
made at five different locations on each sample.
2.4.3 Bead Labeling Procedure
FAM, SE (succinimidyl ester) is a fluorescein derivative with a functional group
that provides an efficient and convenient way to selectively attach to primary amines (R-
NH2), located on the protein. Amide bonds formed in this reaction are as stable as
peptide bonds. Bead labeling was done by Irena Nikcevic and the procedure is detailed in
our publication[26].
2.4.4 PDMS Sample Preparation
The PDMS samples were prepared by mixing the base and curing agent at a 10:1
ratio (m/m), degassed in a vacuum to remove air bubbles, then poured into Petri dishes,
degassed again, and finally cured on a level hotplate at 80°C for 1 h. Samples were cut
into 1 x 2 cm rectangles, and had a uniform thickness of ~1 mm. Samples were cleaned
ultrasonically and rinsed with 70% ethanol and deionized water, and then air-dried.
2.4.5 Surface Treatments
Several surface treatments were used that have been shown to resist adsorption in
other systems. BSA is a common blocking agent for biological assays[27], Sigmacote is a
43 silanizing agent used to resist cell adhesion in cell culture studies[28], Teflon is an inert
[29] fluoropolymer well know for its highly slippery nature , and CHF3 plasma treatment
has been shown to deposit a Teflon-like fluoropolymer coating[30].
The adsorption experiments were first done on flat PDMS samples, later followed
by experiments inside the microchannels. PDMS controls received no surface treatment
after cleaning. BSA treated samples were incubated with 1% BSA for 48 h and then
dried for 24 hrs. Sigmacote treated samples were dip-coated with Sigmacote for 20 s and
air-dried. Samples treated with Teflon were dip-coated with Teflon AF for 20 s and then
cured at 65° C for 10 min, then the temperature ramped to 110° C for 20 min, then
ramped down to 35°C for 8 h before allowing them to cool to room temperature.
Samples treated with CHF3 plasma were processed at 300 W for 240 s with a CHF3 gas flow rate of 20 sccm.
2.4.6 Data Analysis Procedure
After labeling beads, images were recorded with the MetaMorph imaging software. The averages of greyscale values across all image pixels in a given region was used to measure the total fluorescence from a given area. The fluorescence intensity measured from each successive concentration of beads was used to establish a calibration curve.
2.4.7 Results and Discussion
44 Of all the treatments, Teflon AF was by far the most effective, as seen in Figure 2.8. The other surface treatments failed to reduce bead adsorption by more than a few percent, while Teflon AF treatment eliminated all but 0.3% of the bead adsorption.
Comparison of Surface Treatments
100
90
80
70
60
50
40
30 % of beads remaining after water rinse
20
10
0 Bare PDMS Sigmacote 1% BSA CHF3 Plasma Teflon AF Surface Treatment
Figure 8. Comparison of the ability of different surface treatments to resist the adsorption of microbeads.
A useful tool for surface characterization is the water contact angle measurement.
The contact angle is the angle at which an gas/liquid interface meets a solid surface, and is defined by the properties of the substance in each phase. An illustration of this can be seen in Figure 2.9. For a water contact angle measurement, the liquid and gas are water and air respectively, so the contact angle depends only on the properties of the solid surface. A rounded drop with a high contact angle indicates a hydrophobic surface, while
45 a flat drop with a low contact angle signifies a hydrophilic surface. Contact angle measurements were made in this study to determine whether the hydrophobicity/hydrophilicity of the surface was a critical factor that caused beads to adsorb. The contact angles were measured using the sessile drop technique on the treated and untreated PDMS samples and are given in Table 2.
Hydrophobic Hydrophilic Surface Surface
Figure 9. The water contact angle is a measure of surface hydrophobicity/hydrophilicity.
Table 2.2 Contact angle measurements on treated PDMS surface
Surface Treatment Contact angle
Bare PDMS 109° (±1)
BSA 105° (±2)
Sigmacote 53° (±7)
Teflon AF 114° (±1)
CHF3 Plasma treatment 111° (±1)
As shown, the water contact angles obtained for the different surface treatments are similar to untreated PDMS, with the exception of the 1% BSA treatment. While most treatments produced hydrophobic surfaces, with highest values obtained for CHF3
46 plasma, treatment with 1% BSA rendered the surface hydrophilic, and seemed to be the
least reproducible coating. Contact angles measurements suggest that hydrophobicity is
likely not the determining factor for bead adsorption under the conditions tested, as no
correlation between the hydrophobicity of the surface and increased or decreased bead adsorption was observed.
Roughness and impurities in the PDMS can change the microscopic surface area and surface properties of the polymer so that it is more likely to interact with the microbead. It was thought that surfactant treatment would be able to reduce bead adsorption by masking the surface groups on the streptavidin coated microbeads.
Surfactant molecules cover up the “hydrophobic regions of proteins, making them more solvated and less prone to adsorption or agglomeration. However, results indicated that only slightly fewer beads were prevented from sticking to the PDMS surface, suggesting that the observed interactions are not primarily hydrophobic. Altering the pH resulted in sticking that was slightly less than that of pure water, although the result was not dramatic enough to be considered the driving force causing bead adsorption.
2.5 Conclusions
This work has show that we have the ability to rapidly and inexpensively prototype microfluidic devices using lithography and cast PDMS. The primary immunoassay functions that must be integrated with the microchip to produce an
effective microbead-based device are controlling the movement of beads and detecting of
the electrochemical or fluorescent signal. The optically transparent nature of PDMS
makes fluorescent detection feasible, and electrochemical detection in a fluidic channel
47 made of another material was also explored. The manipulation of microbeads remains a
significant challenge, however, although the observation that the teflon coating nearly eliminated sticking offers promise that this problem will be solved in the near future.
The fact that very few beads adsorbed to the Teflon AF surface should not be surprising. Fluorocarbon polymers are well known for their resistance to adhesion and fouling. The failure of the other surface treatments to halt the adsorption of beads, as well as the high quantity of adsorption on bare PDMS, suggest the existence cooperative binding interactions between the bead and substrate surfaces. Both the beads and the
PDMS surface contain potentially reactive surface groups. The chemistry of the bead surface is primarily determined by the attached streptavidin. Streptavidin, like all proteins, has reactive and charged surface groups exposed to solution. In addition, there may be patches of polystyrene exposed where the streptavidin does not completely cover the surface of the bead. The PDMS surface can possibly contain unreacted functional groups that interact with either the protein or the exposed polystyrene on the microbead.
These surface interactions are possibly the reason why such a high quantity of beads stuck to the PDMS. If the exact nature of the surface interactions were determined, then specific chemistries could be used to counteract the adsorption using blocking agents or a specific surface treatment. Until the chemistry is developed to make a PDMS surface compatible with streptavidin coated microbeads, the best solution for preventing adsorption is the placement of a chemically inert layer of a polymer such as Teflon AF between the surface and solution.
Problems related to the nonspecific adsorption of beads must be solved before a practical bead-based PDMS microfluidic device can be used for immunoassay
48 measurements. However, results of these experiments give indication that this problem may be solved using surface modification strategies.
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53
Chapter 3:
Immunoassay using IDA Microelectrodes
54
3.1 Introduction
As discussed in Chapter 1, various detection methods associated with different
modes of signal transduction have been developed in immunoassay techniques to
recognize an antibody/antigen binding event. This chapter focuses on the use of
interdigitated array microelectrodes as the detection element for paramagnetic bead-based
immunoassay.
3.1.1 Electrochemical Immunoassay Background
Possibly the most common detection technique not relying on optical
measurements is electrochemical detection, which has strengths and weaknesses due to
the way the signal is measured. Although it is sensitive, easy to miniaturize, requires
relatively simple instrumentation, and is capable of making measurements in turbid
samples that prohibit optical detection, electrochemical detection is limited by the fact
that physical contact between the analyte and electrodes is necessary. This contact is the
source of the sensitivity of electrochemical detection, but it also generates the possibility
of electrode contamination and fouling. In addition, analyte must diffuse to the surface of the electrode for current to be generated and provide a detectable signal, and that current is very sensitive to the rate that the redox-active molecules are able to diffuse to the surface. Mass transfer related to convection is much faster than Fickian diffusion under the parameters typical of most electrochemical measurements, so quantitative techniques can be sensitive to changes in stirring speed or flow rate.
55 3.1.2 Miniaturized Electrochemical Immunoassay
Electrochemical instrumentation can be easily miniaturized, providing a system
requiring low operating power. This makes electrochemistry suitable for on-board
detection, particularly when compactness and portability are important. Nonetheless, a
direct consequence of miniaturization is that only tiny currents are generated, and
improved electronics/shielding are often required for accurate measurements. A stable
reference electrode is often necessary. The typical Ag/AgCl reference electrode that is
used for larger experiments consists of a hollow plastic or glass tube with an enclosed
AgCl-plated Ag wire and a semi-permeable membrane that permits electrical contact
between the interior and exterior solutions, but prevents the fill solution from mixing with
the sample. The tube is filled with a chloride-containing electrolyte solution. The
solution is typically saturated with silver chloride to prevent silver chloride leaching from
the wire. As long as the chloride concentration remains constant, the reference potential
remains steady. In miniaturized systems, it is sometimes not practical, or even possible,
to scale down a reference electrode of this type. Instead, a silver wire coated with silver
chloride is used as the reference. There are some limitations to this practice, however. In
order for the reference potential to remain stable, the chloride concentration in the sample
must remain constant. This can be achieved by using buffered solutions of fixed chloride
concentration. In addition, silver chloride is very slightly soluble, so extended storage in
an aqueous solution will eventually leach away the silver chloride.
Another significant problem faced when using microelectrodes is electrode fouling. Unlike larger electrodes, microelectrodes cannot be mechanically polished
because they are so thin, so their lifespan can be very limited when working with a
56 species that tends to adsorb to the surface. Generally, fouling primarily occurs when
working with high concentrations, so a conscious effort to limit the amount of time
between experiments that the electrodes are exposed to fouling solutions can increase the
usable time of the electrode.
3.1.3 Analytical Advantages of Microelectrodes
Microelectrodes (i.e. electrodes with one dimension ≤50 µm) offer analytical
advantages over conventional, larger electrodes. In addition to being able to sample
smaller sample volumes, the drastic reduction of the electrode diameter to the low
micrometer range results in a hemispherical diffusion profile in close proximity to the
electrode surface.[1] This is in contrast to the planar diffusion profile observed on macroscaled electrodes, and results in an enhanced collection efficiency of an electroactive species at the surface during a voltammetric experiment. The practical result is that the signal-to-noise ratio increases, because the signal generated is proportional to the current generated, while the noise is proportional to the area of the electrode. The improved signal/noise ratio generally results in a lower detection limit. Therefore, voltammetry is unique in that its performance can actuall improve as the sensing electrodes become smaller. As a result, small volume analyses down to a micron drop diameter size have been demonstrated at microelectrodes with bead-based immunoassays.[2]
3.1.4 IDA Electrodes
57 A relatively recent addition to the arsenal of amperometric detection is the interdigitated array electrode (IDA). An IDA consists of a pair of interdigitated fingers with a narrow gap between them. During detection, each electrode is held at a different potential to promote redox cycling of the analyte. Illustrated in Figure 3.1 is the redox cycling of p-aminophenol, a product of an enzyme label commonly used in amperometric assays.
58
Figure 3.1. Redox cycling of p-aminophenol (PAP) and p-quinone imine (PQI). The
PAP is produced by the enzyme β-Galactosidase on the substrate p-aminophenyl galactopyranoside (PAPG)
59 One electrode is held at an oxidizing potential and the other at a reducing potential for
each half of the redox couple. If the analyte starts out in its reduced form, it interacts with
the surface of the oxidizing electrode (anode) in an electrochemical reaction, producing
an electric signal and an oxidized product. Instead of stopping there, as is the case with
an ordinary electrode, a cycle is initiated. The oxidized species diffuses to the other
electrode, where it is converted into its reduced form, generating more signal and
allowing the cycle to continue. This has been extensively reported in the literature and in
review articles.[3] High sensitivity is typically required for many biological assays such as
immunoassay, and small detection volumes are preferred due to the high cost or scarcity
of reagents and rapid analysis times associated with smaller dimensions and shorter
diffusional distances. For these reasons, IDAs have been used as the detection element
for enzyme assays and immunoassays.[4] Multiple strategies have been devised to
improve the sensitivity and utility of electrochemical assays using IDAs. Some assays have used the electrode surface as the solid support of the immunoassay[5], while others
incorporate particles such as microbeads for their increased surface area and easy
dispersal throughout a sample[6]. Photolithographic microfabrication techniques allow
IDA microelectrodes to be combined with microfluidic channels, resulting in devices that
can do an entire immunoassay procedure on-chip.[7] In another assay strategy, a trench
IDA electrode was fabricated to optimize detection efficiency.[8] The redox cycling rate
and thus the amplification of the electrode[9] are determined by the properties and
dimensions of the IDA, and include gap width, electrode width and height, number of
fingers and the electrode material. These variables have been investigated using model
analytes[10, 11]. An added benefit of the IDA electrode is that a high level of sensitivity can
60 be achieved on an unstirred sample, which is important for some assay procedures
because it gives more flexibility in sensor design by removing the necessity of stirring or
maintaining a controlled flow rate.
The observation was made that the cycling efficiency of IDA electrodes is proportional to the diffusion length (We/4 + Gap), where We is the electrode width and the gap is the distance between fingers. When the number of redox cycles is plotted against the dimensions of the IDA (using We/4 + Gap) a dramatic effect is seen as the diffusion length shrinks to the low microns [9]. A redox cycle number of approximately
5 is observed with a 7.5 µm IDA, while the 0.75 µm IDA had a redox cycle number
greater than 40. This amplification was approximated empirically by Aoki et al[12] with
the following equation that predicts the current of an IDA based on its geometrry:
⎡ ⎤ ⎢ ⎥ ⎢ ⎧ ⎛ w ⎞⎫ ⎥ ⎪ ⎜ f ⎟⎪ 0.19 I lim = nFDC * mb⎢0.637ln⎨2.55 1+ ⎬ − ⎥ ⎜ w ⎟ 2 ⎢ ⎩⎪ ⎝ g ⎠⎭⎪ ⎛ w f ⎞ ⎥ ⎢ ⎜1+ ⎟ ⎥ ⎜ w ⎟ ⎣⎢ ⎝ g ⎠ ⎦⎥
Where Ilim is the steady state current, nF is the number of electrons transferred per mole
of analyte, D is the diffusion coefficient, C* is the concentration of the bulk analyte, m is
the number of paired fingers in the IDA, b is the length of each individual finger, wf is the width of the finger, and wg is the gap between fingers. In practice, this equation indicates
that current increases exponentially as electrode dimensions are reduced.
Recently, IDAs with sub-micron electrode gaps have been reported that are
capable of a higher level of amplification due to the even more rapid redox cycling.[13-15]
61 The increased amplification associated with the shorter diffusion path of the analyte, when combined with very small sample volumes, results in very high sensitivity and low detection limits. These analytical characteristics are ideal for assays that quantitate pathogenic biological organisms. Such assays typically require very low detection limits because the organisms can be harmful at extremely low doses. Many strategies are used to increase the signal that can be generated from a captured analyte, such as using an enzyme label An enzyme label that rapidly converts many substrate molecules into a detectable product increases the number of detectable molecules dramatically. Similarly, an electrode geometry that cycles redox-active molecules will amplify the signal that is produced from each enzyme product. The combination of two different efficient amplification strategies, when combined with preconcentration of analyte into a microvolume, promises to produce high sensitivity and very low limits of detection. This work addresses the feasibility of a paramagnetic bead-based electrochemical immunoassay with an interdigitated array nanoelectrode as the detection element.
3.2 Characterization of Commercial IDAs
3.2.1 Materials
IDA electrodes (3 µm -5 µm) were from ABTECH Scientific Inc.(Richmond,
VA). K3Fe(CN)6 was obtained from J.T. Baker Chemical Co. (Phillipsburg, NJ). p- aminophenol was from Sigma-Aldrich ( St. Louis, MO ). KCl was obtained from Fisher
Scientific (Fair Lawn, NJ).
3.2.2 Charazterization of IDAs
62 Before attempting immunoassay measurements with our own microfabricated
IDAs, we made measurements with 5 µm finger width, 5 µm gap commercial IDAs in order to work out immunoassay procedures and to identify potential problems that could arise during the immunoassay. Some light microscopy images of the 5 µm IDAs can be seen in Figure 3.2A-C.
Figure 3.2A
63
Figure 3.2B
Figure 3.3C
Figure 3.2A-C. Light microscopy images of 5 µm gap IDAs
64 Electrochemical measurements were made by pipetting a drop of analyte on to the IDA surface, and then carefully placing a silver wire coated with silver chloride as a reference, and a platinum auxiliary wire both into the drop. Cyclic voltammetry was done with
K3Fe(CN)6 to characterize the system. The results can be seen in Figs 3.3 - 3.4. As expected, the measured currents at each electrode are roughly mirror images of each other, with the oxidation current (anodic) being slightly higher.
Dual Potential Cyclic Voltametry of Ferrocyanide on Gold IDA
0.00005
0.00004
0.00003
0.00002
0.00001
10 mM 0 1 mM 0.1 mM -0.00001
Current (A) Current 0.01 mM
-0.00002
-0.00003
-0.00004
-0.00005
-0.00006 0.7 0.6 0.5 0.4 0.3 0.2 0.1 0 -0.1 -0.2 Potential (V)
Figure 3.3. CV with Ferrocyanide on 5µm IDA electrode, 100 mV/sec.
65 Dual Potential Cyclic Voltametry of Ferrocyanide on Gold IDA
0.000006
0.000004
0.000002
0 1 mM 0.1 mM 0.01 mM
-0.000002 (A) Current
-0.000004
-0.000006
-0.000008 0.7 0.6 0.5 0.4 0.3 0.2 0.1 0 -0.1 -0.2 Potential (V)
Figure 3.4. Close up of 100 mV/sec CV on 5 µm IDA with 10 mM concentration traces removed to show detail.
A calibration curve was constructed for PAP over an assay-relevant concentration range and can be seen in Figure 3.5. A linear relationship between current and concentration was obtained for all concentrations down to 312 nM, the lowest concentration measured.
Immunoassay measurements were then made using the MS2 bacteriophage as the analyte. The MS2 samples were prepared, captured on beads, and exposed to secondary antibody labeled with β-galactosidase according to the procedure described in section 3.5.
The detection was done by pipetting a 45 µL drop of PAPG on the array surface, then adding 5 µL of beads. The increasing current associated with the enzyme catalyzed
66 production of PAP was measured, and the slope of this current plotted vs. time was used to quantitate the analyte concentration. The results of the assay can be seen in Figure 3.6.
While the calibration was somewhat linear, the lower concentrations were very irreproducible. We suspect that the detected currents were very sensitive to the manner in which the pipetted beads dispersed on the IDA surface. Because of this irreproducibility, we decided to use a different detection procedure with the nanoIDA.
Another limitation of this technique was that the microbeads were allowed to settle on the array surface. However, inspection of the electrode surface before and after measurements did not show any evidence that the beads damaged the surface.
Figure 3.5. Calibration plot for PAP on 5 micron IDA using anodic current.
67
Calibration Plot 4.5
4
y =R 2 0.4018x - 0.0177 3.5 = 0.9992
3
2.5
2
1.5
1 slope (nA/sec)
0.5
0 0 2 4 6 8 10 12 -0.5 Concentration (ug/mL)
Figure 3.6. Calibration plot for MS2 bacteriophage using 5 µm IDA for detection.
Anodic current was used for the measurement.
3.3 Immunoassay with NanoIDA
3.3.1 IDA Fabrication
The IDA nanoelectrode was fabricated by Xiaoshan Zhu (Department of
Electrical and Computer Engineering, University of Cincinnati) using an e-beam lithography lift-off technique to pattern the nanometer-scaled IDA fingers, and conventional lithography to pattern the contacts. A 300 nm thick layer of PMMA (495K
,Microchem, Newton, MA) was spin-coated onto an oxidized silicon wafer, and then the
68 wafer was baked on a 180 °C hot plate for 2 min. Subsequently, the nanopatterns were exposed by e-beam (Raith 150 e-beam lithography system (Ronkonkoma, NY). After developing the exposed PMMA, a Ti/Au (100 Å/ 1000 Å) layer was deposited on the patterned sample surface using e-beam evaporation, and the deposited sample was dipped into acetone for lift-off. The positive photoresist (Shipley 1818)(Shipley Co.,
Marlborough, MA) was coated on a glass substrate, and then the wafer was baked in a 90
°C oven for 30 min. Next, the baked Shipley 1818 was exposed under UV light, and immersed in chlorobenzene for 45 s. After immersion, the sample was dried in a 120 °C oven for 30 s. The sample was then developed and dried, and a Ti/Au (100 Å/ 1000 Å) layer was deposited on the patterned sample surface. The deposited sample was dipped into acetone for lift-off.
Wires were attached to the working electrodes with conductive silver epoxy, then insulated with insulating epoxy. This process was complicated by the fact that the solvents in the insulating epoxy tended to dissolve the conducting epoxy, resulting in shorted connections. This was solved by insulating the connections in several steps, each step applying the insulating epoxy in very small amounts.
3.3.2 Physical Characteristics of Nanoelectrode
A low-power image of the nanoIDA can be seen in Figure 3.7A. The two working electrodes were wired to the potentiostat with conductive silver-based epoxy, while the on-chip reference and auxiliary were not. The grey square in the center is the actual array, but the fingers are too small to be seen at this scale. Figure 3.7B shows a closer view of the entire array, and Figs. 3.7C-D show a high-magnification view of the
69 individual fingers. These SEM measurements indicate a uniform finger width of approximately 220 nm, with an inter-electrode gap of approximately 620 nm. On some samples, the fingers of the IDA were slightly misaligned (Figure 3.8), although this did not seem to affect the performance of the electrode. The misalignment resulted from the total area of the electrode being larger than the pattern that could be written with the e- beam lithography system, necessitating multiple, successive, e-beam writing steps.
70
Figure3.7A
Figure 3.7B
71
Figure 3.7C
Figure 3.7D
Figure 3.7A-D. SEM images of nanoIDA.
72
Figure 3.8 SEM image of IDA showing the slight misalignment of electrode fingers.
3.4 E. coli Immunoassay
3.4.1 Materials
Dynabeads M-280 streptavidin were obtained from Dynal ASA (Oslo, Norway). Heat- killed E. coli O157:H7 positive control, phosphatase-labeled goat anti-E. coli O157:H7 antibody and goat anti-E. coli O157:H7 antibody were from KPL, Inc. (Gaithersburg, Maryland). PBS powder (10x), THAM, MgCl2 5H2O, 8-tube strip microcentrifuge tubes, round-bottom 96-well microtiter plates, Centricon YM50 and disposable culture glass tubes were from Fisher Scientific
(Pittsburgh, PA). Neodymium-Iron-Boron rare earth magnets were from K & J Magnetics
(Warminster, PA). Sulfo-NHS-LC-biotinylation kit was from Pierce (Rockford, IL). Glycine was from Matheson Coleman & Bell Manufacturing Chemists (Norwood, OH). BSA, biotin and
73 NaN3 were from Sigma (St. Louis, MO). Tween 20 was from Aldrich Chemical Company
(Milwaukee, WI), and PAPP was from MP Biomedicals, Inc. (Aurora, OH).
3.4.2 Buffer Preparation
PBS (10x) was prepared by dissolving one 10x PBS powder package in 1 L of distilled water. The reaction buffer (1x PBS-R) was prepared by adding 5 g of BSA to 2.5 mL of Tween
20 and 50 mL of 10x PBS and diluting the solution to 500 mL with deionized water. The 2x
PBS-R buffer was prepared by adding 5 g of BSA to 2.5 mL of Tween 20 and 50 mL of 10x PBS and diluting the solution to 250 mL with deionized water. The detection buffer (Tris, 0.1 M,
MgCl2, 5 mM, NaN3, 0.02 % (w/v), glycine, 10 mM, pH 9) was prepared by dissolving 3.029 g of THAM, 0.25 g of MgCl2 5H2O, 0.05 g of NaN3 and 0.1876 g of glycine in 200 mL of deionized water. The solution was adjusted to pH 9 and filled up to 250 mL mark of a 250 mL- volumetric flask. PBS buffer (0.1 M) was prepared by dissolving the PBS buffer package in the sulfo-NHS-LC-biotinylation kit into 500 mL of deionized water.
3.4.3 Preparation of Biotinylated Antibodies
Goat anti-E. coli O157:H7 antibody was biotinylated by Jian Tu , a postdoc in our laboratory, using the sulfo-NHS-LC-biotinylation kit from Pierce. Stock NHS-biotin solution
(10 mM) was prepared by dissolving 1.1 mg of NHS-biotin in 200 µL of distilled water. The reaction was initiated by immediately adding 7 µL of the stock solution to a Centricon YM50
(MW50000 cutoff) receiving vial containing 250 µL of 2 mg/mL antibody. The mixture was incubated at room temperature for 30 min with gentle shaking. Reaction by-products were
74 removed using the Centricon YM-50 and centrifugation (1000 g for 2 min) with 0.1 M PBS buffer. The biotinylated antibody was concentrated during centrifugation. The concentration of the biotinylated antibody was determined by measuring UV absorbance at 280 nm using the extinction coefficient of IgG (1.35 for 1 mg/mL IgG). The amount of biotinylation was determined using a HABA assay [16] and determined to be 1-2 biotin/IgG. An equal volume of glycerol was added to the biotinylated antibody to prevent freezing damage and the antibody solution (0.38 mg/mL) was stored at -20o C.
3.4.4 Paramagnetic Bead-Based Immunoassay for E. coli O157:H7
The capture beads (5 µL bead suspension/sample) were prepared in a glass culture tube by mixing a suspension of streptavidin-coated beads (60 µL of 6.7 x10 8 beads/mL) with biotinylated-goat anti-E.coli O157:H7 antibody (60 µL of 38 µg/mL) to give a final concentration of 19 µg/mL. The streptavidin-coated beads were then washed with 60 µL of 1x
PBS-R once and resuspended in 60 µL of 1x PBS-R. The mixture was incubated for 30 min. with gentle shaking and washed with 1x PBS-R to remove the excess antibody. The beads were resuspended in 60 µL of 1x PBS-R and evenly distributed into 12 wells in a round-bottomed 96- well microtiter plate (5 µL beads/well). Then various concentrations of heat-killed E. coli
O157:H7 (5 µL) from 5 x 102 cfu/mL to 5 x 108 cfu/mL, were incubated with the capture beads
(5 µL) for an additional 30 min with gentle shaking along with a buffer blank to evaluate non- specific binding. They were then washed twice with PBS-R. The E. coli-complexed beads were prepared for detection by incubating with 20 µL of 10 µg/mL phosphatase-labeled goat anti-E. coli O157:H7 antibody for 30 min and washed with 1x PBS-R twice, and Tris pH 9 buffer four
75 times. The beads were suspended in 20 µL of Tris buffer and transferred to a strip of microcentrifuge tubes for detection. All the incubations were done at room temperature.
3.4.5 Nano IDA Detection
The E. coli-complexed beads (5 µL) were added to 50 µL of PAPP (1 mg/mL), which was freshly prepared and filtered through a 0.45 µm filter. The bead suspension was mixed with the substrate and incubated at room temperature for 5 min. An aliquot (10 µL) of the supernatant solution was removed while the beads were magnetically immobilized at the bottom of the tube. and placed on the IDA. IDA measurements were made by applying +300 mV and -200 mV potentials to the two working electrodes of the IDA. The current was then recorded for 3 min for each measurement. In between runs the IDA was carefully cleaned with distilled water by gently adding and removing a drop with a pipette. The results of the E. coli assay can be seen in Figure
3.9. The lowest concentration detected that was distinguishable from the blank was 1.2 x 107 cfu/mL, which is not a very low detection limit. However, a very limited number of measurements were actually made because the array failed after several measurements.
76 E.coli immunoassay detected by nanoIDA time (s) 0.1e 0 50 100 150 200 -0.4
-0.9
-1.4
-1.9 Current (µA) -2.4 PAPP 8 -2.9 E.coli= 2.56x10 cfu/mL 7 E.coli= 1.28x10 cfu/mL E.coli= 2.56x105 cfu/mL
Figure 3.9 E. coli detection with nanoIDA
3.5 MS2 Immunoassay
3.5.1 Materials
Dynabeads M-280 streptavidin were obtained from Dynal ASA (Oslo, Norway). Rabbit anti-MS2 antibody was from Tetracore, Inc. (Gaithersburg, MD). MS2 was generously given by
Michael Goode (Aberdeen Proving Ground, MD). β-Galactosidase conjugated to rabbit anti-
MS2 antibody was custom-made by American Qualex (San Clemente, CA). PBS powder (10x),
THAM, MgCl2. 5H2O, 8-tube stripe microcentrifuge tubes, round-bottom 96-well microtiter plates, Centricon YM50 and disposable culture glass tubes were from Fisher Scientific
(Pittsburgh, PA). Neodymium-Iron-Boron rare earth magnets were from K & J Magnetics
(Warminster, PA). BSA, biotin, p-aminophenyl-β-D-galactopyranoside (PAPG) and NaN3 were
77 from Sigma (St. Louis, MO). Tween 20 was from Aldrich Chemical Company (Milwaukee,
WI).
3.5.2 Preparation of the Immunoassay Buffers
PBS (10x) was prepared by dissolving a 10x PBS powder package in 1 L of deionized ultrafiltered water. The reaction buffer (1x PBS-R) was prepared by adding 5 g of BSA to 2.5 mL of Tween 20 and 50 mL of 10x PBS and then diluting the solution to 500 mL with deionized ultrafiltered water. The 2x PBS-R buffer was prepared by adding 5 g of BSA to 2.5 mL of
Tween 20 and 50 mL of 10x PBS and diluting the solution to 250 mL with deionized ultrafiltered water. The detection buffer (PBS-D) was prepared by dissolving 0.25 g of MgCl2 5H2O, 0.05 g of NaN3 in 200 mL of deionized ultrafiltered water and 50 mL of 10x PBS. PBS buffer (0.1 M) was prepared by dissolving a PBS buffer package into 500 mL of deionized ultrafiltered water.
3.5.3 Paramagnetic Bead-Based Immunoassay for MS2
Capture beads (10 µL bead suspension/sample) were prepared by mixing a suspension of streptavidin-coated beads (125 µL of 6.7 x108 beads/mL) with biotinylated rabbit anti-MS2 antibody (125 µL of 80 µg/mL) in a glass culture tube. The streptavidin-coated beads were previously washed with the reaction buffer three times and resuspended in 125 µL of the reaction buffer. The mixture was incubated for 25 min with gentle shaking and washed with the reaction buffer once to remove excess antibody. The beads were resuspended in 125 µL of the reaction buffer and distributed into the wells of a round-bottomed 96-well microtiter plate (10 µL beads/well). Then, the appropriate MS2 concentrations from 10 ng/mL to 20 µg/mL and a buffer blank (10 µL of 10, 20, 50, 100, 200, 500 ng/mL, 1, 2, 5, 10, 20 µg/mL,) were incubated with the
78 capture beads (10 µL) for an additional 10 min with gentle shaking, and washed with the reaction buffer three times. The antigen-complexed beads (10 µL) were prepared for detection by incubating with β-galactosidase-labeled rabbit anti-MS2 antibody (15 µL of 250 µg/mL) for 30 min and washed with the reaction buffer twice, and the detection buffer (PBS-D) four times. All the incubations were done at RT.
3.5.4 Nano IDA Detection
The MS2-complexed beads (5 µL) were added to 45 µL of freshly-prepared PAPG (1.5 mg/mL) in a clean test tube and filtered through a 0.45 µm filter. The bead suspension was mixed with the substrate and incubated at room temperature for 5 min. The beads were immobilized magnetically at the bottom of the test tube and an aliquot (10 µL) of the solution, free of beads, was removed and placed on the IDA The IDA measurement was made by applying +300 mV and -200 mV to the two working electrodes of the IDA. Although reference and auxiliary electrodes were fabricated on the chip, off-chip electrodes were used for the measurements because previous experiments using the on-board electrodes had fried the device.
The current was measured for 3 min. The IDA was rinsed with distilled water between measurements.
3.5.5 Immunoassay Results
A bead-based assay was done to capture the MS2 on 5 µL of beads. The beads were mixed with 45 µL of substrate, and after a 5 min. incubation, a 10 µL detection volume was taken from the supernatant while the beads were immobilized with a rare earth magnet at the bottom of the tube. As illustrated in Figure 3.10, the drop containing enzyme product was placed
79 directly on the array, and the fine wires used for reference and auxiliary electrodes were inserted carefully. A limitation of this arrangement is that the microdrop tends to evaporate rapidly.
Therefore, measurements were made immediately after the drop was placed on the array.
Unfortunately, only a limited number of measurements could be made before the electrode failed. Measurements for the blank, 10, 20, and 50 ng/mL were made successfully, then the IDA was damaged during the 100 ng/mL sample detection. We are unsure if the IDA was destroyed by debris, or if the current generated by the higher analyte concentration caused the failure. The electrode response as a function of time can be seen in Figure 3.11. It can be seen that the anodic current was slightly larger than the cathodic. This is due to the fact that the redox cycling is not
100% efficient, and it is the PAP that is formed as the enzyme product that is oxidized. After an initial decline, the currents stabilized after approximately 40 s. A possible explanation for the current reduction is electrode fouling, which we have observed in prior work with PAP. The cathodic currents at 100 s were plotted vs. the analyte concentration for the calibration plot seen in Figure 3.12. 100 s was chosen because the currents seemed fairly stable by this time, although some instability and a decrease in current was observed at approximately 140 seconds for the 50 ng/mL sample. The cathodic currents were used because they were slightly more stable than the anodic, and both were nearly the same magnitude but opposite in charge due to the redox cycling. The detected signal for an MS2 immunoassay rose as concentration was increased between 10 ng/mL and 50 ng/mL. The data suggest that detection limits below this are definitely possible, as the noise stabilized greatly after a few minutes.
A limitation of an enzyme label system, such as β-galactosidase and PAPG, that converts substrate to a detectable product is that no concentration of substrate is optimal for all analyte concentration ranges. At high analyte concentrations, a higher substrate concentration is
80 required to ensure that product formation is not reduced due to substrate depletion. This is especially important in microliter volumes. When detecting very low concentrations, the background signal caused by the non-enzyme-catalyzed conversion of substrate to product, as well as the slight electrochemical activity of the substrate, limits the detection sensitivity. More optimization of substrate concentration and sample volume could be done to lower further the limit of detection. The arrays tended to fail after several measurements, so analysis of more data points and further optimization of this system was not possible. Despite the observed fragility, the performance of the IDA electrodes illustrates that sensitive nano or even pico-volume measurements may be possible that can’t be done with traditional electrodes. When cycling was stopped by turning “off” the potential of the cathodic fingers with the anodic set to an oxidizing potential, the current was unmeasurable at even the highest assay-relevant concentrations. Based on these observations, we estimate the current enhancement to be greater than 100X compared to the system without cycling.
Ag/AgCl Platinum Reference Auxiliary
Working + Working - Silicon Substrate
Figure 3.10. Diagram showing how immunoassay measurements were made on the IDA.
81
Figure 3.11. Plot of the current response as a function of time for immunoassay samples containing 0(a), 10(b) 50 (c) and 100 (d) ng/mL of MS2 phage.
82 Calibration Plot
-5
-4.5
-4
-3.5
-3
-2.5
Current (nA) Current -2
-1.5
-1
-0.5
0 0 102030405060
Concentration (ng/mL)
Figure 3.12. Calibration curve for MS2 using nanoIDA
3.6 Conclusions
3.6.1 Problems Associated with the Microscale
As can be seen in the 10x view of the IDA (Figure 3.7A) the electrodes were designed to have microfabricated auxiliary and reference electrodes in addition to the dual working electrodes. The reference electrode was to be plated with Ag/AgCl, and the small auxiliary electrode would only be required to carry a small amount of current because the two working electrodes are nearly identical in area and should almost completely balance out their currents.
After numerous destroyed electrodes, it was determined that the small reference and auxiliary electrodes were being damaged soon after the cycling was initiated. We surmise that during the course of an experiment the differences in current between the two working electrodes became too much for the small counter electrode to balance in an acceptable potential range, resulting in
83 destruction of the electrode. As a result, a fine silver wire coated with silver chloride was used as a reference, and a small platinum wire as the auxiliary.
The electrodes were also very susceptible to damage from dust particles or stray microbeads from the immunoassay. Care was taken to keep the electrodes covered at all times, and to filter any fluids that would touch the electrode surface. Despite these precautions, the electrodes tended to fail after a few hours of use. SEM images of a damaged electrode (Figure
3.13A-B) illustrate the damage that can be caused by a small piece of debris. It is suspected that a dust particle from the air or a stray microbead damaged the surface during the gentle distilled water rinse. This is probable because the array fingers are so small that a 2.8 micron bead is large enough to settle across two to three of the gaps. In order for a device of this size to be practical and reusable, a protective strategy must be devised. Possiblities include a permeable protective membrane, a flow cell, or a microfluidic network to isolate the array from particles in the air.
Figure 3.13A. Damaged IDA electrode
84
Figure 3.13B Closeup of damaged region of IDA electrode.
3.6.2 E. coli Assay
The E. coli assay was in many ways a learning experience that led to the much more successful MS2 measurements, even though the number of MS2 measurements was very limited as well. Several important lessons were learned concerning precautions that must be observed when working with such a fragile device. All fluids, including the deionized water rinse, must be filtered immediately before coming into contact with the IDA surface. Also, the IDA surfaces could not be exposed to the ambient air for any significant amount of time, or particles would settle on the surface. This assay demonstrated the challenges associated with interfacing a milli- to micro- volume procedure with a nanometer scaled detection element, but did not result in more sensitivity or a lower limit of detection than conventional microelectrode detection. 3.6.3
MS2 Assay
85 A highly sensitive immunoassay for MS2 bacteriophage was shown using a sub-micron gap interdigitated array nanoelectrode. The limit of detection achieved was nearly 20 times greater than with a rotating disk electrode, and nearly ten times greater than with a 2.4 µm IDA.
The information in Table 1 illustrates the sensitivity that can be gained by redox cycling using essentially the same immunoassay procedure for MS2 as with other electrode types and geometries. The redox cycling on the interdigitated array electrodes generated a greater current amplification than the enhanced diffusion due to hydrodynamic flow on the rotating disk. In addition, a smaller detection volume is required for measurement with the IDA, further improving the detectable amount.
Electrode type Limit of Detection Amplification
Rotating Disk 200 ng/mL [6] convection/diffusion
Electrode
2.4 µm gap IDA 90 ng/mL[6] diffusion/redox
cycling
620 nm gap IDA <10 ng/mL diffusion/redox
cycling
Table 1. Comparison of nanoIDA results to previous work with the same MS2 assay.
3.6.4 Future outlook
Nanometer scaled IDAs show great promise for sensitive amperometric detection in small volumes. For bead-based immunoassay applications to become practical, however, strategies must be developed that limit the possibility of beads or other debris coming into contact with the electrode. The microfluidic strategies discussed in Chapter 2 offer a solution to
86 the problem of debris from the room falling on the IDA surface, but the precise manipulation of microbeads on a scale smaller than the naked eye can see remains a significant challenge.
87
References
[1] A. C. Micheal, and R. M. Wightman, Microelectrodes, in Laboratory Techniques in
Electroanalytical Chemistry 2nd Ed, in Kissinger, P. T., and Heineman, W. R., (Eds.), Marcel
Dekker, 1996, pp. 367-402.
[2] C. A. Wijayawardhana, H. B. Halsall, and W. R. Heineman, Micro volume rotating disk electrode (RDE) amperometric detection for a bead-based immunoassay, Analytica Chimica
Acta 399 (1999) 3-11.
[3] Osamu Niwa, Electroanalysis with interdigitated array microelectrodes, Electroanalysis 7
(1995) 606-613.
[4] O. Niwa, Y. Xu, H. B. Halsall, and W. R. Heineman, Small-volume voltammetric detection of 4-aminophenol with interdigitated array electrodes and its application to electrochemical enzyme immunoassay, Analytical Chemistry 65 (1993) 1559-1563.
[5] M. Diaz-Gonzlez, M. B. Gonzlez-Garcia, and A. Costa-Garcia, Recent Advances in
Electrochemical Enzyme Immunoassays, Electroanalysis 17 (2005) 1901 - 1918.
[6] J. Thomas, in Chemistry, 2003 p. 181 University of Cincinnati, Cincinnati.
[7] J.-W. Choi, K. W. Oh, J. H. Thomas, W. R. Heineman, H. B. Halsall, J. H. Nevin, A. J.
Helmicki, H. T. Henderson, and C. H. Ahn, An integrated microfluidic biochemical detection system for protein analysis with magnetic bead-based sampling capabilities, Lab on a Chip 2
(2002) 27-30.
88 [8] J. H. Thomas, S. K. Kim, P. J. Hesketh, H. B. Halsall, and W. R. Heineman, Bead-Based
Electrochemical Immunoassay for Bacteriophage MS2, Analytical Chemistry 76 (2004) 2700-
2707.
[9] O. Niwa, M. Morita, and H. Tabei, Electrochemical behavior of reversible redox species at interdigitated array electrodes with different geometries: consideration of redox cycling and collection efficiency, Analytical Chemistry 62 (1990) 447-452.
[10] M. Paeschke, U. Wollenberger, C. Kohler, T. Lisec, U. Schnakenberg, and R. Hintsche,
Properties of interdigital electrode arrays with different geometries, Analytica Chimica Acta 305
(1995) 126-136.
[11] A. J. B. Junhong Min, Characterization and Optimization of Interdigitated
Ultramicroelectrode Arrays as Electrochemical Biosensor Transducers, Electroanalysis 16
(2004) 724-729.
[12] K. Aoki, M. Morita, O. Niwa, and H. Tabei, Quantitative analysis of reversible diffusion- controlled currents of redox soluble species at interdigitated array electrodes under steady-state conditions, Journal of Electroanalytical Chemistry 256 (1988) 269-282.
[13] M. Paeschke, T. Lisec, U. Schnakenberg, R. Hintsche, and U. Wollenberger, Highly sensitive electrochemical microsensensors using submicrometer electrode arrays, Sensors and
Actuators B: Chemical 27 (1995) 394-397.
[14] A. E. Cohen, and R. R. Kunz, Large-area interdigitated array microelectrodes for electrochemical sensing, Sensors and Actuators B: Chemical 62 (2000) 23-29.
[15] X. Zhu, J.-W. Choi, and C. H. Ahn, A new dynamic electrochemical transduction mechanism for interdigitated array microelectrodes, Lab on a Chip 4 (2004) 581-587.
89 [16] V. G. Janolino, J. Fontecha, and H. E. Swaisgood, A spectrophotometric assay for biotin- binding sites of immobilized avidin, Applied biochemistry and biotechnology. Part A : enzyme engineering and biotechnology (1996) 1-7.
90
Chapter 4:
Carbon Nanotube-Based EIS Biosensor
91
4.1 Introduction
4.1.1 Chemical Structure of Carbon Nanotubes
Carbon nanotubes (CNTs) are allotropes of carbon, and have a structure that can be described as planar sheets of graphite (graphene) rolled into cylinders. The diameter of the tubes is in the low nanometers, while the lengths can be as long as 10’s of millimeters. Because of the covalent bonding between carbon atoms along the length of the tube, carbon nanotubes are very strong in that direction. The bonding structure of CNTs is similar to that of graphite, in that the carbon atoms are sp2 hybridized, thus containing both σ-bonds and π-bonds. The strength of the sp2 hybridized carbon bond is what gives nanotubes their amazing tensile strength[1].
There are several different nanotube structures that correspond to the different helicities, or ways that graphene sheets can be “rolled up”. The structures are classified as “armchair”,
“zigzag” and “chiral”, and can be seen in Figure 4.1.[2]. The helicity of the nanotube affects the electronic characteristics of the nanotubes, as armchair nanotubes are conducting, while zigzag and chiral nanotubes can be either conducting or semiconducting[3].
92
Figure 4.1 Diagram showing different ways of “rolling” graphene to form nanotubes, showing a) armchair, b) chiral, and c) zig-zigzag conformations. Adapted from Saito et al.[2].
CNTs may be of single wall (SWCNT) or multi wall (MWCNT) construction. SWCNTs consist of a single graphene sheet rolled into a tube, while MWCNTs consist of multiple layers of such tubes rolled up to form concentric cylinders. Because MWCNTs consist of tubes of many different diameters, they exhibit metallic conductivity, as statistically the majority of the tubes will be conductive [4]. A brief comparison of SWCNTs and MWCNTs is seen in Table
4.1
93
Single Walled Nanotubes Multi Walled Nanotubes
1-5 nm diameter 2.5-30 nm diameter generally higher purity, but cost more approximately 0.33 nm between layers used in molecular electronics more prone to structural defects may be metallic or semiconducting much less expensive to produce
Table 4.1. Comparison of single walled and multi walled nanotubes.
4.1.2 Physical Properties of Nanotubes
Carbon nanotubes are some of the strongest materials ever discovered, with a tensile strength on the order of 63 GPa[5], roughly 100 times that of steel. They are also extremely stiff, with an elastic modulus near 1 TPa. Because they have a relatively low density compared to other ultrastrong materials (1.3 g/cm3), they have the highest specific strength (strength divided by density) of any known material[6]. These remarkable mechanical properties make carbon nanotubes a very attractive material for micro- and nanoscale devices, especially for structures such as needles that are exposed to high levels of mechanical stress.
4.1.3 Electrical Properties
Carbon nanotubes also possess interesting electrical properties that make them suitable for molecular electronics, sensing, and other nanomaterial applications [7-12]. Of particular interest for electrochemical sensing applications is the high conductivity of metallic nanotubes.
94 Nanotubes have also been shown to conduct electrons ballistically, which means that individual electrons move down the length of the tube without scattering and losing spin information. This effect is very important for molecular wiring and electronics, but has not been used for the type of sensing applications described in this dissertation. One limitation of nanotubes for sensor devices is the high contact resistance observed between nanotubes and metals, although there is work being done to minimize this effect[13]
4.1.4 Sensing Applications
Because of their unique properties, there has been much interest in using carbon nanotubes for sensing applications[14]. Probably the most commonly seen electrochemical sensors using carbon nanotubes are enzyme modified electrodes [15-20]. The common enzyme, substrate, and mediator schemes used for nanotube electrodes are similar to those seen with other enzyme electrodes. In general, the enzyme of interest is used to catalyze reactions involving biologically relevant molecules, such as glucose, lactate, pyruvate, or glutamate, to form electrochemically detectable products such as H2O2 or NADH. In other cases, a mediator is used instead of the direct detection of redox active product. Other sensor systems involving nanotubes include DNA biosensors[21], immunosensors[22],and impedance based sensors[23-25].
4.1.5 Electrochemical Impedance Spectroscopy
Electrochemical impedance spectroscopy (EIS) is an electrochemical technique that makes use of a phase-sensitive voltmeter (lock-in amplifier) to measure the phase and amplitude of the resulting current when a sinusoidal wave voltage pulse is applied. EIS can be used to investigate the dynamic interface between the electrode and the solution, since the electron
95 transfer, mass transport, and chemical reaction of redox species will all contribute to a change in phase and amplitude of the output signal.
Impedance is similar to resistance in that it is measured as the ratio of applied voltage to measured current, but resistance is for direct current rather than alternating current. Impedance measurements are made by applying a low amplitude AC voltage over a small potential range
(generally 5-10 mV), and then measuring the current. This measurement is then made over a large frequency range, typically 105 Hz or higher down to 0.1 Hz or lower. Because different processes, such as electron transfer, diffusion, double layer charging, etc., have different kinetics, their effects show up in different frequency regions. For example, capacitive effects such as double layer charging are dominant at high frequency, while the low frequency region is dominated by diffusion and surface properties.
Traditionally, EIS has been used to characterize circuits and to evaluate corrosion and insulative coating failure.[26] Recently EIS has become popular for sensing applications because it can directly measure surface binding without the use of a label [27-31]. When a chemical change is made to the surface of an electrode, the electric properties such as dielectric constant and resistance will change, and can be detected by EIS. This change is due to the binding of the molecule itself, and so a label is not required. However, some labels can be used to augment the sensitivity of EIS by drastically changing the surface properties of the electrode [32]. The exact nature of the change of electronic surface properties is not known, but many theoretical models have been constructed considering such factors as displacement of water, change in dielectric properties, and charge state [33]. A general observation has been that the most dramatic changes in EIS are observed when the analyte has drastically different properties than the probe molecule that is immobilized on the electrode.
96 One way that the results of impedance measurements can be displayed is by a Nyquist plot, which plots the real vs. imaginary components of the measured signal. Although many different shapes are possible, the typical shape observed in sensing applications where the impedance decreases as frequency increases is that of a semicircle as seen in Figure 4.2. In well characterized systems, the information presented in the Nyquist plot can be used to generate an equivalent circuit model, which can then be used to estimate a number of key paramaters of an electrochemical system, including the diffusion coefficient of the redox species, the faradaic time relaxation constant τ, and the double layer capacitance [26]. A simpler interpretation that is often seen in sensors work, as opposed to circuit characterization, is to correlate an increased electron transfer resistance with an increase in insulating material bound to the electrode.
Selectivity in this type of system is achieved through the probe molecule immobilized on the electrode surface rather than through the characteristics that can be determined through EIS. This type of measurement, while not using the full capability of EIS spectroscopy, can still be very useful for label-free sensing.
97
Figure 4.2 Typical Nyquist plot for sensing applications.
4.1.6 Goals of This Work
Typical immunoassays are multi-step procedures that involve a capture step, multiple rinses, adding a label for detection and/or amplification, and finally detection.[34] These methods can be very sensitive, and are perfectly suitable for laboratory analysis, but are limited for some applications that require continuous monitoring, rapid response, and portability. Electrochemical impedance spectroscopy (EIS), coupled with the selectivity and sensitivity of biological recognition molecules such as antibodies promises to be an effective label-free sensing technique for biomolecules.[35] EIS has advantages over many other label-free sensing techniques in that it can be used for continuous monitoring, it does not destroy the sample, and the only hardware
98 needed consists of the electrodes and potentiostat. Once optimized, EIS-based biosensors may be able to provide rapid, cheap, dependable and sensitive detection of many biological molecules in sample volumes from the bulk scale of lakes and rivers to sub-microliter clinical samples.
The physical properties of carbon nanotubes make them an excellent electrode material for an EIS biosensor. The aligned MWNTs used in this study are highly conductive, and, when functionalized, the carbon surface can be attached directly to antibodies or other sensing biomolecules via covalent chemical bonds. Also, the remarkable mechanical strength of the nanotubes will enable future biosensors to be needle-thin, and being able to control the molecular geometry of the nanotubes becomes a powerful tool for creating nanoscale structures that optimize the electrode’s analytical performance. A schematic of the sensor can be seen in Figure
4.3[36].
Figure 4.3. Schematic of the nanotube immunosensor (not to scale)
The surface of the electrode can be seen as a forest of exposed conducting nanotubes with antibodies on the ends, immersed in a block of insulating material. The design goal was to further reduce the current observed per molecule bound to the sensor by limiting the electrode surface to
99 an array of small points that can be easily obstructed by target molecules, as illustrated in Figure
4.4. Although this scheme was the goal, fabrication and chararacterization limitations never clearly established that this design was ever truly achieved on the nanoscale. Still, the sensors made and the results obtained indicate that the impedance biosensors based on carbon nanotubes are potentially very useful. As our knowledge and ability to more precisely control the nanoscaled patterning and orientation of nanotubes increases, we expect that more sensitive, reliable, and well characterized sensing devices will be possible.
A low number of blocking A high number of molecules occlude a molecules are required significant portion of the to occlude a significant conducting surface portion of the surface
Conductor
Insulator
Figure 4.4. Advantage of using nanotube composite for electrode material over bulk surface.
4.2 Materials and Methods
100 4.2.1 Synthesis of Nanotubes
P-type 4 in. diameter Si wafers <100> with a typical resistivity of 1-20 Ohm-cm were used. An E-beam evaporator (Airco Temescal, Fairfield ,CA)was used to deposit a 10 nm Al thin film. Then the Al was oxidized using reactive ion etch (RIE) to form a layer of Al2O3. Finally, catalytic iron films of controlled thickness between 1 and 2 nm were deposited on Si, SiO2, and
-7 Al2O3 surfaces using e-beam evaporation. The evaporation was done at approximately 5 x 10 torr and the system was equipped with a film thickness monitor. The final substrate was cut into
5 mm x 5 mm squares with a dicing saw.
The nanotubes were synthesized by thermal CVD in a horizontal 2 in tube furnace,
(EasyTubeTM ET1000, FirstNano, Ronkonkoma, NY), which consists of four mass flow controllers and a vapor delivery system. Argon was used to carry the water vapor to the reaction chamber, and to purge the reactor for 20 minutes while the CVD furnace was heated to 750°C.
The gas flow was then switched to ethylene, water, and hydrogen for specific lengths of time based on process parameters. During this time the hydrocarbon precursor, ethylene, reacts with the catalyst and deposits CNT on the substrate. After the nanotube array was synthesized it was cooled to ambient temperature, completing the last process step. During cooling, ethylene, water, and hydrogen flow were stopped and the system was purged with argon to prevent back flow of air from the exhaust line. The synthesized CNT arrays were characterized by environmental scanning electron microscopy (ESEM).
4.2.2 Biosensor Fabrication
The aligned nanotubes were peeled off the silicon substrate and cast in epoxy using Epon
Resin 862 and the EPICURE curing agent W (Hexion Speciality Chemicals, Houston, TX). The
101 epoxy insulates the nanotubes thermally and electrically, and provides mechanical support. The bottom side of the array was then polished to expose the aligned nanotubes for connection.
Conductive epoxy was used to attach copper wire to the bottom of the array, and insulating epoxy was used to seal the connection. The top side of the array was then polished to expose the nanotubes. A summary of this procedure can be seen in Figure 4.5[36].
102
Figure 4.5. Steps of biosensor fabrication
4.2.3 Immobilization of Antibody
Antibodies were attached covalently to the nanotubes by first oxidizing the electrode surface to present carboxyl surface groups. The nanotubes were opened using H2SO4/70% HNO3, and HCl, followed by electrochemical treatment at 1.5 V (versus Ag/AgCl) in 1.0 M NaOH for 30 s. The carboxyl groups were activated with a 20 minute incubation in a 500 µL solution containing 10 mg of 1-ethyl-3(3-dimethylaminopropyl) carbodiimide hydrochloride (EDC) and 5 mg sulfo-N-
103 hydroxysuccinimide (sNHS) to stabilize the acylisourea intermediate. The activation was quenched in a 1% ethanolamine solution to stop further carbodiimide activity. The electrode was then incubated in an antibody solution containing 20 µL of approximately 1.5 mg/mL donkey anti-mouse IgG (Jackson Immunoresearch Laboratories Inc. West Grove, PA) in 1 mL PBS, pH
7.0 for four hours. A description of this immobilization procedure can be seen in Figure 4.6. The resulting product of antibody covalently attached to the carbon nanotubes was rinsed with deionized water and stored at 4º C and pH 7 until used for testing.
104
H2C CH2 Na CH3 H2C CH3 N H2C N C N CH3 H2C CH3 N H CH H C N N CH H 3 2 3 C C C H2 H2 O O O O C C a. b. nanotube
O O- c. S O d. CH CH2
C C Antibody O O N
O N O O C H C
Figure 4.6. Coupling strategy used to attach antibody to carboxyl groups on exposed nanotube surface. a) EDC is used to activate carboxyl group, b) unstable acyl-isourea intermediate is formed, c) Sulfo-NHS displaces the EDC to form stable intermediate, and d) stable amide bond is formed to amine group on antibody.
The model analyte for this system was mouse IgG, despite the literature indications that a more robust signal could be obtained if the target molecule had significantly different size and surface properties than the probe. The mouse IgG was selected due to the fact that it was readily
105 available and is commonly used as a model analyte, so it can thus be directly compared to literature results in other sensing systems.
Mouse IgG was not the first choice of analyte that we attempted to sense. Initial experiments using both anti-E. coli antibody and polymyxin B to sense for E. coli were unsuccessful. However, irreproducibility of the electrode surface properties was a problem that was continually observed, so it is possible that the unsuccessful results were a result of the electrode fabrication or attachment chemistry rather than an incompatibility of the technique.
4.2.4 Electrochemical Analysis
The electrochemical properties of the biosensor were evaluated by CV with a
Bioanalytical Systems (BAS), Epsilon system with a C3 cell stand with a Faraday cage. A platinum wire and Ag/AgCl reference electrode were used as the auxiliary and reference electrodes, respectively. K3Fe(CN)6 (99% Fisher, Pittsburgh, PA), and KNO3 (99% Fisher,
Pittsburgh, PA) were all prepared fresh for experiments.
The EIS was done with three-electrode cells, with the nanotube electrode as the working electrode, a Ag/AgCl reference electrode, and a platinum wire as the counter electrode. The antigen was mouse IgG from Jackson Immunoresearch Laboratories Inc. (West Grove, PA). EIS measurements were taken with a Gamry Potentiostat (Model: PCI4/750) coupled with the EIS
(Gamry, EIS300, Warminster, PA) software. All testing was done at 0 V DC and 0.1 Hz to 300
KHz, and the sinusoidal potential magnitude was ±20 mV in the redox probe, which was 5 mM
K4[Fe(CN)6], K3[Fe(CN)6] in PBS (pH 7.0).
4.3 Results/Disscussion
106 4.3.1 Nanotube Characterization
Figs. 4.7(a-b) [36] show the ESEM results for the water-assisted nanotube synthesis done by YeoHeung Yun with fixed growth conditions of: 200 SCCM of H2 flow, 200 SCCM of C2H4 flow, and at 750° C with 3 h growth time. Fe/Al2O3/SiO2/Si (5 mm × 5 mm) was cut from one wafer and Fe was patterned into 100 µm × 100 µm squares with 100 µm space in between using a shadow mask. The length of CNT arrays continued to increase for up to 3 hours of growth time.
The resulting CNT forest is highly aligned and has high density, making it suitable for further sensor development. The nanotubes can be easily peeled off of the silicon substrate to be used for sensor construction. Figure 4.7(c) is the ESEM image of aligned MWCNT patterned arrays showing the alignment and high density. Figure 4.8 shows the final fabricated device. The nanotube average diameter was 20 nanometers, with an aspect ratio of 200,000:1. The resistivity of the nanotube tower was measured by casting a nanotube tower with epoxy and polishing both ends of the nano-composite, and then connecting a copper wire to the electrode using conductive epoxy. The volume resistivity is approximately 0.11 ohm ⋅ cm.
107
Figure 4.7A
Figure 4.7B
108
Figure 4.7C
Figure 4.7. ESEM images of (A) aligned high density MWCNT patterned arrays on Si substrate,
(B) side view of patterned nanotube array, (C) high resolution view of individual nanotubes in high density array. This array was fabricated with the growth conditions of: 200 SCCM of H2 flow, 200 SCCM of C2H4 flow, 100 SCCM bubbler flow, 750º C growth temperature.
109 Figure 4.8. Wired nanotube tower-epoxy electrode.
4.3.2 Electrochemical Analysis
The results of a cyclic voltammetry scan using the K3Fe(CN)6 redox probe are shown in
Figure 4.9. A decrease in plateau current for reduction and oxidation is observed with each successive functionalization step. The figure shows, that immobilizing the antibody on the nanotube surface reduces the observed current. This is due to the diffusional barrier of a large insulating molecule being attached to the electrode surface. Exposure to the target antigen resulted in a further reduction in current, as the conducting surface .became more insulated.
(a)
(b)
(c)
110 Figure 4.9. Cyclic voltammetry of 6 mM K3Fe(CN)6 in 1.0 M KNO3 with nanotube tower electrode with 100 mV/s scan rate: after (a) functionalization of nanotube array, (b) immobilization of antibody and (c) antigen binding
The Nyquist plots of EIS results in Figure 4.10[36] show that the sensor response depends on analyte concentration. For a control experiment, an electrode functionalized with anti-mouse
IgG was used, but donkey IgG was used as the analyte. No dramatic increase in impedance was observed. The presence of the target antigen (mouse IgG) bound to the immobilized antibody progressively increased the electron transfer resistance, as indicated by the radius of the curve.
The plot of concentration vs. impedance derived from this plot can be seen in Figure 4.11. The lowest concentration successfully detected and clearly distinguished from the blank with this sensor was 500 ng/mL. This is not very sensitive by immunoassay standards, as some highly sensitive procedures have detection limits in the pg/mL range or below[34]. However, a recent publication by Kuramitz et al. used an automated fluidic system to detect another protein analyte, ovalbumin[37]. In that system a 470 ng/mL detection limit was achieved, so the 500 ng/mL detection limit for IgG using a rapid analysis technique is still relevant. Also, the same system also reports a detection limit of 990 ng/mL for MS2 phage. Thus, a rough calculation of comparative detection limits, assuming that ovalbumin and IgG behave similarly, places the EIS biosensor roughly 100 times less sensitive than the bead based assay using the IDA for detection discussed in Chapter 3. This is expected, however, as the EIS biosensor is still early in development, and achieves detection in essentially one step.
111 Concentration Plot
4.5
4
3.5
3 a b 2.5 c d 2 e f
1.5 Z Imaginary (MegaOhm)
1
0.5
0 024681012 Z Real (MegaOhm)
Figure 4.10. Electrochemical impedance spectra of immunosensor response with the addition of different concentrations of antigen (inner to outer curve); (a) without antigen, with (b) 500 ng/mL,
(c) 1µg/mL, (d) 5 µg/mL, (e) 10 µg/mL, and (f) 100 µg/mL of antigen.
112 Calibration Plot
18
16
14
12 Z (MOhms) 10
8
6 0 20 40 60 80 100 Concentration (µg/mL)
Figure 4.11. Calibration plot for mouse IgG on the impedance sensor. The Z values used for this plot were obtained by taking the square root of the sum of the squares of the real and imaginary components of the impedance at a frequency of 0.1 Hz (the lowest frequency measured).
In Figure 4.12[36], the response of the sensor was monitored as a function of time. The electron transfer resistance increased as more antigen became bound to the electrode, until a saturation point was reached. The fact that the majority of the signal change occurred during the first 30 min. of incubation indicates that this system is suitable for rapid detection. Faster measurements were not possible because the entire impedance spectrum was measured. For a
113 more optimized sensor, a single frequency could be monitored, dramatically reducing analysis time.
Incubation Plot
4
3.5
3
2.5 a b 2 c d e 1.5
1 Imaginary Z (MegaOhm) Imaginary
0.5
0 0 2 4 6 8 10 12 Real Z (MegaOhm)
Figure 4.12. Electrochemical impedance spectra for a nanotube immunosensor with (a) donkey anti-mouse IgG immobilization, and mouse IgG binding after the incubation for half (b), one (c), two (d) and four (e) hours incubation (inner to outer curve) EIS is done at DC potential 0 V with frequencies between 0.1Hz and 300KHz. Sinusoidal potential magnitude is ±20 mV in 5 mM
K4[Fe(CN)6], K3[Fe(CN)6] with PBS (pH 7.0).
114
4.4 Conclusions and Future Directions
Although label-free measurements were made that corresponded to an increase in bound analyte, a much more detailed characterization of the biosensor is required in order to understand the nature of the EIS signal. A number of factors limited our ability to fabricate, functionalize, and test the nanotube biosensors. One major obstacle was the irreproducibility of the electrode surface. Repeated EIS measurements after polishing and functionalizing the nanotube/epoxy composite surface resulted in widely different impedance values. Identical functionalization and polishing procedures yielded results with up to an order of magnitude difference in impedance.
Polish and functionalization steps were required to remove the probe and analyte antibodies after each series of experiments, so it was impossible to make a direct comparison of results obtained from different electrodes, or even the same electrode before and after polishing. Because of this, future sensors will use a reactive ion etch (RIE) plasma polish rather than a mechanical one.
Attempts were made to verify the amount of immobilized antibody by using fluorescently labeled probes. We were not able to make quantitative measurements because the background fluorescence of the conductive epoxy provided too much background emission to observe the labeled probes. Alternative host materials were evaluated, with very limited success. A brief summary of rejected host materials can be seen below in Table 4.2.
115
Material Benefits Limitations tetraethoxysilane (TEOS) inert, insulating, limited to < 50µm layers,
transparent glass shatters when polished methyltriethoxysilane (MTES) similar to TEOS but less fragments when polished
prone to shattering poly (methyl methacrylate) more transparent than high background, cannot
(PMMA) epoxy polish like epoxy. poly(dimethylsiloxane) optically transparent, easy too soft to polish
to cast in thick layers
Table 4.2 Rejected host materials
The polish step was the critical shortcoming of several of these materials, and so they may be viable host materials once the plasma etching system becomes available.
An alternative method for constructing the arrays would be to use very short nanotubes (<
10 µm) rather than the millimeter plus arrays currently used. The advantage of this method would be that the well established coatings such as spin-on glass or silicon nitride could be used for the insulating material. The problem with short nanotubes is that they must be grown on a conductive substrate, because they are too small to peel from a wafer and be wired with conductive epoxy.
This presents a problem, as most vapor-deposited metallic films cannot stand up to the temperatures required for nanotube deposition. To solve this problem, highly doped silicon can be used as the substrate material[38].
116 This study has shown the feasibility of an EIS-based biosensor using carbon nanotubes as an electrode material. Admittedly, a great deal more research is required to make it a practical alternative to the existing heterogeneous immunoassay procedures typically used today. Further experiments will evaluate non-specific binding and its effect on sensor performance, different sizes, geometries, and fabrication techniques, alternative molecular recognition strategies, and further investigation of the EIS technique.
117
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Appendix A: Publications List
Microfluidic immunosensor systems. Biosensors & Bioelectronics 20, 2488-2503, (2005). A Bange, HB Halsall, WR Heineman.
Carbon nanotube array immunosensor development.. IEEE Nano2006 2, 766-769. A Bange, HB Halsall, WR Heineman, Y-H Yun, MJ Schulz, V Shanov.
Microscale immunosensors for biological agents. (In: Microfluidics, BioMEMS and Medical Microsystems III. eds I Papautsky, I Chartier) Proc SPIE 5718, 142-151 (2005). A Bange, DK Wong, CJ Seliskar, HB Halsall, WR Heineman.
Adsorption of fluorescently labeled microbeads on PDMS surfaces. (In: Microfluidics, BioMEMS and Medical Microsystems III. eds I Papautsky, I Chartier) Proc SPIE 5718, 159-168 (2005). I Nikvecic, A Bange, ET Peterson, I Papautsky, WR Heineman, HB Halsall, CJ Seliskar.
Carbon nanotube array smart materials. Proc SPIE, 617205-617212 (2006) in Smart Structures and Materials 2006: Smart Electronics, MEMS, BioMEMS, and Nanotechnology (Varadan, V. K., Ed.). YH Yun, A Bange, VN Shanov, WR Heineman, HB Halsall, SK Pixley, M Behbehani, Z Dong, Y Tu, S Yarmolenko, S Neralla, MJ Schulz.
High sensitivity carbon nanotube tower electrodes. Sensors & Actuators B 120, 298-304 (2006). YH Yun, VN Shanov, MJ Schulz, Z Dong, A Jazieh, WR Heineman, HB Halsall, DKY Wong, A Bange, Y Tu, S Subramaniam.
A nanotube composite microelectrode for monitoring dopamine levels using cyclic voltammetry and differential pulse voltammetry. Proc Institution of Mechanical Engineers, Part N, J Nanoengineering and Nanosystems (2006). Y-H Yun, A Bange, VN Shanov, WR Heineman, HB Halsall, DKY Wong, M Behbehani, S Pixley, A Bhattacharya, Z Dong, MJ Schulz, in press.
The Columbi eggs of nanotechnology. IEEE Nano2006. 2, 698-701 MJ Schulz, Y-H Yun, VN Shanov, S Neralla, S Yarmolenko, J Sankar, Y Tu, A Gorton, G Choi, G Seth, A Bange, HB Halsall, WR Heineman.
A carbon nanotube needle biosensor. J Nanosci & Nanotech. 7, 2293-2300 (2007) Y-H Yun, A Bange, VN Shanov, WR Heineman, HB Halsall, Z Dong, A Jazieh, Y Tu, DKY Wong, S Pixley, M Behbehani, MJ Schulz.
123 A nanotube array immunosensor for direct electrochemical detection of antigen-antibody binding. Sensors & Actuators: B: Chemical 123, 177-182 (2007). Y-H Yun, A Bange, WR Heineman, HB Halsall, VN Shanov, Z Dong, S Pixley, M Behbehani, A Jazieh, Y Tu, DKY Wong, A Bhattacharya, MJ Schulz.
Carbon nanotubes grown on stainless steel to form plate and probe electrodes for chemical/biological sensing. . J Nanosci & Nanotech 7, 891-897 (2007). Y-H Yun, R Gollapudi, V Shanov, MJ Schulz, Z Dong, A Jazieh, WR Heineman, HB Halsall, D Wong, A Bange, Y Tu, S Subramaniam.
On-line carbon nanotube-based biosensors in microfluidic channels. In: Nanosensors, Microsensors, and Biosensors and Systems 2007, edited by Vijay K. Varadan, Proc. SPIE. 6528, 65280T 1-10, (2007). Y-H Yun, Z Dong, VN Shanov, A Bange, WR Heineman, HB Halsall, L Conforti, A Bhattacharya, MJ Schulz.
Imaging Fluorescently Labeled Microbeads on Polymer Surfaces Using Epifluorescence Microscopy. Nauka Tehnika Bezbednost , 2, 35-40 (2005). I. Nikcevic, A. Bange, E. T. K. Peterson, I. Papautsky, W. R. Heineman, H. B. Halsall, and C. J. Seliskar,.
Electrochemical detection of MS2 phage using a bead-based immunoassay with a nanoIDA. Electroanalysis. A Bange, J Tu, X Zhu, C Ahn, HB Halsall, WR Heineman. Accepted.
124