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Development of Synthetic Conical Nanopores for Protein Sensing Applications

Permanent Link: http://ufdc.ufl.edu/UFE0024356/00001

Material Information

Title: Development of Synthetic Conical Nanopores for Protein Sensing Applications
Physical Description: 1 online resource (149 p.)
Language: english
Creator: Sexton, Lindsay
Publisher: University of Florida
Place of Publication: Gainesville, Fla.
Publication Date: 2009

Subjects

Subjects / Keywords: artificial, conical, nanopore, pi, protein, resistivepulse
Chemistry -- Dissertations, Academic -- UF
Genre: Chemistry thesis, Ph.D.
bibliography   ( marcgt )
theses   ( marcgt )
government publication (state, provincial, terriorial, dependent)   ( marcgt )
born-digital   ( sobekcm )
Electronic Thesis or Dissertation

Notes

Abstract: Abstract of Dissertation Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy DEVELOPMENT OF SYNTHETIC CONICAL NANOPORES FOR PROTEIN SENSING APPLICATIONS By Lindsay Taylor Sexton May 2009 Chair: Charles R. Martin Major: Chemistry The goal of this research is to develop protein sensing devices from artificial conical nanopores. In the first part of this work, single conical nanotubes are used as resistive-pulse sensing devices. A key challenge for this sensing paradigm is building selectivity into the protocol so that the current pulses for the target analyte can be distinguished from current pulses for other species that might be present in the sample. It is demonstrated here that this can be accomplished with a protein analyte by adding to the solution an antibody that selectively binds the protein. Because the complex formed upon binding of the antibody to the protein is larger than the free protein molecule, the current-pulse signature for the complex can be easily distinguished from the free protein. The second part of the research also involves resistive-pulse sensing of protein analytes. Proteins of various sizes were detected with a conical nanotube sensor. The effect of protein size on translocation through a narrow nanotube tip was examined. The size of the protein was found to have a dramatic effect on current-pulse duration. The current-pulse frequency was also affected by the protein size and nanotube tip opening diameter. These studies are important towards the optimization of protein resistive-pulse sensors. In the third part, a new method for optimizing protein resistive-pulse sensing is investigated. Previously, all resistive-pulse sensing work has been done at potentials ? plus or minus1 V. In this work, proteins were sensed at much higher potentials (i.e., up to 4 V). High potential sensing results in a significant decrease in the standard deviation of current-pulse duration for protein analytes. Decreasing the standard deviation in duration allows for better discrimination of analytes, and allowed for two proteins in a mixture to be distinguished. In the last part of this work, a new type of sensor is developed from single conical nanopores. Protein molecules are immobilized on the surface of the nanopore walls and the isoelectric point of the immobilized proteins are determined from current-voltage curves. Isoelectric point determination is made based on the ion current rectification phenomenon exhibited by conical nanopores. At pHs above and below the isoelectric point of the immobilized proteins, the nanopore surface will carry a charge, and therefore current-voltage curves will show ion current rectification. However, at the pH corresponding to the isoelectric point of the immobilized proteins, there will be no surface charge and the current-voltage curves will show no ion current rectification.
General Note: In the series University of Florida Digital Collections.
General Note: Includes vita.
Bibliography: Includes bibliographical references.
Source of Description: Description based on online resource; title from PDF title page.
Source of Description: This bibliographic record is available under the Creative Commons CC0 public domain dedication. The University of Florida Libraries, as creator of this bibliographic record, has waived all rights to it worldwide under copyright law, including all related and neighboring rights, to the extent allowed by law.
Statement of Responsibility: by Lindsay Sexton.
Thesis: Thesis (Ph.D.)--University of Florida, 2009.
Local: Adviser: Martin, Charles R.
Electronic Access: RESTRICTED TO UF STUDENTS, STAFF, FACULTY, AND ON-CAMPUS USE UNTIL 2010-05-31

Record Information

Source Institution: UFRGP
Rights Management: Applicable rights reserved.
Classification: lcc - LD1780 2009
System ID: UFE0024356:00001

Permanent Link: http://ufdc.ufl.edu/UFE0024356/00001

Material Information

Title: Development of Synthetic Conical Nanopores for Protein Sensing Applications
Physical Description: 1 online resource (149 p.)
Language: english
Creator: Sexton, Lindsay
Publisher: University of Florida
Place of Publication: Gainesville, Fla.
Publication Date: 2009

Subjects

Subjects / Keywords: artificial, conical, nanopore, pi, protein, resistivepulse
Chemistry -- Dissertations, Academic -- UF
Genre: Chemistry thesis, Ph.D.
bibliography   ( marcgt )
theses   ( marcgt )
government publication (state, provincial, terriorial, dependent)   ( marcgt )
born-digital   ( sobekcm )
Electronic Thesis or Dissertation

Notes

Abstract: Abstract of Dissertation Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy DEVELOPMENT OF SYNTHETIC CONICAL NANOPORES FOR PROTEIN SENSING APPLICATIONS By Lindsay Taylor Sexton May 2009 Chair: Charles R. Martin Major: Chemistry The goal of this research is to develop protein sensing devices from artificial conical nanopores. In the first part of this work, single conical nanotubes are used as resistive-pulse sensing devices. A key challenge for this sensing paradigm is building selectivity into the protocol so that the current pulses for the target analyte can be distinguished from current pulses for other species that might be present in the sample. It is demonstrated here that this can be accomplished with a protein analyte by adding to the solution an antibody that selectively binds the protein. Because the complex formed upon binding of the antibody to the protein is larger than the free protein molecule, the current-pulse signature for the complex can be easily distinguished from the free protein. The second part of the research also involves resistive-pulse sensing of protein analytes. Proteins of various sizes were detected with a conical nanotube sensor. The effect of protein size on translocation through a narrow nanotube tip was examined. The size of the protein was found to have a dramatic effect on current-pulse duration. The current-pulse frequency was also affected by the protein size and nanotube tip opening diameter. These studies are important towards the optimization of protein resistive-pulse sensors. In the third part, a new method for optimizing protein resistive-pulse sensing is investigated. Previously, all resistive-pulse sensing work has been done at potentials ? plus or minus1 V. In this work, proteins were sensed at much higher potentials (i.e., up to 4 V). High potential sensing results in a significant decrease in the standard deviation of current-pulse duration for protein analytes. Decreasing the standard deviation in duration allows for better discrimination of analytes, and allowed for two proteins in a mixture to be distinguished. In the last part of this work, a new type of sensor is developed from single conical nanopores. Protein molecules are immobilized on the surface of the nanopore walls and the isoelectric point of the immobilized proteins are determined from current-voltage curves. Isoelectric point determination is made based on the ion current rectification phenomenon exhibited by conical nanopores. At pHs above and below the isoelectric point of the immobilized proteins, the nanopore surface will carry a charge, and therefore current-voltage curves will show ion current rectification. However, at the pH corresponding to the isoelectric point of the immobilized proteins, there will be no surface charge and the current-voltage curves will show no ion current rectification.
General Note: In the series University of Florida Digital Collections.
General Note: Includes vita.
Bibliography: Includes bibliographical references.
Source of Description: Description based on online resource; title from PDF title page.
Source of Description: This bibliographic record is available under the Creative Commons CC0 public domain dedication. The University of Florida Libraries, as creator of this bibliographic record, has waived all rights to it worldwide under copyright law, including all related and neighboring rights, to the extent allowed by law.
Statement of Responsibility: by Lindsay Sexton.
Thesis: Thesis (Ph.D.)--University of Florida, 2009.
Local: Adviser: Martin, Charles R.
Electronic Access: RESTRICTED TO UF STUDENTS, STAFF, FACULTY, AND ON-CAMPUS USE UNTIL 2010-05-31

Record Information

Source Institution: UFRGP
Rights Management: Applicable rights reserved.
Classification: lcc - LD1780 2009
System ID: UFE0024356:00001


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DEVELOPMENT OF SYNTHETIC CONICAL NANOPORES FOR PROTEIN SENSING APPLICATIONS By LINDSAY TAYLOR SEXTON A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY UNIVERSITY OF FLORIDA 2009 1

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2009 Lindsay Taylor Sexton 2

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To my parents, Billy and Fran Taylor, and my husband, Galen 3

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ACKNOWLEDGMENTS I would like to thank my advisor, Dr. Charles Martin, for his guidance and support during my time at the University of Florida. I am grateful to have had an advisor who allowed for independent thinking and scientific creativity. I also thank Dr. Martin for teaching me how to be a professional scientist. Working in the Martin group has truly been a pleasure. I acknowledge the entire Martin group for their support, assistance, and friendship over the past five years. I would like to thank Drs. Zuzanna Siwy, Lane Baker, Youngseon Choi and Hitomi Mukaibo for their insightful advice and experimental ideas. I thank Kaan Kececi, Lloyd Horne, Gregory Bishop, Stefanie Sherrill, Pu Jin, John Wharton, JaiHai Wang, and Dooho Park for sharing their ideas and offering support on my projects. I would also like to thank my other friends in the Martin group, Otonye Braide, Funda Tongay and Jillian Perry, who have helped make my time here enjoyable. I would like to thank my husband, Galen, for always being by my side, as well as my parents, Billy and Fran Taylor, for their continued love and support. Finally, I would like to thank God for blessing me with loving and caring family and friends, who help me to grow and learn every day. 4

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TABLE OF CONTENTS page ACKNOWLEDGMENTS ...............................................................................................................4 LIST OF TABLES ...........................................................................................................................8 LIST OF FIGURES .........................................................................................................................9 ABSTRACT ...................................................................................................................................12 CHAPTER 1 INTRODUCTION AND BACKGROUND...........................................................................14 Introduction.............................................................................................................................14 Fabrication of Conical Nanopores..........................................................................................15 The Track-Etch Method..................................................................................................15 Formation of Latent Ion Tracks.......................................................................................15 Ion Track Etching............................................................................................................16 Materials..........................................................................................................................19 Conical Nanopore Characterization and Properties................................................................20 Electron Microscopy.......................................................................................................20 Electrochemical Measurements.......................................................................................21 Electric Field Focusing....................................................................................................22 Ion Current Rectification.................................................................................................23 Tailoring Surface Chemistry of Conical Nanopores..............................................................25 Electroless Gold Plating..................................................................................................26 Chemisorption of Thiols/Generating Biocompatible Surfaces........................................27 EDC/Sulfo-NHS chemistry.............................................................................................28 Biological Nanopores.............................................................................................................29 Resistive Pulse Sensing..........................................................................................................30 Other Nanopore-Based Sensing Strategies.............................................................................33 Dissertation Overview............................................................................................................34 2 RESISTIVE PULSE SENSING OF PROTEINS AND PROTEIN/ANTIBODY COMPLEXES USING A CONICAL NANOTUBE SENSOR.............................................43 Introduction.............................................................................................................................43 Experimental...........................................................................................................................44 Materials..........................................................................................................................44 Pore Etching and Nanotube Preparation..........................................................................45 Current-Pulse Measurements...........................................................................................46 Finite Element Simulations.............................................................................................47 Results and Discussion...........................................................................................................48 Steady-State Current and Current-Pulse Data for BSA...................................................48 Effect of Potential on Current-Pulse Frequency..............................................................51 5

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Effect of Tip Diameter on BSA Current-Pulse Frequency and Duration........................51 Current-Pulse Data for SA, SA/anti-BSA-Fab, and BSA/anti-BSA-Fab........................53 Scatter Plot.......................................................................................................................56 Conclusions.............................................................................................................................57 3 RESISTIVE-PULSE SENSING OF PROTEINS USING A CONICAL NANOTUBE SENSOR.................................................................................................................................67 Introduction.............................................................................................................................67 Experimental...........................................................................................................................69 Materials..........................................................................................................................69 Pore Etching and Nanotube Preparation..........................................................................69 Current-Pulse Measurements...........................................................................................70 Results and Discussion...........................................................................................................71 Steady-State Current and Current-Pulse Events for BSA, Phosphorylase B, and -Galactosidase...............................................................................................................71 Current-Pulse Frequency.................................................................................................73 Scatter Plot and Histograms............................................................................................77 Conclusions.............................................................................................................................79 4 RESISTIVE-PULSE SENSING OF PROTEINS AT HIGH POTENTIALS USING A CONICAL NANOPORE SENSOR.......................................................................................90 Introduction.............................................................................................................................90 Experimental...........................................................................................................................91 Materials..........................................................................................................................91 Pore Etching and Nanotube Preparation..........................................................................92 Current-Pulse Measurements...........................................................................................92 Results and Discussion...........................................................................................................93 Steady-State Current and Current-Pulse Events for BSA and -Galactosidase..............93 Effect of Potential on Protein Translocation...................................................................96 Current-Pulse Events for a Mixture of BSA and -Galactosidase..................................98 Conclusions.............................................................................................................................99 5 DETERMINATION OF THE SURFACE ISOELECTRIC POINT OF PROTEINS IMMOBILIZED ON POLYMER NANOPORE MEMBRANES BY CURRENT-VOLTAGE CURVES...........................................................................................................108 Introduction...........................................................................................................................108 Experimental.........................................................................................................................110 Materials........................................................................................................................110 Pore Etching..................................................................................................................110 PET Surface Modification.............................................................................................111 Current-Voltage Measurements and Isoelectric Point Determination...........................112 Isoelectric Focusing.......................................................................................................113 Results and Discussion.........................................................................................................113 Surface Protein Determination......................................................................................113 6

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Determination of Isoelectric Points from Current-Voltage Curves...............................117 PET and Ethanolamine Modified Nanopores.........................................................118 BSA, Phosphorylase B and Amyloglucosidase Nanopores Modified via EDC/Sulfo-NHS Chemistry................................................................................119 BSA, Phosphorylase B and Amyloglucosidase Nanopores Modified via Nonspecific Adsorption......................................................................................120 Conclusion............................................................................................................................121 6 CONCLUSION.....................................................................................................................135 LIST OF REFERENCES.............................................................................................................138 BIOGRAPHICAL SKETCH.......................................................................................................149 7

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LIST OF TABLES Table page 2-1 Calculated diffusion coefficient for BSA in the nanotube tip, and simulated electric field strength in the tip, for nanotubes with the indicated tip diameters............................66 2-2 Current-pulse-duration () data for the indicated proteins and protein mixtures...............66 3-1 Current-pulse frequency (f p ) and duration () data for the indicated proteins and nanotube tip diameters.......................................................................................................89 3-2 Calculated diffusion coefficient for the proteins in the nanotube tip, for nanotubes with the indicated tip diameters.........................................................................................89 4-1 Summary of current-pulse duration data obtained with three different bare PET conical nanopore sensors.................................................................................................107 5-1 Percent atomic composition of PET membrane before (bare) and after modification with ethanolamine and BSA............................................................................................134 5-2. Average change in nanopore tip diameter with type of molecule and method of immobilization.................................................................................................................134 5-3 Comparison of protein pI values determined from current-voltage curves, IEF and literature...........................................................................................................................134 8

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LIST OF FIGURES Figure page 1-1 Diagram of the ion track-etching method..........................................................................35 1-2 Diagram of the electrochemical cell used for chemical etching and all electrochemical measurements..........................................................................................35 1-3 Schematic of a conical nanopore in a polymer membrane showing the base diameter and tip diameter..................................................................................................................36 1-4 Plot of current versus time recorded during a first step etch of a conical nanopore..........36 1-5 Track-etching of a conical pore, showing bulk etch rate, v B track etch rate, v T and cone half angle, ...............................................................................................................37 1-6 Chemical structures of polymers typically used for track-etching....................................37 1-7 Scanning electron micrographs of nanopores track-etched in various polymer materials.............................................................................................................................38 1-8 Scanning electron micrograph of conical gold nanocones deposited in a conical nanopore membrane...........................................................................................................38 1-9 A typical currentvoltage curve used to calculate the tip diameter of the conical nanopore.............................................................................................................................39 1-10 Distribution of the electric field across a conical nanopore...............................................39 1-11 Model of ion current rectification......................................................................................40 1-12 Schematic of the Au electroless plating procedure............................................................41 1-13 Reaction mechanisms for EDC chemistry.........................................................................41 1-14 Illustration of the resistive-pulse sensing method..............................................................42 1-15 -Hemolysin protein nanopore embedded in a lipid bilayer support.................................42 2-1 Conical nanotube sensor element.......................................................................................59 2-2 Current-voltage curves used to calculate the diameter of the tip opening after each step of the sensor-fabrication process................................................................................59 2-3 Current-time transients for a PEG-functionalized conical nanotube sensor......................60 2-4 Expanded views of typical current pulses associated with tip-to-base translocation........60 9

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2-5 Effect of exposure to BSA on the current-voltage curves for a PEG-functionalized nanotube.............................................................................................................................61 2-6 BSA current-pulse frequency versus transmembrane potential.........................................61 2-7 BSA current-pulse frequency versus nanotube tip diameter..............................................62 2-8 Histograms of BSA current-pulse-duration data for nanotubes with three different tip diameters............................................................................................................................62 2-9 Finite element simulation data...........................................................................................63 2-10 Histograms of current-pulse-duration data for control solutions.......................................63 2-11 Histograms of current-pulse-duration data........................................................................64 2-12 Current-time transient for a PEG-functionalized conical nanotube sensor.......................64 2-13 Scatter plot of current-pulse magnitude (i) versus current-pulse duration ().................65 3-1 Conical nanotube sensor element.......................................................................................81 3-2 Current-time transients for a PEG-functionalized conical nanotube sensor......................82 3-3 Expanded views of typical current pulses associated with tip-to-base translocation of proteins...............................................................................................................................83 3-4 BSA current-pulse frequency versus BSA concentration..................................................84 3-5 BSA current-pulse frequency versus transmembrane potential.........................................84 3-6 BSA current-pulse frequency versus nanotube tip diameter..............................................85 3-7 Histograms of current-pulse amplitude data for protein solutions.....................................86 3-8 Histograms of current-pulse duration data for protein solutions.......................................87 3-9 Scatter plot of current-pulse magnitude (i) versus current-pulse duration () for protein solutions.................................................................................................................88 4-1 Current-voltage curves of a PEG functionalized nanotube sensors before and after application of 5 V applied transmembrane potential.......................................................100 4-2 Current-time transients for a bare PET conical nanopore sensor with buffer only.........100 4-3 Current-time transients for a bare PET conical nanopore sensor with BSA....................101 4-4 Expanded views of typical current pulses associated with tip-to-base translocation of BSA and -galactosidase.................................................................................................101 10

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4-5 Current-voltage curves of a bare PET conical nanopore sensor taken before and after sensing a 100 nM BSA solution......................................................................................102 4-6 Scatter plot of current-pulse magnitude (i) versus current-pulse duration () for BSA at various applied transmembrane potentials..........................................................102 4-7 Histograms of BSA current-pulse duration data..............................................................103 4-8 Histograms of BSA current-pulse amplitude data...........................................................104 4-9 Histograms of BSA and -galactosidase current-pulse duration data.............................105 4-10 Current-time transient for a bare PET conical nanopore sensor with a solution of BSA and -galactosidase on the tip side of the membrane.............................................105 4-11 Histogram of current-pulse duration for a solution containing a mixture of BSA and -galactosidase.................................................................................................................106 5-1 Schematic of the EDC/sulfo-NHS chemistry..................................................................123 5-2 XPS spectra before and after membrane modification....................................................124 5-3 Current-voltage curves taken in 1 M KCl before and after modification with a protein, and after further modification with an antibody.................................................126 5-4 Plot of rectification ratio versus pH for three PET single nanopore membranes............126 5-5 Current-voltage curves taken after chemical etching of PET and after modification with ethanolamine via EDC/sulfo-NHS chemistry..........................................................127 5-6 Plot of rectification ratio versus pH for three ethanolamine modified single nanopore membranes.......................................................................................................................127 5-7 Current-voltage curves taken in 1 M KCl before and after modification to measure the change in tip diameter................................................................................................128 5-8 Current-voltage curves taken in 10 mM KCl, 10 mM PBS buffer before and after modification with proteins at pH 1.6 and 5......................................................................130 5-9 Plot of rectification ratio versus pH for protein modified nanopores..............................132 5-10 Plot of rectification ratio versus pH for protein modified nanopores..............................133 11

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Abstract of Dissertation Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy DEVELOPMENT OF SYNTHETIC CONICAL NANOPORES FOR PROTEIN SENSING APPLICATIONS By Lindsay Taylor Sexton May 2009 Chair: Charles R. Martin Major: Chemistry The goal of this research is to develop protein sensing devices from artificial conical nanopores. In the first part of this work, single conical nanotubes are used as resistive-pulse sensing devices. A key challenge for this sensing paradigm is building selectivity into the protocol so that the current pulses for the target analyte can be distinguished from current pulses for other species that might be present in the sample. It is demonstrated here that this can be accomplished with a protein analyte by adding to the solution an antibody that selectively binds the protein. Because the complex formed upon binding of the antibody to the protein is larger than the free protein molecule, the current-pulse signature for the complex can be easily distinguished from the free protein. The second part of the research also involves resistive-pulse sensing of protein analytes. Proteins of various sizes were detected with a conical nanotube sensor. The effect of protein size on translocation through a narrow nanotube tip was examined. The size of the protein was found to have a dramatic effect on current-pulse duration. The current-pulse frequency was also affected by the protein size and nanotube tip opening diameter. These studies are important towards the optimization of protein resistive-pulse sensors. 12

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In the third part, a new method for optimizing protein resistive-pulse sensing is investigated. Previously, all resistive-pulse sensing work has been done at potentials V. In this work, proteins were sensed at much higher potentials (i.e., up to 4 V). High potential sensing results in a significant decrease in the standard deviation of current-pulse duration for protein analytes. Decreasing the standard deviation in duration allows for better discrimination of analytes, and allowed for two proteins in a mixture to be distinguished. In the last part of this work, a new type of sensor is developed from single conical nanopores. Protein molecules are immobilized on the surface of the nanopore walls and the isoelectric point of the immobilized proteins are determined from current-voltage curves. Isoelectric point determination is made based on the ion current rectification phenomenon exhibited by conical nanopores. At pHs above and below the isoelectric point of the immobilized proteins, the nanopore surface will carry a charge, and therefore current-voltage curves will show ion current rectification. However, at the pH corresponding to the isoelectric point of the immobilized proteins, there will be no surface charge and the current-voltage curves will show no ion current rectification. 13

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CHAPTER 1 INTRODUCTION AND BACKGROUND Introduction In recent years, there has been a growing interest surrounding nanostructured materials. However, the concept of nanoscience was raised over 40 years ago in Richard Feynmans 1959, visionary lecture, There is plenty of room at the bottom. 1 Nanostructured materials are generally considered to be matter with dimensions between 1-100 nanometers (nm). These materials often have unique properties which have potential applications in a wide variety of sectors, including biotechnology, energy, electronics and environmental monitoring to name a few. 2-17 Nanopores are one type of nanostructured material that have received much attention. This attention stems from the critical roles that biological nanopores play in almost all life processes. 18,19 Biological nanopores are present in the cellular membranes of all living cells. There are a variety of different types of biological nanopores, however, they consist of a membrane-bound protein pore that spans the thickness of the cellular membrane. 19 They are the primary devices by which cellular communication occurs via transport of ions and neutral molecules across the cell membrane. 18,19 Recently there has been a great deal of interest in developing artificial analogues of biological nanopores. Artificial nanopores offer the advantages of increased stability, and the ability to control pore dimensions. Artificial nanopores have been fabricated in various materials, using a variety of techniques. 20-65 In the Martin group, we have been exploring conically-shaped artificial nanopores in polymer membranes prepared by the track-etch method. 55-65 The conical shape is particularly advantageous for certain applications (vide infra). 66-78 Artificial conical nanopores have been used to study transport properties, 68-76,79-83 for 14

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bioseperations, 84,85 as templates for the deposition of other materials, 86,87 and as sensing devices. 88-94 The research presented here explores sensing-based applications of these conical nanopores. In particular, this research has focused on the development of protein sensing devices. Chapter 1 is divided into seven additional sections providing the background information for the research. The fabrication of conical nanopores along with the properties they exhibit will be discussed. Biological nanopore sensors will also be reviewed, as well as nanopore-based sensing paradigms. Fabrication of Conical Nanopores The Track-Etch Method The track-etch method entails the chemical removal of latent ion tracks, formed during the irradiation of a membrane sample with a high energy, heavy ion beam. This method allows for microand nano-meter sized pores to be prepared with various dimensions and geometries. The track-etch method has been practiced commercially for decades to prepare multi-pore membranes that are used, for example, for filtration applications 95-98 or as templates for the deposition of other materials. 86,99,100 This method has also been used to create single pore membranes that have been implemented as sensing devices. 88-94 Formation of Latent Ion Tracks Membranes prepared by the track-etch method are created by first bombarding the membranes with a beam of high-energy particles (>1 MeV/nucleon) from a nuclear reactor or cyclotron (Figure 1-1A). Every ion that penetrates the membrane creates a linear damage track that spans the entire thickness of the membrane, which is typically 5-10 m (Figure 1-1B). The number of latent damage tracks formed, which corresponds to the pore density after chemical 15

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etching, is determined by the exposure time to the particle beam. Multi-pore membranes with pore densities ranging from 10 5 to 10 9 pores per cm 2 are commercially available. 101 A procedure for preparing single-track membranes was developed at Gesellschaft fuer Schwerionenforschung (GSI) in Darmstadt, Germany. They use a defocused heavy ion beam to irradiate the polymer membrane with a single ion. 57,58 Single-ion irradiation is achieved by placing a shutter in between the ion beam and the membrane and an ion detector behind the membrane. When the ion detector registers that a single ion has traversed the membrane the shutter is closed, thus precluding any further exposure of the membrane to the beam. The efficiency of this irradiation process depends on the type and energy of the irradiating ions, the radiation sensitivity of the material and the storage conditions of the membrane after irradiation. 67 Ion Track Etching After irradiation, the latent damage tracks can then be chemical etched to create pores (Figure 1-1C). In the commercial process, the ion-tracked membrane is simply immersed into the etching solution, and the damage tracks are etched from both faces of the membrane (isotropic etching). This yields cylindrical pores through the membrane; the pore diameter is determined by the type of material, concentration of etchant, etching time and etchant temperature. For reasons that will be discussed in detail below, conically shaped nanopores have many interesting characteristics that make them well suited for certain applications, such as sensing. The etching process for preparing such conically-shaped nanopores was first developed by Apel et al. 59 In this process, the ion-tracked membrane is first placed between two halves of an electrochemical cell shown in Figure 1-2. 59 An etching solution is added to one side of the cell and a stopping solution is added to the other side. The latent damage track is preferentially 16

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etched from the face of the membrane in contact with the etching solution. When the etchant breaks through to the other side of the membrane the stopping solution neutralizes the etchant. The etching process is stopped by placing the nanopore membrane in water or the stopping solution. The resulting nanopore is conical in shape with the large-diameter (base) opening facing the etch solution and the small-diameter (tip) opening facing the stopping solution (Figure 1-3). During the etching process Pt electrodes are placed on either side of the membrane and a potential is applied. The current across the cell is monitored; initially the current is zero, however, breakthrough is marked by a sudden increase in the ionic current (Figure 1-4). The electrodes are arranged so that the anode is in half-cell containing the etch solution. After breakthrough, an electro-stopping process occurs. 59 To understand this process, consider the case of poly(ethylene terephthalate) (PET). The etching solution used for PET is 9 M NaOH and the stopping solution is 1M formic acid and 1 M KCl. Placing the anode in the etch solution causes the OH etchant to be electrophoretically driven away from the nanopore tip opening electro-stopping. This results in conical nanopores with very small tip diameters (<5 nm). 59 A key issue in artificial nanopore sensor design is reproducible fabrication of the nanopore sensor element. This has proven to be a challenge with artificial nanopore fabrication techniques. 28,32-35,102,103 The anisotropic chemical etching of conical nanopores gives good reproducibility and control in the base diameter of conical track-etched nanopores. 63-65 Reproducibility of the base diameter is achieved by stopping the first etch step after a predetermined amount of time. However, it has been found that if the track-etching process is stopped after a predetermined amount of time, the tip diameter will vary greatly from sample to sample. This is believed to be due to the nature of the anisotropic etching process. The mixing 17

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of etching and stopping solutions in the nanopore tip makes it difficult to control the etch rate in this region. 63 Recently, however, a second isotropic etching step was developed to fine tune and accurately tailor the nanopore tip opening. 63 The second isotropic etch step involves an analogous setup to anisotropic etching, except that a more dilute etchant is placed on both sides of the membrane. This isotropic chemical etching process allows for the entire length of the conical nanopore to be uniformly etched at a slower, more controllable, rate. A transmembrane potential is again applied with platinum wire electrodes, and the current is monitored as a function of time. Instead of stopping the second etch at a predetermined time, the process is stopped at a predetermined current value. If the second etch is stopped after a certain amount of time, the variability in tip diameter, resulting from the first etch step, is retained. However, it has been demonstrated that the current flowing through the nanopore can be correlated to the tip opening diameter. 63 Therefore, by stopping the isotropic etching process at a pre-determined current value, accurate and highly reproducible tip diameters are obtained. The two step etching procedure allows for the base diameter of the conical pores to be fixed in the first, anisotropic, etch step and the tip diameter in the second, isotropic, etch step. 63 The quality of the track-etch process is distinguished by the track-etch-ratio. 56,60 The track-etch-ratio is defined as the ratio of the track etch rate, v T to the bulk etch rate, v B The parameter v B is influenced by the concentration of the etchant, and the etchant temperature. 60 While the parameter, v T is influenced by the sensitivity of the material to tracking, post-irradiation conditions, as well as the etching conditions. 60 The parameters v B and v T determine the shape of track-etch pores as well (Figure 1-5). For high track-etch ratio, the cone half-angle, 18

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, is defined as the ratio v B /v T For materials with a high track-etch velocity, the etch cone becomes cylindrical. Materials A variety of membrane materials are well-suited for the track-etch technique. However, polymer membranes have seen the greatest use due to their chemical and mechanical robustness and high susceptibility to selective ion-track etching. 67 A number of polymer materials are suitable for preparing ion track-etched conical nanopores, including poly(carbonate) (PC), poly(ethylene terephthalate) (PET), poly(propylene) (PP), poly(vinylidenefluoride) (PVDF), and poly(imide) (PI). The chemical structure of three commonly used polymers are shown in Figure 1-6, along with scanning electron micrograph (SEM) images of pores etched in these materials in Figure 1-7. The ideal etching parameters, such as etchant composition, and etching temperature differ for each material. For example, ion tracks in PET membranes are typically etched with a 9 M NaOH etchant and a 1 M formic acid plus 1 M potassium chloride (KCl) stopping solution, at room temperature. 59,62 Upon nanopore breakthrough, hydroxide etchant is simply neutralized by the formic acid. The NaOH hydrolyzes the ester bonds in PET resulting in the formation of carboxylate and hydroxyl groups inside the pore. 83 In contrast, ion tracks in Kapton membranes are etched with a NaOCl etchant with an active chlorine content of 13%, and a 1 M potassium iodide (KI) stopping solution, at a temperature of 50 o C. 60,62,69 Upon etchant breakthrough at the nanopore tip opening, an oxidation-reduction reaction occurs, whereby iodide ions catalyze the reduction of hypochlorite ions to produce chloride ions. This stop-etch reaction yields iodine, yellow in color, which 19

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provides a colorimetric indicator of breakthrough. Etching of Kapton results in the formation of carboxylate groups inside the pore via hydrolysis of imide bonds. 69,104 Etching rates also differ from one material to the other. The two etching properties that have the greatest influence on the shape of conical nanopores are the bulk etch rate, v B and the track etch rate, v T of the material. The bulk etch rate for PET has been determined to be ~2.17 nm/min. 59,62 The track etch rate is ~10 m/hour for 12 m thick PET. 59 Kapton has a bulk etch rate of 0.42.04 m/hour and a track etch rate of 3.12 + 0.65 m/hour for 12 m thick Kapton. 69 For a conically shaped pore, the ratio of v B /v T determines the cone angle of the pores. Kapton has a much higher v B /v T ratio than PET, which results in conical pores with much larger base diameters and cone angles. 59,60,62,69 Conical Nanopore Characterization and Properties Electron Microscopy The three-dimensional shape of conical nanopores is generally characterized from field-emission scanning electron microscopy (FE-SEM) images of gold nanopore replicas (Figure 1-8). The gold replicas are obtained by electrolessly depositing gold inside the empty nanopores. The electroless deposition method (vide infra) also leaves a layer of gold along both faces of the nanopore membrane surface. The nanopore replicas can be liberated by removing one or both layers of gold from the membrane surface and then dissolving away the membrane material. The base diameters, d b of conical nanopores obtained after the first etch step have also been characterized and measured with FE-SEM. Electron micrographs of the base openings of conical pores (Figure 1-7) in track-etched multi-pore membranes can be used to determine the bulk etch rate, v B of the material being etched. The bulk etch rate of the material determines the diameter of the base opening in conical nanopores. If the bulk etch rate is known, then the base diameter of single conical nanopores can be calculated by multiplying v B by the total etching 20

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time of the first step etch. For example, the bulk etch rate of PET was determined to be 2.17 nm/min from FE-SEM images. 59,63 If the anisotropic etching process is stopped after 2 hours then the base diameter of a conical nanopore in PET will be ~520 nm. Electrochemical Measurements Current-voltage curves are typically used to determine the tip opening diameter, d t of single conical track-etched nanopores. This electrochemical method for tip size determination entails mounting the membrane containing the conical nanopore in the same cell setup used for the etching process (Figure 1-2). An electrolyte solution, of known ionic conductivity, is introduced into both sides of the cell along with electrodes. A current-voltage curve for the electrolyte-filled nanopore is then obtained via a linear scan of the transmembrane potential, while measuring the resulting ion current flowing through the nanopore (Figure 1-9). The slope of this current-voltage curve is equal to the ionic conductance, G (in Siemens, S) of the electrolyte-filled nanopore. Since conical nanopores with small tip diameters rectify ion current (vide infra), the linear portion of the current-voltage curve (between -200 and +200 mV) is used to calculate the nanopore tip diameter. 89 The equation for the ionic conductance of a conical pore is, 59,63,88,89 L G4ddtb (1-1) where is the specific conductivity of the electrolyte solution (S cm -1 ), L is the length of the nanopore (thickness of the membrane), d b is the experimentally determined base opening diameter, and d t is the diameter of the tip opening. Since all other parameters except for d t in Equation 1-1 are known, d t can be calculated. Post-anisotropic etching tip opening diameters typically range from 1 nm. 63 It is important to note that this equation can only be rigorously applied to conical nanopores after the first etching step. 63 21

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In order to determine the tip diameter after the second, isotropic, etch step one must take into account the change in both base and tip diameters that occur during the second etch step. A detailed mathematical model has been developed that takes into account this change, and allows one to calculate the single conical nanopore tip diameter from current voltage curves after the second etch. 63 However, this model predicts that for final tip diameters under ~50 nm the change in the base diameter is negligible compared to the change in the tip diameter. 63 This allows for Equation 1-1 to be used to calculate tip diameters after the second step etch for conical nanopores with tip diameters less than 50 nm. Electric Field Focusing An important feature of the conical nanopore sensor is that the voltage drop caused by the ion current flowing through the nanopore is focused at the nanopore tip. 66,92 Indeed, calculations done by Lee et al. indicate that even when a modest transmembrane potential is applied across an electrolyte-filled nanopore, the electric field strength in the nanopore tip is enormous. 66 For example, with an applied transmembrane potential of 1 V and a conical pore with geometry: tip diameter = 60 nm, base diameter = 2.5 mm, length = 6 mmthe magnitude of the electric field at the nanopore tip was modeled to be on the order of 1.5 MV per m (Figure 1-10). 66 Furthermore, finite element simulations of a nanopore at an applied transmembrane potential of 1 V, with a base opening diameter of 520 nm, a length of 12 mm and tip opening diameters ranging from 10 nm, have been done. 92 These studies have shown that the electric field strength inside the nanopore tip increases with decreasing tip opening diameter. A consequence of this focusing effect is that the ion current is extremely sensitive to analyte species present in or near the nanopore tip. That is, there is an analyte sensing zone just inside the tip. 66,88,89 This focusing effect makes conically shaped nanopores better suited for sensing applications than cylindrical nanopores. 22

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Because of this electric-field focusing effect it is not the total length of the pore (membrane thickness) that is relevant to the sensing application but rather the effective length. This effective length is the distance from the tip opening where the majority of the electric field is dropped, put another way, the length of the analyte-sensing zone. If we use the criterion that the effective length is that length over which 80% of the voltage is dropped, then conical nanopores with bases of 5 m and tips of 20 nm have an effective length, as determined by the finite-element method, 66 of 50 nm. Furthermore, such simulations show that the effective length can be controlled by varying the cone angle of the conical nanopore. 66 This is important because it allows for the length of the sensing zone to be tailored to the size of the analyte species to be detected. Ion Current Rectification Another important characteristic exhibited by conical nanopores is the rectification of ion current. Rectification is a term that comes from electronics, referring to devices that conduct electrons only in one direction. 67 Ion current rectifiers show nonlinear current-voltage curves, and have a preferential direction of ion flow through their channels. 18,19 Single conical nanopores in polymer membranes show both ion selectivity, with a preferential direction of cation flow from the tip opening of the pore to the base, and exhibit nonlinear current-voltage curves. 70-73 Several models have been developed to explain ion current rectification in artificial nanopore systems. In the model developed by Siwy et. al. 72 there are three requirements for ion current rectification. These requirements are (i) an asymmetric pore shape, (ii) a tip diameter with dimension comparable to Debye layer at the nanopore wall and (iii) a surface charge on the pore wall. Artificial conical nanopores in polymer membranes possess all of these characteristics, which allows for ion current rectification to occur. 23

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The chemical etching of the ion tracks in PET and Kapton membranes leaves carboxylate groups on the nanopore surface. 62 These carboxylate groups are deprotonated at pHs above the polymers isoelectric point (~3 for both polymers), 62 which results in a negative surface charge. Therefore, current-voltage curves taken with single conical nanopores show ion current rectification at pHs above the polymers isoelectric points (i.e., when negative surface charge is present). 62,69-72 It is important to note that ion current rectification is only observed in conical nanopores when the tip diameter opening is small. This is because in order for rectification to occur the thickness of the Debye layer at the nanopore wall must be of comparable dimensions to that of the pore tip. The Debye layer forms in solution close to the pore surface in order to compensate for the negative surface charge. As long as the thickness of this layer is comparable to the tip radius, the negatively charged conical nanopore will preferentially transport cations and reject anions. 72 The asymmetric shape of conical nanopores has also been proven necessary in order to observe rectification. Studies on the transport properties of cylindrical pores have been conducted, 105 which revealed that track-etched cylindrical nanopores of the same limiting diameter do not rectify ion current. The model proposed by Siwy et. al. is then based on an electrostatic ratchet in which the asymmetry in negatively charged nanopores creates an electrostatic trap for cations at positive applied potentials. 72 The electrostatic trap effectively forms the off state for the artificial nanopores. At negative applied potentials the electrostatic trap is eliminated and the on state is observed. For nanopores with a positively charged surface the ratchet model will be reversed. Another model, introduced by White et. al., 78 also requires an asymmetric pore shape and a pore wall surface charge. In this model, ions of charge opposite to that of the pore walls are able to traverse the pore with much greater ease than those of charge similar to that of the pore walls. 24

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For example, when a transmembrane potential is applied so that ions of charge opposite to that of the pore walls are driven through the conical pore from tip to base, ions of charge similar to that of the pore walls will be driven towards the tip from the base side (Figure 1-11A). However, since these ions have the same charge sign as the pore surface, they will be less likely to exit the pore through the tip (depending on the size of the tip and the surface charge density) due to electrostatic interactions. This is expected to result in a local buildup of ions within the nanopore. 78 Therefore, in this case, the nanopore is highly conductive, and large ionic currents result (Figure 1-11B). When a transmembrane potential is applied so that ions of charge opposite to that of the pore walls are driven through the conical pore from base to tip, ions of charge similar to that of the pore walls will be driven towards the base from the tip side (Figure 1-11C). However, again electrostatic interactions will prevent these ions of like charge from entering the tip of the pore. Thus, a local depletion of ions within the nanopore will result. 78 This leads to a less conductive state and, consequently, smaller ionic currents (Figure 1-11D). Tailoring Surface Chemistry of Conical Nanopores A key function of biological nanopores is their ability to selectively detect and transport specific analytes. It is likewise important to be able to control the surface chemistry of artificial nanopores in order to create both biocompatible and analyte selective devices. There are several strategies currently used to control artificial nanopore surface chemistry. One method employs the electroless deposition of gold onto the nanopore walls followed by modification with simple thiol-based chemistry. Another method takes advantage of surface carboxylate groups produced during chemical etching to attach amine terminated molecules using 1-ethyl-3-[3-dimethylaminopropyl]carbodiimide hydrochloride/N-hydroxysulfosuccinimide (EDC/sulfo-NHS) chemistry. 25

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Electroless Gold Plating The electroless gold deposition method is a form of template synthesis. The template synthesis method 86,99,100,106-125 for the preparation of nanomaterials was pioneered by the Martin group, and involves depositing a desired material into the nanopores of a solid host. This method is useful for preparing hollow nanotubes and solid nanowires. In the electroless deposition of metals, 126 a reducing agent is used to plate metals from solution onto a solid surface. A catalyst is needed to accelerate the rate of the plating reaction on the surface. The procedure that is used to deposit Au along the walls of conical polymer nanopores first involves sensitizing the membrane with Sn(II). This is done by immersing the nanopore membranes in methanol for five minutes and then into a 50/50 water/methanol solution that is 0.026M in SnCl 2 and 0.07M in triflouroacetic acid for 45 min. The Sn(II) sensitizer will bind to the pore walls and membrane surface via electrostatic complexation with the negatively-charged surface functional groups of the polymer formed during chemical etching. 126 The membrane is then washed again for 5 min. in methanol and placed in an aqueous ammoniac solution that is 0.029M in AgNO 3 for 7.5 min. During this step a surface redox reaction occurs and the surface bound Sn(II) is oxidized to Sn(IV) and Ag(I) is reduced to elemental Ag. This deposits silver nanoparticles on the pore walls. 126 The membrane is again washed in methanol for 5 min. before being placed in a gold-plating bath that is 7.9x10 -3 M in Na 3 Au(SO 3 ) 2 0.127 M Na 2 SO 3 0.025M in NaHCO 3 and 0.625 M in formaldehyde at 4 o C. The pH of this solution is first adjusted to 10 with 1 M H 2 SO 4 by dropwise addition. This pH, as well as the lower temperature, are necessary for uniform plating. While in this gold-plating bath another surface redox reaction occurs. Since the standard reduction potential of gold is more positive than that of silver, gold galvanically displaces the silver to yield gold nanoparticles on the pore surface. These nanoscopic gold 26

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particles catalyze the subsequent reduction of Au(I) to Au(0) using formaldehyde as the reducing agent. 126 2 Au(I) + HCHO + 3OH HCOO + 2H 2 O + 2 Au(0) (1-2) This process occurs spontaneously and produces elemental gold via redox chemistry without using electrodes, hence the name electroless. Electroless deposition yields conical gold nanotubes lining the pore walls as well as gold surface layers that cover both faces of the polymer membrane. These surface layers are typically too thin to block the openings of the conical gold nanotube at the membrane surfaces. The surface layers can be removed via a tape-peel method or by swabbing the surfaces with an ethanol-wetted swab. A schematic of the plating process is shown in Figure 1-12. The thickness of the walls of the gold nanotube can be varied by varying the gold plating time, and this provides another means for controlling the diameter of the tip opening of the nanotubes. Gold nanotubes with tip diameters of molecular dimensions (1 nm) can be obtained. Plating for longer periods of time will result in the nanopores being completely filled with gold to yield solid gold nanocones. As previously mentioned the resulting nanocones can be liberated from the membrane and imaged via SEM. Chemisorption of Thiols/Generating Biocompatible Surfaces If conical nanopores in polymer membranes are to be used for biological samples, a means for preventing non-specific protein adsorption to the pore walls must be developed. This can be accomplished by chemisorbing a poly(ethylene glycol) (PEG) thiol to the gold nanotubes. 92,127-130 Typically, a commercially-available, thiolated-PEG is used for this purpose. 92,127 PEG surfaces are ideal for preventing non-specific adsorption because they are hydrophilic and non-ionic. The process of chemisorption simply entails immersing the membrane containing the gold nanotube in a PEG-thiol solution for several hours. The PEG-functionalized gold nanotube is then 27

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removed and rinsed with high purity water. The tip opening diameter after functionalization can be measured electrochemically, as previously described. After chemical functionalization, the conical nanotube can be used for applications which utilize biomolecules, in particular, proteins. Other thiolated molecules have also been attached to the gold coated nanotube walls. For example, this approach has been used to attach biochemical molecular-recognition agents to conical nanotube walls. This allowed for the sensors to be selective to, for example, proteins that bound to these agents. 93 EDC/Sulfo-NHS chemistry EDC/sulfo-NHS chemistry is used to couple carboxylate groups to primary amines (Figure 1-13). The EDC/Sulfo-NHS procedure is commonly used for protein conjugation and immobilization of proteins to a surface. 131-133 Many of the polymers used for preparing track-etched nanopores have carboxylate groups present after etching. Therefore, the EDC/sulfo-NHS method can be used to attached proteins and other molecules with amine terminal groups to the nanopore walls. 90,134-136 EDC can be used alone to couple carboxylate and primary amine groups. EDC will react with a carboxylate group to form an amine-reactive O-acylisourea intermediate. If the intermediate encounters a primary amine group the two molecules will be joined by a stable amide bond. However, the O-acylisourea intermediate is unstable, short lived and the reaction of the intermediate with an amine does not occur quickly. Therefore EDC alone is not very efficient in coupling carboxylate and amine groups. In the presence of water the O-acylisourea intermediate normally hydrolyzes and the carboxyl group is regenerated. For this reason sulfo-NHS is often used to increase the efficiency of EDC-mediated coupling. The addition of sulfo-NHS stabilizes the amine-reactive intermediate by converting it to an amine-reactive sulfo-NHS 28

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ester. The sulfo-NHS ester intermediate has sufficient stability to allow for the primary amine groups to be coupled with the carboxylate groups again via a stable amide bond. Biological Nanopores The growing interest surrounding nanopores arises impart because of the critical roles biological nanopores play in many physiological processes of living organisms. 18,19 Biological nanopores and nanochannels are present in the cellular membranes of all living cells. These channels are formed by membrane proteins that span the entire thickness of the cell membrane (~4 nm). They are the primary device used by cells to communicate with other cells. 18,19 Intercellular communication occurs through transport of ions and neutral molecules through the channels. 18,19 The transport of ions by way of the protein channels is often a highly selective and controlled process. Controlled transport can be achieved because biological ion channels are often selective for certain ions and open and close through gated mechanisms. Gating of ion channels occurs when they open and close in response to certain stimuli, such as deformations in the cell membrane, the presence of a ligand or some other signaling molecule, or changes in membrane potential. 18,19 For example, with voltage-gated ion channels the opened and closed states are dependent on membrane potential. 137 When ion channels are in the opened or on state, ions are allowed to pass through the channels and when in the closed or off state, ion transport is blocked. Many of the transport properties of these biological ion channels are not well understood. Investigating these properties can often be a challenge with biological systems due to their fragile nature. Constructing artificial nanopore devices with transport properties similar to those of biological ion channels could give new insight into the physical and chemical principles of biological ion channel operation. 67 Already, very recent advances in single artificial nanopore 29

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design are shedding light on the mechanisms by which naturally occurring ion channels function. 66,69-76 Another motivation for studying transport in nanopore membranes comes from the possible implementation of these membranes into single-molecule chemical and biochemical sensing devices. In previous work, it has been shown that single, biological transmembrane protein nanopores embedded in lipid bilayer membranes can function as a single-molecule sensing device using the resistive-pulse sensing method (vide infra). 138-165 Biological nanopores have proven to function as extremely versatile and selective resistive-pulse sensors. Recently, it has been shown that artificial single nanopore systems can also be used as platforms for single-molecule resistive-pulse sensing devices. 20-24,26-54,88-92,102,103,166,167 Currently, there is a large research effort focused towards fabricating single nanopores in synthetic materials. 55-65 Resistive Pulse Sensing The resistive pulse sensing method, 102,103,166-168 which is sometimes referred to as stochastic sensing, 138-152 entails mounting a membrane containing a single nanopore between two halves of an electrochemical cell filled with an electrolyte solution. A transmembrane potential is applied, and the resulting ion current flowing through the electrolyte-filled nanopore is recorded versus time (Figure 1-14A). As an analyte, with dimensions comparable to the nanopore diameter is driven through the pore a momentary block in the ion current is observed, yielding a downward current-pulse (Figure 1-14B). The concentration of the analyte can be determined from the frequency of these current-pulse events and the identity of the analyte is encoded in the magnitude and duration of the current pulse. 102,103,166-168 Current work in the field of resistive-pulse sensing is aimed at the detection and characterization of molecules, ions and biopolymers. 24-94,138-171 Sensing of such molecular-sized analytes is possible if the diameter of the nanopore sensor element is of molecular dimensions. 30

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Nanopores in both biological 138-171 and artificial 24-94 membranes have been used to sense such analytes. In particular, a number of prototype resistive-pulse sensing devices have been developed from biological nanopores. The most commonly used protein nanopore is -hemolysin (Figure 1-15). 138-162 -Hemolysin nanopores, either in their wild state or engineered form, have been used to detect DNA, 140,142,155,157-162 nitroaromatic compounds, 143 metal ions, 144 small organic molecules, 147 anions, 146 proteins, 148,149 polymers 153 and enantiomers of drug molecules. 141 There are two main advantages that are offered from this biological nanopore. The first is that the pore size is reproducible from sample to sample and measurement to measurement. The second advantage offered by the -hemolysin nanopore is the selectivity imparted through engineering of the pore. Bayley and coworkers have performed numerous modifications to the -hemolysin pores through genetic engineering and chemical modification, which has allowed for highly selective sensors to be developed. 138-152 The work that has been done with biological nanopores has been extremely influential in the development of molecular resistive-pulse sensors. This work currently stands as the benchmark by which all other resistive-pulse sensing devices are evaluated. However, it seems unlikely that a practical sensing device will ever be developed from biological nanopores due to the fragility of the lipid bilayer membrane support. 139,167 The planar lipid bilayers cannot endure a wide range of pHs, temperatures, applied transmembrane potentials, or solvents, and they are sensitive to vibrations. 139,167 As a result, there has been a significant amount of research focused towards developing artificial analogues of biological nanopores to be used as resistive-pulse sensors. 24-94 Ideally, an artificial nanopore sensor would have the same sensing capabilities and 31

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offer the same advantages of the biological nanopores, but show chemical and mechanical stability over a wide range of conditions. A variety of techniques have been used to fabricate single, artificial nanopores in synthetic materials. The most widely used methods employ ion or electron beams to create single pores in silicon nitride and silicon oxide membranes. 24-39 These nanopores have been used to study mainly DNA, 27,32-37 but some protein sensing 38,39 work has also been reported. Single pores have also been prepared using soft lithographic techniques, 40-45 and have been used to sense colloidal particles 45 and DNA. 42 Nanopores created with this technique have also been used to directly detect the binding of antigens to antibody-coated colloidal particles. 43 Single carbon nanotubes embedded in an epoxy membrane 46-50,102 have been used as the sensing element for the detection of nanoparticles. 47,49,50 A femtosecond-pulsed laser-based technique has been developed to create single pores in glass, which have been used to examine immune complexes. 52 Base etching of silicon wafers 54 and track-etching of nanopores in polymer membranes 55-65 are other methods that have been used to fabricate single nanopores. The Martin group, along with others, 55-65 have utilized the track-etching method to prepare single, conically-shaped nanopores in polymer membranes. The Martin group works with conical nanopores because the geometry has proven to be especially advantageous for resistive-pulse sensing applications. 88-92 Compared with cylindrical pores of the same limiting diameter; the conical pore has a smaller resistance and can therefore generate higher ion currents for a given voltage. 69 The conical pores are also less susceptible to clogging and allow for single molecule transport. 69,91 As previously discussed, the conical shape causes the electric field through the pore to be focused at the pore tip opening, which creates a highly sensitive detection zone within the tip. 66,92 Conically shaped nanopores in polymer 32

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membranes also exhibit many of the transport properties seen in biological ion channels 62,70,72,73 and can act as mimics of voltage-gated ion channels. 62,74-76 Other Nanopore-Based Sensing Strategies Other sensing strategies have also been used with single conical nanopores etched in polymer membranes. A single conical Au nanotube in a PET membrane was used to design a new type of protein biosensor. 93 The nanotubes were modified with various biochemical molecular recognition agents (MRAs) to detect analytes in solution with an on/off response. Like the resistive-pulse sensing method, this sensing protocol also involves passing an ion current through the single nanotube. However, current-pulse translocation events were not observed in this case. Instead, as the analyte bound to the surface bound MRAs the current flowing through the nanopore was permanently shut off. Blockage of the ion current occurred because the diameter of the analyte was of comparable dimensions to that of the nanotube tip. This sensor has been shown to be a highly sensitive and selective type of biosensor, and it should be possible to modify the Au nanotube surface with a wide range of MRAs to selectively detect a wide variety of analytes. Conical nanopore sensors which make use to the ion current rectification phenomenon have also been developed. 94 These devices consisted of a single conical nanopore in a Kapton membrane, and were used to detect drug molecules. Prior to addition of a cationic drug molecule, the single nanopore exhibited ion current rectification. However, after exposure of the nanopore to the drug molecule the extent ion current rectification began to change. The change in rectification was due to the cationic molecule adsorbing onto the nanopore walls and thus changing the surface charge of the pore. It was found that the magnitude of the change in rectification scaled with the concentration of the drug molecule. 33

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Dissertation Overview The goal of this research is to explore one area were these nanopores show potential use sensing. In particular, the research has been focused on developing protein sensing devices. Chapter 1 has reviewed necessary background information for this dissertation including, track-etching, tailoring the nanopore surface, ion current rectification, biological nanopores, and resistive-pulse sensing. In Chapter 2, a strategy for selectively detecting protein molecules using the resistive-pulse sensing method is introduced. Individual protein molecules were first sensed and the resistive-pulse events examined. An antibody, which selectively bound to the protein molecule, was then introduced into solution. The resulting protein/antibody complex was then sensed and the resistive-pulse events were compared to that of the individual protein molecule. In Chapters 3 and 4, the resistive-pulse sensing of individual protein molecules is further explored. In Chapter 3 three different protein molecules were sensed individually, and the effect of protein size on translocation was examined. The effect of high potential sensing was then examined as a means to optimize resistive-pulse sensing of proteins in Chapter 4. In Chapter 5, a different sensing paradigm was used which makes use of the ion current rectification phenomenon exhibited by single conical nanopores. Protein molecules were immobilized onto the nanopore walls and the changes in rectification after immobilization were examined. From the ion current rectification it was possible to determine the isoelectric point (pI) of the immobilized protein molecules. 34

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Figure 1-1. Diagram of the ion track-etching method. A) Irradiation of a membrane with swift, heavy ions results in B) the formation of latent damage tracks along the ions path. C) Chemical etching selectively removes the damage tracks resulting in the formation of pores. Figure 1-2. Diagram of the electrochemical cell used for chemical etching and all electrochemical measurements. 35

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Figure 1-3. Schematic of a conical nanopore in a polymer membrane showing the base diameter and tip diameter (drawing not to scale, see Figure 1-8). 0204060801001200.00.20.40.60.8 Current (nA)Time (min)Breakthrough Figure 1-4. Plot of current versus time recorded during a first step etch of a conical nanopore. 36

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Figure 1-5. Track-etching of a conical pore, showing bulk etch rate, v B track etch rate, v T and cone half angle, Figure 1-6. Chemical structures of polymers typically used for track-etching. A) poly(carbonate) (PC), B) poly(ethylene terephthalate) (PET) and C) poly(imide) (PI). 37

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Figure 1-7. Scanning electron micrographs of nanopores track-etched in various polymer materials. A) PC, 67 B) PET and C) PI. 79 Figure 1-8. Scanning electron micrograph of conical gold nanocones deposited in a conical nanopore membrane. The nanocones have been liberated from the track-etched polymer membrane. 38

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Figure 1-9. A typical currentvoltage curve used to calculate the tip diameter of the conical nanopore. Figure 1-10. Distribution of the electric field across a conical nanopore. The nanopore used for simulations had a base diameter opening of 2.5 mm, a tip opening diameter of 60 nm and a thickness of 6 mm with 1 V applied across the nanopore in 1 M KCl. White hash marks are added to the section of the nanopore where the majority of the electric field is focused. [adapted from Lee, S.; Zhang, Y.; White, H. S.; Harrell, C. C.; Martin, C. R. Analytical Chemistry 2004, 76, 6108-6115.] 39

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Figure 1-11. Model of ion current rectification. Model proposed by White et. al. 169 (drawing of pore not to scale, see Figure 1-8). 40

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Figure 1-12. Schematic of the Au electroless plating procedure. [adapted from Menon, V. P.; Martin, C. R. Analytical Chemistry 1995, 67, 1920-1928.] Figure 1-13. Reaction mechanisms for EDC chemistry. Formation of a stable amide bond occurs between a carboxylate molecule and a molecule with a terminal primary amine group via EDC/Sulfo-NHS chemistry. [adapted from Pierce Biotechnology, http://www.piercenet.com.] 41

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Figure 1-14. Illustration of the resistive-pulse sensing method (drawing of pore not to scale, see Figure 1-8). Figure 1-15. -Hemolysin protein nanopore embedded in a lipid bilayer support. [adapted from Bayley, H.; Jayasinghe, L. Molecular Membrane Biology 2004, 21, 209-220.] 42

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CHAPTER 2 RESISTIVE PULSE SENSING OF PROTEINS AND PROTEIN/ANTIBODY COMPLEXES USING A CONICAL NANOTUBE SENSOR Introduction There is increasing interest in using nanopores in synthetic 24-31,36-54,88-94,102,166,168,170 or biological 138-149,153-164,171 membranes as resistive-pulse sensors for molecular and macromolecule analytes. The resistive-pulse method, 168 which when applied to such analytes is sometimes called stochastic sensing, 138-149 entails mounting the membrane containing the nanopore between two electrolyte solutions, applying a transmembrane potential difference, and measuring the resulting ion current flowing through the electrolyte-filled nanopore. In simplest terms, when the analyte enters and translocates the nanopore, it transiently blocks the ion current, resulting in a downward current pulse. The current-pulse frequency is proportional to the concentration of the analyte, and the identity of the analyte is encoded in the current-pulse signature, as defined by the average magnitude and duration of the current pulses. 102,103,166-168 A key challenge is building analyte selectivity into the sensor itself or into the sensing protocol. For sensors based on the biological nanopore -hemolysin, this has been accomplished by attaching an analyte-selective molecular-recognition agent (MRA) to the nanopore. 138-152 When the analyte is present, it binds to this MRA, yielding current pulses of duration determined by the chemical kinetics of the analyte/MRA interaction. As a result, the current-pulse duration for the analyte is, in general, longer than those for species that do not bind to the MRA. While we have attached MRAs to artificial nanotubes to make highly selective protein sensors, these devices did not use the resistive-pulse method. 93 Another approach for introducing analyte selectivity entails attaching the MRA to a pore-translocating reporter species instead of to the nanopore itself. 43,155 The reporter is first sent through the nanopore in the absence of the analyte to yield current pulses characteristic of the 43

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free reporter. When the analyte is subsequently added, it binds to the MRA on the reporter. Because the resulting reporter/analyte complex is larger in size, the current-pulse signature changes in a predictable way (e.g., longer duration pulses are observed), and it is this change that signals selective detection of the analyte by the MRA-functionalized reporter. 43,155 This study investigates a related strategy for introducing analyte selectivity to resistive-pulse sensing. 52 While this method also makes use of an analyte-selective MRA, it does not require attachment of the MRA to a reporter species. Instead, the MRA used is of dimensions comparable to those of the analyte to be detected. The analyte is first sent through the sensor to obtain current pulses for the free analyte. The MRA is then added to yield the analyte/MRA complex, which is then sent through the sensor. Because the complex is larger than either the free analyte or the free MRA, longer duration current pulses are observed, which signals selective detection of the analyte. This concept is proven here using bovine serum albumin (BSA) as the analyte and a Fab fragment from an antibody to BSA (anti-BSA-Fab) as the MRA. The sensor element was a poly(ethylene glycol) (PEG)-functionalized 127,172 conical gold nanotube 93 prepared by the track-etch method 59,63 in a poly(ethylene terephthalate) (PET) membrane (Figure 2-1). Experimental Materials The anti-BSA-Fab was obtained from Sigma Aldrich; SDS-PAGE showed it to have a molecular weight (MW) of ~50 kDa. BSA (MW ~66 kDa) was obtained from Sigma Aldrich, as was the control protein streptavidin (SA, MW ~52 kDa). Poly(ethylene terephthalate) (PET) membranes, 12 m thick, which contained a single heavy-ion induced damage track, were obtained from GSI (Darmstadt, Germany). A thiolated poly(ethylene glycol) (PEG-thiol, MW 5 kDa) was obtained from Nektar (Huntsville, AL). All other chemicals were of reagent grade and 44

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used as received. Purified water (obtained by passing house-distilled water through a Barnstead, E-pure water purification system) was used to prepare all solutions. Pore Etching and Nanotube Preparation The same cell was used for etching, electrochemical determination of the dimensions of the pore, and for the resistive-pulse experiments. 63 It is a two-compartment Kel-F cell in which the PET membrane separates the two half-cells. The damage track in the PET membrane was chemically etched into a conically shaped pore using the two-step etching method described in detail previously. 63 Conically shaped nanopores and tubes have two openings the large-diameter (or base) opening at one face of the membrane and the small-diameter (or tip) opening at the opposite face (Figure 2-1A). Previous work has shown that the two-step etching method provides for excellent reproducibility in both the tip and base diameters. 63 The base diameter of the pores used for these studies was 520 nm, as determined by electron microscopy. 63 The diameter of the tip opening was determined using an electrochemical method 59 described in detail in prior work. 88,89 Briefly, the membrane containing the single conical nanopore was mounted in the cell, and an electrolyte solution of measured conductivity was placed on either side of the membrane. For these studies this solution was 1 M KCl, pH 6 with a measured conductivity of 10 S/m. A current-voltage curve was obtained (Figure 2-2), the slope of which is the ionic conductance of the electrolyte-filled nanopore. The conductance is used to calculate the diameter of the tip opening. 59,88,89 The nanopores used for these studies had tip diameters, before deposition of the gold nanotube (vide infra) of 50 nm. An electroless plating method 126 was then used to deposit gold along the pore walls to yield a correspondingly conically shaped gold nanotube within the pore (Figure 2-1). Electroless plating also yields gold surface films covering both faces of the membrane, but these were removed by swabbing the membrane faces with an ethanol-wetted cotton swab. A current45

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voltage curve obtained after plating was used to provide the diameter of the tip opening of the resulting gold nanotube (Figure 2-2). The diameter of the much larger base opening remained essentially unchanged after plating. Figure 2-1B shows an electron micrograph of such a conical gold nanotube. While in the sensing application the nanotube is left embedded in the PET membrane, to obtain this image the membrane was removed and the nanotube collected by filtration. 65 PEG-thiol was attached to the gold surfaces to prevent non-specific protein adsorption. 127,172 This was accomplished by immersing the nanotube membrane into a 0.1 mM solution of the PEG-thiol in purified water at 4 C for ~15 hours. The membrane was then rinsed in purified water, and the diameter of the tip opening was remeasured (Figure 2-2). The tip diameters reported here are the diameters measured after PEG functionalization. Nanotubes with tip diameters between 9 nm and 27 nm were used for these studies. The current-voltage curve for the gold nanotube, before PEG functionalization, shows a nonzero current value at zero applied volts. Because the current is zero at an applied potential of 0 V before gold plating and after functionalization with PEG (Figure 2-2), the nonzero current value for the nanotube is most likely a result of residual capacitive current due to the higher capacitance of the unfunctionalized gold. Current-Pulse Measurements The membrane sample containing the PEG-functionalized conical gold nanotube was mounted in the cell and both half cells were filled with ~3.5 mL of 10 mM phosphate buffer solution (pH = 7.4) that was also 100 mM in KCl. A Ag/AgCl electrode (BAS, West Lafayette, IN) was placed into each half-cell solution and connected to an Axopatch 200B (Molecular Devices Corporation, Union City, CA) patch-clamp amplifier. The Axopatch was used to apply the desired transmembrane potential, and measure the resulting ion current flowing through the 46

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electrolyte-filled nanotube. The current was recorded in the voltage-clamp mode with a low-pass Bessel filter at 2 kHz bandwidth. The signal was digitized using a Digidata 1233A analog-to-digital converter (Molecular Devices Corporation), at a sampling frequency of 10 kHz. Data were recorded and analyzed using pClamp 9.0 software (Molecular Devices Corporation). Unless otherwise stated, the applied transmembrane potential was 1000 mV with polarity such that the Ag/AgCl anode was in the half-cell solution facing the base opening, and the Ag/AgCl cathode in the solution facing the tip opening. Because the pI value of BSA is ~4.8 173 and the pI of the control protein, SA, is ~7.0, 174 both proteins have net negative charge in the pH = 7.4 buffer used here. While the exact pI of the anti-BSA-Fab is unknown, isoelectric focusing of whole polyclonal anti-BSA has shown pI values between 5.5 and 7.2. 175 All proteins were added to the half-cell solution facing the tip opening. Finite Element Simulations COMSOL Multiphysics v. 3.3a software (COMSOL, Inc.) was used to compute the electric field strength in and near the tip of the electrolyte-filled nanopore sensor. The software was run on a Dell OptiPlex GX520 (Pentium D CPU, 3.2 GHz, 2 GB RAM). Simulations of this type have been previously described. 66 COMSOL Multiphysics uses the finite element method to solve the partial differential equations that govern a system. Laplaces equation, 2 = 0, was solved for the electrostatic potential, The simulation included an electrolyte layer of 600 m thickness on either side of the membrane with the electrolyte-filled conical nanotube between. The tube was assumed to be 12 m long (the membrane thickness), with a base diameter of 520 nm. The tip diameter was varied from 10 to 25 nm. The tube was divided in two along its long axis (axis of symmetry), and the simulation was done for only one of the halves. Simulating half of the tube allowed us to use a 47

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larger number of elements, which improved accuracy. The number of elements used to compute each result was between 150,000 and 165,000. Results and Discussion Steady-State Current and Current-Pulse Data for BSA In the absence of protein, a steady-state ion current (no current-pulse events) of ~820 pA was observed for the PEG-functionalized nanotube with tip diameter of 17 nm (Figure 2-3A). As will be shown by the simulations (vide infra), conically shaped nanopores and tubes have an analyte-detection zone just inside the tip opening. 66,88,89 When BSA is added to the solution facing the tip opening, current pulses associated with electrophoretic transport of BSA through the detection zone were observed (Figure 2-3B). While the concentration dependence has not yet been studied in detail, as would be expected, 88,89,147,148,155,176 the current-pulse frequency is higher for higher BSA concentrations (Figure 2-3C). That these current pulses are due to electrophoretic transport is supported by the fact that when the polarity is reversed, no current pulses are observed (Figure 2-3D). This is because with this polarity, BSA is driven electrophoretically away from the nanotube membrane. As will be discussed below, the steady-state current is higher at reversed polarity (Figure 2-3D vs Figure 2-3A) because after exposure to BSA the nanotube acts as an ion current rectifier. 73-75 Figure 2-4A shows an expanded view of a BSA current pulse. The current drops precipitously at the start and then tails upward with time. The duration of the pulse () is defined as the time interval between the precipitous drop and the time when the current returns to the baseline value. The current-pulse amplitude (i) is the difference in current between the baseline value and the lowest current within the pulse. Analogous current-pulse shapes have been observed for other charged analytes sent from tip-to-base through conically shaped nanotubes 48

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and pores, for example, for the BSA/anti-BSA complex (Figure 2-4B). This shape is to be expected because the analyte is most effective at blocking the current when it is in the tip and least effective at blocking the current when it is far removed from the tip. Hence, the shape of the current pulse in principle could provide information about the length and geometry of the detection zone. Close inspection of the current-time transients in Figure 2-3 reveals an unexpected result: The steady-state current (between pulses) in the presence of 100 nM BSA (1060 pA) is higher than the steady current in the presence of 50 nM BSA (1000 pA), which is higher than the steady current in the absence of BSA (820 pA). When the BSA solution was removed and replaced with buffer, the baseline current decreased to 860 pA but never returned to the lower pre-BSA-exposure value. To eliminate the possibility that the higher current observed in presence of BSA is due to a change in the conductivity of the buffer, the buffer conductivity with and without 100 nM BSA was measured. The conductivity was the same for both solutions. To explore the origins of this interesting effect, current-voltage curves for the PEG-functionalized nanotube before and after exposure to 100 nM BSA were obtained (Figure 2-5). Before exposure, the current-voltage curve is linear, indicating that the PEG-functionalized nanotube is acting as an ohmic resistor, where the relevant resistance is that of the electrolyte-filled nanotube. 105 In the presence of 100 nM BSA, the current at both positive and negative potentials increases, but the increase at negative potentials is much more dramatic (Figure 2-5). Essentially identical results for unfunctionalized conical gold nanotubes before and after exposure to Cl have been previously obtained by the Martin group. 73 Because Cl adsorbs to gold, after exposure the nanotube had fixed negative surface charge, and this charge was balanced by incorporating an equivalent number of cations from the electrolyte into the tube. 49

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These additional mobile cations made the ionic conductivity of the nanotube higher, and this accounted for the dramatic increase in current at negative potentials upon exposure to Cl 73 As per Figure 2-5, a much smaller increase in current was observed at positive potentials. This is because nanotubes with small tip openings and fixed surface charge are ion-current rectifiers, which causes the current at positive potentials to be suppressed. 73 That essentially identical results are observed upon exposure of the PEG-functionalized nanotubes to BSA indicates that, like Cl the anionic BSA becomes attached to the nanotube walls. The most likely mechanism of attachment is via nonspecific adsorption to the underlying gold. While PEG is attached to the nanotube walls to suppress nonspecific adsorption, 127,172 the PEG monolayer consists, at best, of a close-packed array of the large PEG molecules across the gold surface. Because there is only one thiol per PEG molecule, this means that there is bare gold underneath the umbrella of the PEG molecules. Furthermore, it is unlikely that a perfectly close-packed PEG monolayer can be obtained, so BSA can access this bare gold through defects in the PEG layer. This nonspecific adsorption argument is supported by the fact that the initial linear I-V curve cannot be regenerated by rinsing and soaking in pure buffer (Figure 2-5). If BSA simply partitioned into the PEG, we would expect this process to be completely reversible. It is interesting to note, however, that in prior work from the Martin group on PEG-functionalized gold nanotubes, 172 we conducted protein transport experiments over a 5-day period without any evidence for nonspecific adsorption. That a small amount of nonspecific adsorption can be detected by the current-voltage curve measurement (Figure 2-5) shows how sensitive this electrochemical method is to adsorbed surface charge. 50

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Effect of Potential on Current-Pulse Frequency The pulse frequency (f p ) was determined by counting the number of pulses in 5-min windows (e.g., Figure 2-3B) and then averaging the counts from four such windows. In analogy to data obtained for other charged analytes with both artificial 88,89 and biological 157 nanopores, there is a threshold voltage below which BSA current pulses are not observed, and f p increases exponentially with applied potential above this threshold (Figure 2-6). This is because the BSA molecule pays an entropic penalty when it enters a pore with a tip opening of comparable size to the molecule. 88,89,157 Because BSA is charged, this entropic barrier can be overcome by driving the BSA molecule electrophoretically into the nanotube tip. 88,89,157 Effect of Tip Diameter on BSA Current-Pulse Frequency and Duration The BSA molecule is shaped roughly like an American football with a long axis of ~14 nm and a short axis of ~4 nm. 173 When the tip diameter is smaller than the 14 nm long axis, f p is low, but there is a jump in f p for tips with diameters larger than the long axis (Figure 2-7). This again reflects the entropic penalty paid by the molecule when it enters the tip. The penalty is higher for tips with diameters smaller than the 14 nm BSA long axis because the BSA molecule loses a degree of rotational freedom in such very small tips. Figure 2-8 shows a histogram of BSA current-pulse duration () data obtained for nanotubes with three different tip diameters. Each histogram was fitted to a Gaussian distribution (solid curve), 27,142,158-160 which provided the average pulse duration and standard deviation of the average. Tips with diameters of 9, 17, and 27 nm gave pulses with average values of 520 190, 520 110, and 450 290 ms, respectively. A similar independence of on pore diameter was observed for DNA translocation through a synthetic nanopore sensor. 24 51

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A qualitative explanation for this interesting result can be obtained by considering the electrophoretic velocity, V, of a charged analyte molecule, which is given by 177 V = |z|e / 6r (2-1) where z and r are the charge and radius of the analyte, respectively, e is the electronic charge, E is the electric field strength, and is the solution viscosity. The term in the denominator is the molecular friction that opposes transport, 178,179 which is related to the diffusion coefficient (D) by the StokesEinstein equation D = kT / 6r (2-2) Substituting Equation 2-2 into Equation 2-1 gives V = |z|eD / kT (2-3) Taking the reciprocal of both sides and multiplying by the length of the detection zone, l t for the nanotube sensor, provides the following equation for the current pulse duration, : = l t kT / |z|eD (2-4) Equation 2-4 shows that if is independent of tip diameter (Figure 2-7), then the product D must be independent of tip diameter. To test this prediction, values for the diffusion coefficient for the BSA molecule in the nanotube tip, D tip are needed. D tip is less than the diffusion coefficient in bulk solution because the molecular friction term is larger in the tip due to collisions of the BSA with the nanotube walls (i.e., hindered diffusion). 178,179 D tip can be calculated using the Renkin equation 178,179 5395.009.2104.21soltipDD (2-5) where D sol is the diffusion coefficient in bulk solution, 180 and is the ratio of the diameter of the BSA molecule to the diameter of the nanotube tip. The 4 nm short-axis diameter of BSA was 52

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used in these calculations. Table 2-1 shows D tip values for nanotubes with tip diameters in the range of 10-25 nm. As expected, D tip decreases with decreasing tip diameter. The electric field strength in the nanotube tip can be calculated via finite-element simulation. 66 Figure 2-9A shows the results for a nanotube with tip diameter of 17 nm and base diameter of 520 nm. While the applied transmembrane potential was only 1 V, the field strength just inside the tip is nearly 2 MV m -1 Furthermore, Figure 2-9B shows that field strength in the tip increases linearly with the reciprocal of the tip diameter. The product of ED tip is shown in the last column of Table 2-1, and we see, as predicted, that this product is nearly independent of tip diameter. There are, however, two caveats. First, the calculation uses the values of E and D tip just inside the tip. Yet because the nanotube diameter becomes larger from tip to base, the diffusion coefficient (D) will increase, and E will decrease, as the BSA molecule translocates. Interestingly, again we see the offsetting effects of D and E. Second, because E is linearly related to the reciprocal of the tip diameter (Figure 2-9B), and the relationship between D tip and diameter is nonlinear (Equation 2-5), ED tip will be constant only over a limited range of tip diameters. Current-Pulse Data for SA, SA/anti-BSA-Fab, and BSA/anti-BSA-Fab A histogram of data for a solution that was 100 nM in the control protein SA (Figure 2-10A) provided an average current-pulse duration of SA = 470 140 ms. The histogram for a solution that was 100 nM in the anti-BSA-Fab (Figure 2-10B) provided an average current-pulse duration of Fab = 400 110 ms. Because SA is not bound by the anti-BSA-Fab, a solution that contains both of these proteins should show current pulses for the free SA and for the free Fab. However, because the average pulse durations for these two proteins are the same (Table 2-2), the current-pulse data for 53

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the SA/anti-BSA-Fab solution show only one population of current pulses (Figure 2-11A). An average pulse duration of 460 120 ms was obtained, identical to Fab and SA (Table 2-2). The situation for a solution containing both BSA and the anti-BSA-Fab should be different because the Fab binds BSA to yield a complex (vide infra) that is larger than any of the individual proteins. Studies of DNA translocation through biological 155,159 and artificial 24,30 nanopore sensors have shown that both the average current-pulse duration and the standard deviation of the average increase with the size of the DNA. 24,30,155,159 These results suggest that one should see longer duration current pulses, and larger standard deviations, for solutions containing both BSA and the anti-BSA-Fab. The histogram for a solution that was 100 nM in BSA and 270 nM in the anti-BSA-Fab (Figure 2-11B) shows that this is the case, yielding an average pulse duration for this BSA/anti-BSA-Fab mixture of 2200 650 ms (Table 2-2). That addition of anti-BSA-Fab yields a complex that is larger than the free BSA is further verified by the current-pulse frequency data. Prior to addition of the Fab, a nanotube with a 17 nm tip gave f p for the 100 nM BSA solution of 10.5 0.8 min -1 (Figure 2-3C), but after addition of 270 nM Fab f p was 1.2 0.4 min -1 (Figure 2-12). This drop in frequency occurs because the activation energy for entry of the larger complex into the nanotube tip is higher than the activation energy for the free BSA. There is, however, an interesting point that requires further discussion. The current-pulse data summarized in Figure 2-11B were obtained for a solution that was 100 nM in BSA and 270 nM in the anti-BSA-Fab. If the stoichiometry of binding between the BSA and the anti-BSA-Fab were 1:1, the solution should be ~100 nM in the complex BSA 1 /anti-BSA-Fab 1 and ~170 nM in excess anti-BSA-Fab. This would suggest that one should see two populations of current pulses, one with an average duration of 400 ms for the excess anti-BSA-Fab and one with longer pulse 54

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duration for the complex. Furthermore, again assuming 1:1 stoichiometry, there should be ~1.7 times as many of the short-duration pulses. This is not observed experimentally. Indeed, there is no clear evidence for a set of short duration (~400 ms) current pulses in Figure 2-11B. To understand why this is, it is important to point out that BSA is a multivalent antigen; that is, a single BSA molecule can bind more than one anti-BSA. 181-188 The size and composition of the BSA/anti-BSA complexes formed depend on the relative concentrations of BSA and anti-BSA in solution. A variety of methods have been used to study complex size in the presence of excess antibody. For example, quasi-elastic light scattering was used to determine complex size in the presence of three different monoclonal antibodies, which bound to different epitopes on the BSA molecule. 182,183 When all three antibodies were present, the average size of the BSA/anti-BSA complex was larger than when only two were present, showing that BSA can bind up to three anti-BSA molecules. 183 Similar studies were conducted using sucrose density gradient ultracentrifugation to determine complex size. 184 These studies showed that the maximum complex size occurred in solutions containing a large excess of antibody. 184 Studies have also been conducted with polyclonal antibodies and BSA using size-exclusion high performance liquid chromatography. 185 Complex size was investigated in solutions containing up to 9 times antibody excess and 9 times antigen excess. The largest BSA/ anti-BSA complexes formed when a 3-fold excess of antibody was used. 185 Other studies done with whole polyclonal antibodies showed that when BSA was in low to moderate degrees of excess, the predominant complex formed was BSA 1 /anti-BSA 1 186 These studies also found that a small amount of BSA 2 anti-BSA 1 formed with excess BSA. This is to be expected because the whole antibody is bivalent. This would not be possible with the anti-BSA-Fab used here. 55

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The results of these previous investigations suggest that the predominant complex in the solution containing 100 nM BSA and 270 nM anti-BSA-Fab (Figure 2-11B) has stoichiometry BSA 1 /anti-BSA-Fab 3 This explains why current pulses due to the free Fab are not seen in Figure 11B. These prior studies would also suggest that if solutions with ~1:1 stoichiometry were used, the smaller BSA 1 /anti-BSA-Fab 1 complex would predominate, and a shorter average current-pulse duration would be observed. Figure 2-11C shows that this is indeed the case (Table 2-2). Scatter Plot Scatter plots of current pulse amplitude, i, versus current-pulse duration, are often used to summarize resistive-pulse data. 24,26,30,89,140,160,162 The scatter plot for a solution that was 100 nM in BSA before and after adding 270 nM anti-BSA-Fab (Figure 2-13) shows that it is easy to distinguish the pulses for the BSA/anti-BSA-Fab complex from the pulses for the free BSA. These data also reinforce the point that the spread in values for the complex is larger than that for the free BSA. These results prove the major premise of this work, that a unique current-pulse signature can be obtained when an antibody that binds an analyte protein is added to a solution containing this protein. Furthermore, the control studies with SA/anti-BSA-Fab solutions show that a protein that does not bind to the antibody does not yield this unique set of current pulses. However, a perplexing issue arises from the scatter plot: Why does the larger BSA/anti BSA-Fab complex show current pulses of smaller i than the pulses for the free BSA? This is perplexing because one might expect the larger BSA/anti-BSAFab complex to be more effective at blocking the nanotube tip than the smaller free BSA. As a result, a larger i would be anticipated for the complex. This argument assumes, however, that resistive-pulse sensing of a molecule in a nanopore is completely analogous to resistive-pulse sensing of a particle with the well-known Coulter counter. 102 In the Coulter case, it is simply assumed that the particle 56

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displaces a volume element of electrolyte solution in the pore, and, as a result, the resistance of the pore always increases when the particle is present. There have been two recent reports that indicate that this simple pore-blocking analogy is not always applicable to molecular resistive-pulse sensing. 36,170 In both of these studies, DNA analytes produced upward, as opposed to downward current pulses; that is, the resistance of the nanopore decreased when the DNA analyte was in the pore. These results were interpreted by noting that when a highly charged analyte enters the nanopore, it must bring its charge-balancing counterions with it. As result, there is a transient introduction of additional charge carriers when the analyte is in the nanopore, and this accounts for the upward current pulses. 36,170 It is possible that smaller amplitude current pulses are observed for the BSA/anti-BSAFab complex, relative to free BSA, because the increased size (makes amplitude larger) is partially compensated for by an increase in charge (makes amplitude smaller) as the complex translocates the nanopore tip. Another possibility is that the conformation of the protein changes upon binding by the Fab, which makes the complex less effective at blocking the tip. Further studies will be necessary before a definitive conclusion can be reached. Conclusions In this work it was demonstrated that nanotube resistive-pulse sensors can be used to detect protein analytes and that selectivity can be obtained by adding an antibody to the target protein to the analyte solution. It was also shown that the current-pulse signature for the protein/antibody complex can be used to obtain information about the size of this complex. Hence, the resistive-pulse sensor can be used as a tool to study the stoichiometry of binding between an antibody and an antigen 52 or, in general, between a ligand and a receptor. It was also shown that, because the tip diameter of nanotube sensors prepared by the track-etch method can be controlled at will, 63 the size of the tip can be optimized for detection of the desired analyte (Figure 2-7). In addition, 57

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with this technology on has the ability to conveniently measure the size of the tip after each step of the fabrication process (Figure 2-2). The key to obtaining current pulses for the free BSA and for the BSA/anti-BSA-Fab complex is using a nanotube with a tip diameter comparable to the diameter of these species (10-20 nm). Uram et al. have demonstrated a similar protein-sensing concept using much larger pores (575 nm) prepared by a laser-boring method. 52 Because a much larger pore was used, they could only detect very large protein/antibody complexes consisting of 610-17,300 proteins. Such large complexes formed because Uram et al. used the whole antibody as opposed to the Fab fragment used here. 52 The use of gold as the nanotube material is advantageous because gold can be easily functionalized, for example, to suppress nonspecific protein adsorption, as was done here. Furthermore, the sensor can be evaluated after exposure to the analyte to see if irreversible changes in the device have occurred (Figure 2-5). In addition, the abilities to model the nanotube (Figure 2-9A) and to explore how tube geometry influences field strength in the nanotube tip (Figure 2-9B) are important features of this technology. In the Martin group, we believe these results indicate that gold nanotube resistive-pulse sensors prepared by the track-etch method show promise for development into a practical protein sensing devices. Chapter reproduced with permission from Sexton, Lindsay T.; Horne, Lloyd P.; Sherrill, Stefanie A.; Bishop, Gregory W.; Baker, Lane A.; Martin, Charles R. Journal of the American Chemical Society (2007), 129(43), 13144-13152. Copyright 2007 American Chemical Society. 58

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Figure 2-1. Conical nanotube sensor element. A) Schematic of the PEG-functionalized conical gold nanotube sensor element, showing the base-opening and tip-opening diameters used in these studies. Not to scale, see Figure 2-1B. B) Electron micrograph of such a sensor element after removal from the PET membrane. Note that in the sensing experiment, the nanotube is left embedded in the PET membrane, but it was removed here so that it could be imaged. -1.0-0.50.51.0-20-15-10-551015 Potential (V)Current (nA)-1.0-0.50.51.0-20-15-10-551015 Potential (V)Current (nA) Figure 2-2. Current-voltage curves used to calculate the diameter of the tip opening after each step of the sensor-fabrication process. Black : As-prepared conical nanopore in the PET membrane. Red : After deposition of the conical gold nanotube. Blue : After attachment of PEG to the nanotube walls. 59

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Figure 2-3. Current-time transients for a PEG-functionalized conical nanotube sensor. A) Buffer only. B) Buffer plus 50 nM BSA. C) Buffer plus 100 nM BSA. Applied transmembrane potential for A, B, and C was 1000 mV. D) Buffer plus 100 nM BSA at an applied transmembrane potential of -1000 mV. Tip diameter = 17 nm. Figure 2-4. Expanded views of typical current pulses associated with tip-to-base translocation. A) BSA (100 nM) and B) BSA/anti-BSA-Fab ([BSA] = 100 nM, [anti-BSA-Fab] = 270 nM). Tip diameter = 17 nm. Transmembrane potential = 1000 mV. 60

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-1.0-0.50.51.0-5-4-3-2-112 Potential (V)Current (nA)-1.0-0.50.51.0-5-4-3-2-112 Potential (V)Current (nA) Figure 2-5. Effect of exposure to BSA on the current-voltage curves for a PEG-functionalized nanotube. Black : Before exposure, buffer only. Red : Buffer plus 100 nm BSA. Blue : After removing the BSA solution, rinsing extensively, and returning to buffer only. Tip diameter = 27 nm. 40050060070080090010000102030405060 Current-Pulse Frequency (min-1)Applied Potential (mV)40050060070080090010000102030405060 Current-Pulse Frequency (min-1)Applied Potential (mV) Figure 2-6. BSA current-pulse frequency versus transmembrane potential. Tip diameter = 17 nm. [BSA] = 500 nM. Error bars represent standard deviations obtained by averaging the number of pulses in four 5-min windows of the current-pulse data. 61

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051015202502468101214 Current-Pulse Frequency (min-1)NanotubeTip Diameter (nm) Figure 2-7. BSA current-pulse frequency versus nanotube tip diameter. [BSA] = 100 nM. Applied transmembrane potential = 1000 mV. 02004006008001000 1 01020304050 (ms) Counts 02004006008001000 1 01020304050 (ms) Counts Figure 2-8. Histograms of BSA current-pulse-duration data for nanotubes with three different tip diameters. Tip diameters were: dark gray, 9 nm; light gray, 17 nm; hatched, 27 nm. Solid curves are Gaussian fits. [BSA] = 100 nM. Applied transmembrane potential = 1000 mV. 62

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BElectric Field Strength (MV m-1)Inverse Tip Diameter (nm-1)0.040.060.080.101.01.52.02.53.03.5 Tip end of electrolyte-filled nanotube 0.51.01.52.53.52.03.0 Field Strength (megaVm-1)Electrolyte solution in contact with tip A 8.5 nm 150 nm Tip end of electrolyte-filled nanotube 0.51.01.52.53.52.03.0 0.51.01.52.53.52.03.0 Field Strength (megaVm-1)Electrolyte solution in contact with tip A 8.5 nm 150 nm Figure 2-9. Finite element simulation data. A) Finite-element simulation of electric field strength in and near the tip opening for a nanotube with base opening = 520 nm and tip opening = 17 nm. Membrane thickness = 12 m. Applied transmembrane potential = 1000 mV. B) Plot of electric field strength in the nanotube tip obtained from such simulations versus the inverse of the tip diameter. 020040060080010001200140005101520 (ms)CountsA 020040060080010001200140005101520 (ms)CountsA 20040060080010001200051015202530 (ms)CountsB 20040060080010001200051015202530 (ms)CountsB Figure 2-10. Histograms of current-pulse-duration data for control solutions. A) 100 nM SA. B) 100 nM anti-BSA-Fab. Solid curves are Gaussian fits. Applied transmembrane potential = 1000 mV. Tip diameter = 17 nm. 63

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0200400600800100012001400051015202530 A (ms)Counts 0200400600800100012001400051015202530 A (ms)Counts (ms) 050010001500200025003000350040000246810121416 CountsB (ms) 5001000150020002500051015202530 CountsC Figure 2-11. Histograms of current-pulse-duration data. A) 100 nM SA plus 200 nM anti-BSA-Fab. B) 100 nM BSA plus 270 nM anti-BSA-Fab. C) 100 nM BSA plus 90 nM anti-BSA-Fab. Solid curves are Gaussian fits. Applied transmembrane potential = 1000 mV. Tip diameter for A) and B) is 17 nm. Tip diameter for C) is 27 nm. 840 860 880 880 860 20 sec 840 860 880 880 860 840 860 880 840 860 880 880 860 20 sec 20 sec Figure 2-12. Current-time transient for a PEG-functionalized conical nanotube sensor. A solution 100 nM in BSA and 270 nM in anti-BSAFab added to the tip side of the membrane. Tip diameter = 17 nm. Transmembrane potential = 1000 mV. 64

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(ms)050010001500200025003000350020406080100120140160 i (pA) Figure 2-13. Scatter plot of current-pulse magnitude (i) versus current-pulse duration (). 100 nM BSA only (black ) and 100 nM BSA plus 270 nM anti-BSA-Fab (red ). Applied transmembrane potential = 1000 mV. Tip diameter = 17 nm. 65

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Table 2-1. Calculated diffusion coefficient for BSA in the nanotube tip, and simulated electric field strength in the tip, for nanotubes with the indicated tip diameters. See text and Equation 2-5 for details. Tip Diameter (nm) D tip (x 10 -7 cm 2 s -1 ) a (MV m -1 ) b D tip (x 10 -3 cm V s -1 ) 10 0.40 1.7 3.2 5.4 15 0.27 2.8 2.2 6.2 20 0.20 3.6 1.6 5.8 25 0.16 4.1 1.3 5.3 a Calculated using Eq. 2-5, with a value of 6x10 -7 cm 2 s -1 for D sol 180 b Value obtained from finite-element simulations using a base diameter of 520 nm. Table 2-2. Current-pulse-duration () data for the indicated proteins and protein mixtures. Unless otherwise noted, the tip diameter was 17 nm. Protein(s) Concentration (nM) (ms) BSA 100 520 SA 100 470 Anti-BSA-Fab 100 400 SA/Anti-BSA-Fab 100/200 460 BSA/Anti-BSA-Fab 100/270 2200 BSA/Anti-BSA-Fab (27 nm tip) 100/90 1070 66

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CHAPTER 3 RESISTIVE-PULSE SENSING OF PROTEINS USING A CONICAL NANOTUBE SENSOR Introduction There is increasing interest in using nanopores in synthetic 24-31,36-54,88-94,102,166,168,170 or biological 139-149,153-164,171 membranes as resistive-pulse sensors for molecular and macromolecule analytes. The resistive-pulse method, 168 which when applied to such analytes is sometimes called stochastic sensing, 139-149 entails mounting the membrane containing the nanopore between two electrolyte solutions, applying a transmembrane potential difference, and measuring the resulting ion current flowing through the electrolyte-filled nanopore. In simplest terms, when the analyte enters and translocates the nanopore, it transiently blocks the ion current, resulting in a downward current pulse. The current-pulse frequency is proportional to the concentration of the analyte, and the identity of the analyte is encoded in the current-pulse signature, as defined by the average magnitude and duration of the current pulses. 102,103,166-168 The majority of resistive-pulse sensing work with molecular and macromolecular analytes has utilized the biological nanopore, -hemolysin. 138-162 -Hemolysin nanopores have been used to detect a wide variety of analytes including DNA, 140,142,155,157-162 metal ions, 144 small organic molecules, 147 proteins, 148,149 and polymers. 153 Many studies have been conducted in which such nanopores were used to detect and discriminate ssDNAs of varying chain length. 158,159,162 Branton and coworkers have also demonstrated that -hemolysin pores can be used to discriminate between polynucleotides which differ only in sequence. 160 The data obtained with biological nanopores has been extremely influential in the development of resistive-pulse sensors. However, it seems unlikely that a practical sensing device will ever be developed from biological nanopores due to the fragility of the lipid bilayer membrane that houses the nanopore. 139,167 67

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As a result, there has been a significant amount of research focused towards developing artificial analogues of biological nanopores to be used as resistive-pulse sensors. 24-65,88-92 A variety of techniques have been used to fabricate single, artificial nanopores in various synthetic materials. 24-65 Artificial nanopores have been used as resistive-pulse sensors to study mainly DNA, 27,32-37,42 and nanoparticles. 47,49,50 Resistive-pulse sensing of protein analytes using artificial nanopores is an area in this field that is starting to receive increased interest. 38,39,43,52 Some resistive-pulse protein sensing work with artificial pores has been done, for example, to directly detect the binding of antigens to antibody-coated colloidal particles 43 and to examine immune complexes. 52 However, few studies on the resistive-pulse sensing of individual protein molecules have been reported. 38,39,92 The Martin group has been investigating artificial conical nanopores in polymer membranes prepared by the track-etch method. 55-65 Conical nanopores have been used as resistive-pulse sensors to detect small molecules, 88 DNA, 89,90 and recently, proteins. 92 In Chapter 2, it was shown that individual proteins molecules and protein/antibody complexes could be detected using single conical nanopore sensors. 92 Here the study the transport of individual protein molecules through a single conical nanopore track-etched in a PET membrane is continued. In this work, resistive-pulse sensing of three different protein analytes with varying size is demonstrated. The affect of the protein size on transport was examined by comparing the current-pulse signatures and frequencies of each protein. The three proteins used in this work were bovine serum albumin (BSA), phosphorylase B and -galactosidase. The sensor element was a poly(ethylene glycol) (PEG)-functionalized 127,172 conical gold nanotube 93 prepared by the track-etch method 59,63 in a poly(ethylene terephthalate) (PET) membrane (Figure 3-1). 68

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Experimental Materials Bovine serum albumin (BSA, MW ~66 kDa), phosphorylase B (MW ~97.4 kDa) and -galactosidase (MW ~116 kDa) were obtained from Sigma Aldrich. Poly(ethylene terephthalate) (PET) membranes, 12 m thick, which contained a single heavy-ion induced damage track, were obtained from GSI (Darmstadt, Germany). A thiolated poly(ethylene glycol) (PEG-thiol, MW 5 kDa) was obtained from Nektar (Huntsville, AL). All other chemicals were of reagent grade and used as received. Purified water (obtained by passing house-distilled water through a Barnstead, E-pure water purification system) was used to prepare all solutions. Pore Etching and Nanotube Preparation The same cell was used for etching, electrochemical determination of the dimensions of the pore, and for the resistive-pulse experiments. 63 It is a two-compartment Kel-F cell in which the PET membrane separates the two half-cells. The damage track in the PET membrane was chemically etched into a conically shaped pore using the two-step etching method described in detail previously. 63 Conically shaped nanopores and tubes have two openings the large-diameter (or base) opening at one face of the membrane and the small-diameter (or tip) opening at the opposite face (Figure 3-1A). It has been shown that the two-step etching method provides for excellent reproducibility in both the tip and base diameters. 63 The base diameter of the pores used for these studies was 520 nm, as determined by electron microscopy. 63 The diameter of the tip opening was determined using an electrochemical method 59 described in detail in prior work. 88,89 Briefly, the membrane containing the single conical nanopore was mounted in the cell, and an electrolyte solution of measured conductivity was placed on either side of the membrane. For these studies this solution was 1 M KCl, pH 6 with a measured conductivity of 10 S/m. A current-voltage curve was obtained, the slope of which is 69

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the ionic conductance of the electrolyte-filled nanopore. The conductance is used to calculate the diameter of the tip opening. 59,88,89 The nanopores used for these studies had tip diameters, before deposition of the gold nanotube (vide infra), of 50 nm. The PEG-functionalized nanotube sensors used in this work have been previously described and utilized for resistive-pulse sensing of proteins. 92 Briefly, PEG functionalization was achieved by first using an electroless plating method 126 to deposit gold along the pore walls to yield a correspondingly conically shaped gold nanotube within the pore (Figure 3-1). Electroless plating also yields gold surface films covering both faces of the membrane, but these were removed by swabbing the membrane faces with an ethanol-wetted cotton swab. A current-voltage curve obtained after plating was used to provide the diameter of the tip opening of the resulting gold nanotube. The diameter of the much larger base opening remained essentially unchanged after plating. PEG-thiol was then attached to the gold surfaces to prevent non-specific protein adsorption. 127,172 This was accomplished by immersing the nanotube membrane into a 0.1 mM solution of the PEG-thiol in purified water at 4 C for ~15 hours. The membrane was then rinsed in purified water, and the diameter of the tip opening was remeasured. The tip diameters reported here are the diameters measured after PEG functionalization. Nanotubes with tip diameters of 17 and 23 nm were used for these studies. Current-Pulse Measurements The membrane sample containing the PEG-functionalized conical gold nanotube was mounted in the cell and both half cells were filled with ~3.5 mL of 10 mM phosphate buffer solution (pH = 7.4) that was also 100 mM in KCl. A Ag/AgCl electrode (BAS, West Lafayette, IN) was placed into each half-cell solution and connected to an Axopatch 200B (Molecular Devices Corporation, Union City, CA) patch-clamp amplifier. The Axopatch was used to apply 70

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the desired transmembrane potential, and measure the resulting ion current flowing through the electrolyte-filled nanotube. The current was recorded in the voltage-clamp mode with a low-pass Bessel filter at 2 kHz bandwidth. The signal was digitized using a Digidata 1233A analog-to-digital converter (Molecular Devices Corporation), at a sampling frequency of 10 kHz. Data were recorded and analyzed using pClamp 9.0 software (Molecular Devices Corporation). Unless otherwise stated, the applied transmembrane potential was 1000 mV with polarity such that the Ag/AgCl anode was in the half-cell solution facing the base opening, and the Ag/AgCl cathode in the solution facing the tip opening. Because the pI values of BSA, phosphorylase B, and -galactosidase are ~4.8, 173 5.8-6.3, 189,190 and ~4.6, 191 respectively, all the proteins have net negative charge in the pH = 7.4 buffer used here. All proteins were added to the half-cell solution facing the tip opening and driven electrophoretically through the nanotube sensor from tip to base (Figure 1). Results and Discussion Steady-State Current and Current-Pulse Events for BSA, Phosphorylase B, and -Galactosidase In the absence of protein analyte, a steady-state ion current (no current-pulse events) of ~820 pA was observed for the PEG-functionalized nanotube with tip diameter of 17 nm (Figure 3-2A). As previously described, 66,88,89,92 conical nanopores and nanotubes have an analyte-detection zone just inside the tip opening. When a solution 100 nM in BSA was added to the half-cell facing the tip opening, current-pulses associated with the electrophoretic transport of BSA through the detection zone were observed (Figure 3-2B). After sensing BSA the nanotube sensor was thoroughly rinsed and the steady-state ion current (no current-pulse events) returned (Figure 3-2C). Solutions 100 nM in phosphorylase B and -galactosidase were subsequently 71

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sensed with the same nanotube sensor. These protein solutions also showed current-pulse events associated with electrophoretic transport through the detection zone (Figure 3-2D and 3-2E). After rinsing again these current-pulse events ceased, and the steady-state current with no events was restored. That these current pulses are due to electrophoretic transport is supported by the fact that when the polarity is reversed, no current pulses are observed (Figure 3-2F). This is because with this polarity, the protein molecules are driven electrophoretically away from the nanotube membrane. As will be discussed below, the steady-state current is higher (~2850 pA) at reversed polarity (Figure 3-2F vs Figure 3-2A) because after exposure to proteins the nanotube acts as an ion current rectifier. 73-75 Close inspection of the current-time transients in Figure 3-2 reveals that the steady-state current (between pulses) in the presence of 100 nM BSA (~1060 pA) is higher than the steady current in the absence of BSA (~820 pA). When the BSA solution was removed and replaced with buffer, the baseline current decreased to ~860 pA but never returned to the lower pre-BSA-exposure value. In the presence of 100 nM phosphorylase B and 100 nM -galactosidase the steady-state current again rose to ~960 pA and ~910 pA, respectively. When these protein solutions were removed and replaced with buffer, the baseline current decreased again to ~860 pA. The origins of this effect have been described in detail in prior work. 92 Briefly, the increase in steady-state current is a result of protein molecules nonspecifically adsorbing on portions of the gold nanotube walls that remain exposed under the PEG layer. Protein attachment in this way introduces excess negative charge on the nanotube walls. This causes the nanotube to function as an ion-current rectifier. 73-75 This ion current rectification phenomenon 72

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results in a large increase in current at negative potentials (Figure 3-2F) and only a slight increase at positive potentials (Figure 3-2B). An expanded view of a typical current-pulse event for each protein is shown in Figure 3-3. As reported in Chapter 2, the BSA current pulse drops sharply and then tails upward with time. 92 The current-pulse events for phosphorylase B and -galactosidase were found to exhibit similar shapes (Figure 3-3). This shape reflects the fact that the protein is driven into the tip, where it is most effective at blocking the ion current, and driven toward the base, where it becomes less and less effective at blocking the current. 92 The current-pulse events can be characterized by the current-pulse amplitude (i) and the current-pulse duration (). As per the prior work presented in Chapter 2, 92 i is defined as the difference in current between the baseline and the lowest current within a pulse, and is defined as the time interval between the precipitous drop and the time when the current returns to the baseline value. Current-Pulse Frequency The current-pulse frequency (f p ) was determined by counting the number of pulses in five-minute windows (e.g., Figures 3-2B) and then averaging the counts from four such windows. The current-pulse frequency decreases with protein size for both the 17-nm tip sensor and the 23-nm-tip sensor (Table 3-1). In addition, for the two largest proteins, f p is smaller for the 17-nm tip than for the 23-nm tip. The size of the nanotube tip opening as well and the size of the protein analyte greatly affected current-pulse frequency. The reported hydrodynamic diameters of BSA, phosphorylase B and -galactosidase are 6.8 nm, 180 9.8 nm 192 and 16.6 nm, 193 respectively. There are several factors to consider when analyzing the affects of analyte size and nanotube tip opening on current-pulse frequency. The first is the partitioning of the analyte 73

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species into the nanotube sensor element. If we represent the analyte species as A, then this partition process can be written as, A s A tip (3-1) where the subscripts, s and tip, correspond to the analyte in the contacting solution phase and the analyte in the nanotube tip, respectively. At first glance, this seems like a simple process because the analyte is dissolved in an electrolyte and the same electrolyte floods the nanotube. As a result, one might expect that the partition coefficient, = [A tip ]/[A s ], would be unity, and this is true if the radius of the analyte, r A is much less than the radius of the tip opening, r tip If, however, the analyte and the nanotube tip are of comparable radii, the partition coefficient will be less than unity. This can be quantified by defining the parameter, which is given as = r A /r tip (3-2) The partition coefficient is related to via Equation 3-3, 9 = (1-) 2 (3-3) Equation 3-3 was used to calculate the partion coefficients for the three proteins utilized in this work. The results are summarized in Table 3-2 for each nanotube tip diameter. The partition coefficient decreases with increasing protein size for a given tip diameter because the analyte pays an entropic penalty when it partitions into a nanotube tip with a radius comparable to that of the analyte. The entropic penalty becomes larger as the size of the analyte approaches the size of the nanotube tip opening. This diminution in the partition coefficient is deleterious to resistive-pulse sensing because, in general, an analyte cannot be detected unless it partitions into the pore, and as have been shown, the tendency to partition decreases with increasing analyte size. 74

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However, once the protein analyte partitions into the nanotube, it is electrophoretically transported through the pore and produces a current-pulse. The equation for current-pulse frequency has been previously derived from the electrophoretic flux, 89 and is given as, RTArCEzFDftiptipp/)(2 (3-4) were z is the charge of the analyte, F is Faradays constant, D tip is the diffusion coefficient of the analyte in the nanopore tip, C is the analyte concentration, E is the electric field strength inside the nanopore tip, r tip is the radius of the nanopore tip, A is Avogadros number, R is the gas constant, and T is temperature. Equation 3-4 predicts that an increase in current-pulse frequency can be obtained by increasing analyte concentration, applied transmembrane potential and/or nanotube tip diameter. Indeed these effects have been observed experimentally with BSA. Figure 3-4 shows a plot of f p versus BSA concentration for a PEG functionalized nanotube with a 17 nm tip. The frequency of BSA translocation events scales linearly with concentration over the range examined. It was previously shown that f p increases with both increasing applied potential and increasing nanotube tip opening diameter as well. 92 In these instances, however, f p did not increase linearly, as the equation would predict. The BSA current-pulse frequency was found to increase exponentially with increasing applied potential, above the threshold voltage (Figure 3-5). In analogy to data obtained for other charged analytes with both artificial 88,89 and biological 157 nanopores, there is a threshold voltage below which BSA current pulses are not observed, and f p increases with applied potential above this threshold. This threshold is a result of the entropic penalty the BSA molecule must pay when it enters a pore with a tip opening of comparable size to the molecule. 88,89,157 Because BSA is charged, this entropic barrier can be overcome by driving the BSA molecule electrophoretically into the nanotube tip. 88,89,157 75

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An exponential increase in current-pulse frequency with increasing applied potential has also been observed by Henrickson et. al. 157 A detailed model accounting for this exponential increase was described. 194 This model describes different voltage regimes for each analyte where f p will either be very small (i.e., potentials around the threshold voltage), increase exponentially (i.e., potentials right above the threshold voltage), or increase linearly (i.e., potentials well above the threshold voltage). 194 In Chapter 2, it was also reported that the BSA current-pulse frequency jumps at a certain nanotube tip diameter. 92 This jump was related to the size and shape of the BSA molecule. The BSA molecule is shaped roughly like an American football with a long axis of ~14 nm and a short axis of ~4 nm. 173 When the tip diameter is smaller than the 14-nm long axis, f p is low, but there is a jump in f p for tips with diameters larger than the long axis (Figure 3-6). This again reflects the entropic penalty paid by the molecule when it enters the tip. The penalty is higher for tips with diameters smaller than the 14-nm BSA long axis because the BSA molecule loses a degree of rotational freedom in such very small tips. In the case with phosphorylase B and -galactosidase, the current-pulse frequency was also dependent on tip diameter. As the size of the protein increases, and approaches the size of the nanotube tip opening, the entropic penalty that the molecule pays for entering the tip becomes higher. Therefore, a drop in frequency, relative to that of BSA at the same tip diameter, is observed because the activation energy for entry of the larger proteins into the nanotube tip is higher than the activation energy for BSA (Table 3-1). In analogy to the previously discussed results of f p versus nanotube tip diameter with BSA, the pulse frequency of phosphorylase B and -galactosidase increased with tip diameter (Table 3-1), because the entropic penalty to enter the tip was lower. 76

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According to Equation 3-4, another factor that affects f p is the diffusion coefficient. The reported diffusion coefficients of BSA, phosphorylase B and -galactosidase are 6 x 10 -7 180 4.33 x 10 -7 192 and 3.03 x 10 -7 cm 2 s -1 193 respectively. These reported values are the diffusion coefficients in bulk solution (D sol ). The diffusion coefficient values inside the nanotube tip (D tip ) will be less than the diffusion coefficient in bulk solution due to the effects of hindered diffusion inside the nanotube. 178,179 D tip can be calculated using the Renkin equation 178,179 5395.009.2104.21soltipDD (3-5) where is the ratio defined in Equation 3-2. Table 3-2 shows the values of and D tip calculated for each protein in a nanopore with tip diameters of 17 and 23 nm. The percentage of decrease in the diffusion coefficient for BSA, phosphorylase B and -galactosidase is ~47%, ~87%, and ~95%, respectively, for the 17 nm tip and ~36%, ~75%, and ~92%, respectively, for the 23 nm tip. These calculations again reflect on the entropic penalty paid by the proteins as they enter the nanotube tip, 161 as the larger proteins have a greater percent decrease in diffusion coefficient values. Scatter Plot and Histograms In order to obtain average values, and standard deviations, for the current-pulse amplitude and duration, histograms of the i and data were plotted for each protein (Figure 3-7 and Figure 3-8). The average values and standard deviations were obtained by fitting the histograms to a Gaussian distribution (solid curves). 27,142,158-160 As noted in the introduction, one of the key objectives of this study was to investigate how protein size affects the current-pulse signature. The BSA molecule is shaped roughly like an American football with a short axis of ~4 nm and a long axis of ~14 nm. As previously noted, 77

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the hydrodynamic diameter, calculated from the diffusion coefficient, is 6.8 nm. 180 The hydrodynamic diameters of phosphorylase B and -galactosidase are 9.8 nm 192 and 16.6 nm, 193 respectively. Figure 3-7 shows the i histogram for each protein when a nanotube with a tip diameter of 17 nm was used as the sensor. The average i values for BSA, phosphorylase B, and -galactosidase are 80 pA, 95 pA, and 135 pA, respectively. These data show that while i in general increases with protein size, it would be difficult to distinguish between these three proteins on the basis of the current-pulse amplitude alone. Figure 3-8 shows the corresponding histogram for each protein, and the average values are shown in Table 3-1. We see that the average current-pulse duration increases with the size of the protein with the largest protein, -galactosidase, giving an average pulse duration of almost two seconds. However, the standard deviation of the value also increases with the size of the protein molecule. For reasons discussed in Chapter 2, we have also found that is independent of tip diameter (Table 3-1). 92 These trends in the current-pulse amplitude and duration data can be visualized more clearly via scatter plots of i vs. (Figure 3-9). The large spread in the current-pulse duration data for the largest protein is clearly seen, as is the much smaller spread for the smallest protein, BSA. This plot suggests that -galactosidase could be distinguished from BSA because only -galactosidase produces pulses in the upper right quadrant of this plot, and only BSA produces pulses in the lower left quadrant. Distinguishing phosphorylase B from the other two proteins based purely on such current-pulse data would be problematic. However, in Chapter 2 it was shown that protein specificity can be introduced by adding an antibody that selective binds the target protein-analyte to the analyte solution. 92 Hence, one does not have to rely on the current78

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pulse signature of the protein by itself to determine whether a particular analyte protein is present in the sample. These general trends of increasing and standard deviation of with increasing analyte size have been observed with other resistive-pulse sensors. For example, numerous studies have been conducted in which DNA strands of various lengths were transported through both biological 158,159,162 and artificial nanopores. 24,27,30 These studies have shown that as the length of DNA increases so does the average current-pulse duration 24,27,30,158,159,162 as well as the standard deviation in current-pulse duration. 24,27 In a study conducted by Golovchenko and coworkers, the effect of applied potential on DNA current-pulse duration was also investigated. 24 They found that as the applied potential was increased from 60 mV to 120 mV, the average value of decreased, as did the standard deviation in These results suggest that resistive-pulse sensing of protein analytes with artificial conical nanopores could be further optimized by increasing the applied transmembrane potential. Conclusions In this study it was demonstrated that nanotube resistive-pulse sensors can be used to detect protein analytes of various sizes. It was also shown that the current-pulse frequency for the protein can be used to obtain information about the size of the protein. Hence, the resistive-pulse sensor can be used as a tool to study entropic effects resulting from translocation through a narrow nanotube tip. Furthermore, because the tip diameter of nanotube sensors prepared by the track-etch method can be controlled at will, 63 the size of the tip can be optimized for detection of the desired analyte. In addition, with this technology on has the ability to conveniently measure the size of the tip after each step of the fabrication process. 79

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From the scatter plots and histograms of i and it was shown that current-pulse duration could be a useful tool in discriminating proteins based on size. The increase in standard deviation with increasing protein size, however, presents a problem. Methods for further optimization of protein resistive-pulse sensing have been suggested. Future work will involve optimization of the transmembrane potential applied during sensing, as a means for decreasing the standard deviations in current-pulse duration. 80

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Tip Opening (17 to 23 nm) Base Opening (520 nm) PET Membrane 12 m Electrolyte-Filled Conical Gold Nanotube Tip Opening (17 to 23 nm) Base Opening (520 nm) PET Membrane 12 m Electrolyte-Filled Conical Gold Nanotube Figure 3-1. Conical nanotube sensor element. A) Schematic of the PEG-functionalized conical gold nanotube sensor element, showing the base-opening and tip-opening diameters used in these studies. Not to scale, see Figure 3-1B. B) Electron micrograph of such a sensor element after removal from the PET membrane. Note that in the sensing experiment, the nanotube is left embedded in the PET membrane, but it was removed here so that it could be imaged. 81

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5 sec100 pA Current (pA) 30 sec 30 sec 30 sec 5 sec 5 sec 82010608609609102850ABCDEF 5 sec100 pA Current (pA) 30 sec 30 sec 30 sec 5 sec 5 sec 82010608609609102850 5 sec 5 sec 5 sec100 pA Current (pA) 30 sec 30 sec 30 sec 5 sec 5 sec 82010608609609102850100 pA Current (pA) 30 sec 30 sec 30 sec 5 sec 5 sec 82010608609609102850 30 sec 30 sec 30 sec 5 sec 5 sec 30 sec 30 sec 30 sec 5 sec 5 sec 30 sec 30 sec 30 sec 30 sec 30 sec 30 sec 30 sec 30 sec 30 sec 5 sec 5 sec 5 sec 5 sec 5 sec 5 sec 82010608609609102850ABCDEF Figure 3-2. Current-time transients for a PEG-functionalized conical nanotube sensor. A) Buffer only. B) Buffer plus 100 nM BSA. C) Buffer only after sensing 100 nM BSA. D) Buffer plus 100 nM phosphorylase B. E) Buffer plus 100 nM -galactosidase. Applied transmembrane potential for A-E was 1000 mV. F) Buffer plus 100 nM BSA at an applied transmembrane potential of -1000 mV. Tip diameter = 17 nm. 82

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40 ms2 sec ABC 40 ms2 sec 40 ms2 sec 40 ms2 sec ABC Figure 3-3. Expanded views of typical current pulses associated with tip-to-base translocation of proteins. A) BSA (100 nM), B) phosphorylase B (100 nM), and C) -galactosidase (100 nM). Tip diameter = 17 nm. Transmembrane potential = 1000 mV. 83

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Concentration (nM)Current-Pulse Frequency (min-1)01002003004005000102030405060 Figure 3-4. BSA current-pulse frequency versus BSA concentration. Tip diameter = 17 nm. Transmembrane potential = 1000 mV. Error bars represent standard deviations obtained by averaging the number of pulses in four 5-min windows of the current-pulse data. Applied Potential (mV)Current-Pulse Frequency (min-1)40050060070080090010000102030405060 Figure 3-5. BSA current-pulse frequency versus transmembrane potential. Tip diameter = 17 nm. [BSA] = 500 nM. Error bars represent standard deviations obtained by averaging the number of pulses in four 5-min windows of the current-pulse data. 84

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NanotubeTip Diameter (nm)Current-Pulse Frequency (min-1)051015202502468101214 Figure 3-6. BSA current-pulse frequency versus nanotube tip diameter. [BSA] = 100 nM. Applied transmembrane potential = 1000 mV. 85

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204060801001201401600204060 Countsi(pA)A 4060801001200102030 Countsi(pA)B 40608010012014016018020005101520 Countsi(pA)C 204060801001201401600204060 Countsi(pA)A 204060801001201401600204060 Countsi(pA)A 4060801001200102030 Countsi(pA)B 4060801001200102030 Countsi(pA)B 40608010012014016018020005101520 Countsi(pA)C 40608010012014016018020005101520 Countsi(pA)C Figure 3-7. Histograms of current-pulse amplitude data for protein solutions. A) 100 nM BSA, B) 100 nM phosphorylase B, and C) 100 nM -galactosidase. Solid curves are Gaussian fits. Applied transmembrane potential = 1000 mV. Tip diameter = 17 nm. 86

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020040060080010000510152025303540 CountsDuration (ms) Counts (ms)A 0400800120016002000024681012 CountsDuration (ms) Counts (ms)B 0100020003000400002468101214 CountsDuration (ms) Counts (ms)C 020040060080010000510152025303540 CountsDuration (ms) Counts (ms)A 020040060080010000510152025303540 CountsDuration (ms) Counts (ms)A 0400800120016002000024681012 CountsDuration (ms) Counts (ms)B 0400800120016002000024681012 CountsDuration (ms) Counts (ms)B 0100020003000400002468101214 CountsDuration (ms) Counts (ms)C 0100020003000400002468101214 CountsDuration (ms) Counts (ms)C Figure 3-8. Histograms of current-pulse duration data for protein solutions. A) 100 nM BSA, B) 100 nM phosphorylase B, and C) 100 nM -galactosidase. Solid curves are Gaussian fits. Applied transmembrane potential = 1000 mV. Tip diameter = 17 nm. 87

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0100020003000400050100150200 Amplitude (pA)Duration (ms) (ms) i(pA)0100020003000400050100150200 Amplitude (pA)Duration (ms) (ms) i(pA) Figure 3-9. Scatter plot of current-pulse magnitude (i) versus current-pulse duration () for protein solutions. 100 nM BSA (black ), 100 nM phosphorylase B (red ), and 100 nM -galactosidase (blue ). Applied transmembrane potential = 1000 mV. Tip diameter = 17 nm. 88

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Table 3-1. Current-pulse frequency (f p ) and duration () data for the indicated proteins and nanotube tip diameters. Tip Diameter (nm) Protein f p (min -1 ) (ms) 100 nM BSA 10.5.8 520 100 nM phosphorylase B 2.5.6 1060 17 100 nM -galactosidase 1.50.3 1920870 100 nM BSA 10.1.1 440 100 nM phosphorylase B 7.8.4 663 23 100 nM -galactosidase 5.51.0 1200570 Table 3-2. Calculated diffusion coefficient for the proteins in the nanotube tip, for nanotubes with the indicated tip diameters. See text and Equations 3-2, 3-3, and 3-5 for details. Protein D sol (x 10 -7 cm 2 s -1 ) d (nm) D tip (x 10 -7 cm 2 s -1 ) D tip (x 10 -7 cm 2 s -1 ) Tip Diameter (nm) 17 23 BSA 6.00 6.8 0.400 0.36 1.69 0.296 0.50 2.58 Phosphorylase B 4.33 9.8 0.576 0.18 0.55 0.426 0.33 1.09 -Galactosidase 3.03 16.6 0.976 0.001 0.15 0.722 0.08 0.25 89

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CHAPTER 4 RESISTIVE-PULSE SENSING OF PROTEINS AT HIGH POTENTIALS USING A CONICAL NANOPORE SENSOR Introduction There is increasing interest in using nanopores in synthetic 24-31,36-54,88-94,102,166,168,170 or biological 139-149,153-164,171 membranes as resistive-pulse sensors for molecular and macromolecule analytes. The resistive-pulse method, 168 which when applied to such analytes is sometimes called stochastic sensing, 139-149 entails mounting the membrane containing the nanopore between two electrolyte solutions, applying a transmembrane potential difference, and measuring the resulting ion current flowing through the electrolyte-filled nanopore. In simplest terms, when the analyte enters and translocates the nanopore, it transiently blocks the ion current, resulting in a downward current pulse. The current-pulse frequency is proportional to the concentration of the analyte, and the identity of the analyte is encoded in the current-pulse signature, as defined by the average magnitude and duration of the current pulses. 102,103,166-168 Resistive-pulse sensing with synthetic and biological nanopores is traditionally carried out using an applied transmembrane potential of a few hundred mV or less. 20-54,88-92,138-165 In the case of biological nanopores, sensing at higher potentials is not possible due to the fragile lipid bilayer that houses the nanopore sensor. Planar lipid bilayers, typically used in biological nanopore sensors, can only withstand a few hundred mV applied transmembrane potential before rupturing. 139,167 Sensing with synthetic nanopore devices has typically been carried out using the same applied potentials as the biological sensors they are meant to mimic. 20-53 Synthetic nanopores, however, are capable of enduring much higher potentials. The Martin group has been using conical nanopores in polymer membranes as resistive-pulse sensors. 88-92 Previously reported sensing work with these devices has been carried out at potentials up to V. 88-92 However, the polymer nanopore membranes are capable of 90

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withstanding potentials up to 20 V. 127 Increasing the applied potential used for resistive-pulse sensing could offer many advantages. Previous reports have shown that as the applied transmembrane potential is increased, the standard deviation in current-pulse duration decreases. 24 Decreasing the spread in current-pulse duration would allow for better discrimination of analytes within a mixture. Increasing applied potential should also lead to increased current-pulse frequency since this would also increase the electrophoretic velocity of the charged analytes. This in turn would allow for lower limits of detection to be achieved. Here high potential (>1 V) resistive-pulse sensing of protein molecules is explored. Proteins were first sensed separately with applied transmembrane potentials from 2-4 V. The resulting affect of high potential sensing on current-pulse duration and amplitude was then examined. Two proteins with different molecular weights were then sensed in a mixture. It was determined that the proteins could be distinguished based on their current-pulse durations. The sensor element used was a conical nanopore prepared by the track-etch method 59,63 in a poly(ethylene terephthalate) (PET) membrane. The two proteins examined were bovine serum albumin (BSA) and -galactosidase. Experimental Materials Bovine serum albumin (BSA, MW ~66 kDa) and -galactosidase (MW ~116 kDa) were obtained from Sigma Aldrich. Poly(ethylene terephthalate) (PET) membranes, 12 m thick, which contained a single heavy-ion induced damage track, were obtained from GSI (Darmstadt, Germany). All other chemicals were of reagent grade and used as received. Purified water (obtained by passing house-distilled water through a Barnstead, E-pure water purification system) was used to prepare all solutions. 91

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Pore Etching and Nanotube Preparation The same cell was used for etching, electrochemical determination of the dimensions of the pore, and for the resistive-pulse experiments. 63 It is a two-compartment Kel-F cell in which the PET membrane separates the two half cells. The damage track in the PET membrane was chemically etched into a conically shaped pore using the two-step etching method described in detail previously. 63 Conically shaped nanopores have two openings: the large-diameter (or base) opening at one face of the membrane and the small-diameter (or tip) opening at the opposite face. It has been shown that the two-step etching method provides for excellent reproducibility in both the tip and the base diameters. 63 The base diameter of the pores used for these studies was 520 nm, as determined by electron microscopy. 63 The diameter of the tip opening was determined using an electrochemical method 59 described in detail in prior work. 88,89 Briefly, the membrane containing the single conical nanopore was mounted in the cell, and an electrolyte solution of measured conductivity was placed on either side of the membrane. For these studies, this solution was 1 M KCl, pH 6 with a measured conductivity of 10 S/m. A current voltage curve was obtained, the slope of which is the ionic conductance of the electrolyte-filled nanopore. The conductance is used to calculate the diameter of the tip opening. 59,88,89 The PET nanopore sensors used to here had tip diameters between 40-50 nm. Current-Pulse Measurements The membrane sample containing the single conical nanopore sensor was mounted in the cell, and both half cells were filled with ~3.5 mL of 10 mM phosphate buffer solution (pH = 7.4) that was also 100 mM in KCl. A Ag/AgCl electrode (BAS, West Lafayette, IN) was placed into each half-cell solution and connected to an Axopatch 200B (Molecular Devices Corp., Union City, CA) patch-clamp amplifier. The electrodes were arranged with polarity such that the 92

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Ag/AgCl anode was in the half-cell solution facing the base opening, and the Ag/AgCl cathode in the solution facing the tip opening. The Axopatch was used to apply the desired transmembrane potential and measure the resulting ion current flowing through the electrolyte-filled nanopore. The current was recorded in the voltage-clamp mode with a low-pass Bessel filter at 2 kHz bandwidth. The signal was digitized using a Digidata 1233A analogue-to-digital converter (Molecular Devices Corp.), at a sampling frequency of 10 kHz. Data were recorded and analyzed using pClamp 9.0 software (Molecular Devices Corp.). Because the pI values of BSA and -galactosidase are ~4.8 173 and ~4.6, 191 respectively, both proteins have net negative charge in the pH = 7.4 buffer used here. Proteins were always added to the half-cell solution facing the tip opening. All previously reported resistive-pulse sensing work has been done at potentials at or under V. In the case of biological nanopore sensors, this is because the lipid bilayer support can only withstand potentials of a few hundred mV before rupturing. 139,167 Synthetic nanopores however are capable of withstanding much higher transmembrane potentials. The main impediment on high potential sensing (>1 V) with synthetic nanopores is the instrumentation. The Axopatch 200B software used for this type of sensing has a maximum potential output of V. In order to overcome this barrier, a 9V battery was placed in series with the head stage of the Axopatch 200B. A homemade device was used to control the output from the battery, and transmembrane potentials up to 4 V were applied. Results and Discussion Steady-State Current and Current-Pulse Events for BSA and -Galactosidase In previous studies, described in Chapter 2 and 3, resistive-pulse sensing of protein analytes was carried out using a poly(ethylene glycol) (PEG) functionalized nanotube. 92 When performing sensing at high potentials it was necessary to use a bare PET nanopore with no 93

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further functionalization. This is because the higher potentials were found to disrupt the underlying gold layer of the PEG-functionalized nanopores, and after a period of time the gold layer and PEG monolayer were removed. Figure 4-1 shows current-voltage curves of a PEG functionalized nanotube before and after application of 5 V applied transmembrane potential. After the 5 V transmembrane potential was applied, the nanotube tip opening diameter increased to a value close to what was measured after the second etch step. The bare PET nanopores, however, showed a stable steady-state ion current even at high applied transmembrane potentials. Figure 4-2 shows the steady-state current values recorded for a 40 nm PET nanopore from 1-4 V applied transmembrane potential. It appears that as the potential was increased so did the absolute value of the noise (i.e., peak-to-peak noise) in the ionic current. To evaluate the noise at each potential, the relative noise was used. The relative noise for the PET nanopore was determined by dividing the average steady-state current value by the absolute noise at each potential. The relative noise at 1, 2, 3, and 4 V was 64, 97, 106 and 112, respectively. The relative noise does in fact increase slightly with increasing applied transmembrane potential. However, the current-pulse events associated with protein translocation were large enough in current-pulse amplitude to be distinguished from the noise (vide infra). When a solution 100 nM in BSA was added to the half-cell facing the tip side of the nanopore very few events were observed at 1 V applied potential. However, when the potential was increased to 2 V the frequency of translocation events from BSA increased significantly (Figure 4-3A and 4-3B). In analogy to data obtained for other charged analytes with both artificial 88,89,92 and biological 157 nanopores, there is a threshold voltage below which BSA current-pulses are not observed, and current-pulse frequency increases with applied potential 94

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above this threshold (Figure 4-3C). The threshold results from an entropic penalty that the BSA molecule pays when it enters a pore with a tip opening of comparable size to the molecule. 88,89,92,157 Because BSA is charged, this entropic barrier can be overcome by driving the BSA molecule electrophoretically into the nanopore tip. 88,89,92,157 In previous BSA sensing work with PEG functionalized nanopores, 92 current-pulse events from BSA were observed at smaller applied potentials (i.e., under 1 V). This suggests that the threshold voltage for the bare PET nanopores is somewhat higher than that of the PEG functionalized nanotubes. The reason for this higher threshold is most likely due to the surface charge on the PET nanopore. The chemical etching of PET results in COOH groups on the membrane surface and nanopore walls. 62 Functionalization of the nanopore membranes with PEG neutralizes the majority of this surface charge. When the nanopores are left unfunctionalized there will be an electrostatic repulsion between the negatively charged BSA and the negatively charge nanopore walls. Therefore, higher potentials are needed in order for the BSA molecule to overcome the entropic and electrostatic barrier. As previously mentioned, one potential advantage of sensing at high potentials is the increase in current-pulse frequency that would result. Indeed, this effect can be seen in Figure 4-3. This increase could ultimately lead to lower analyte detection limits. While the effect on detection limit has not been studied here, it was previously shown that current-pulse frequency is also dependent on analyte concentration. 89 Therefore, by increasing applied potential at decreasing analyte concentrations, an improvement in detection limits should also be achievable. Figure 4-4A shows an expanded view of a BSA current pulse. Again referring to previously described work with PEG functionalized nanotubes, the BSA current pulses dropped sharply and then tailed upward with time. 92 The BSA current pulse events obtained with bare 95

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PET nanopore sensors, however, first show a sharp increase in current that then drops and tails upward with time. The current-pulse events for -galactosidase were found to exhibit a similar shape (Figure 4-4B). The -galactosidase peaks, however, showed a larger increase in current amplitude at the beginning of the pulse than those of BSA (Figure 4-4). This suggests that the significant change in current-pulse amplitude is the upward portion of the current-pulse. Upward current-pulses have been observed in other studies. 36 These results were interpreted by noting that when a highly charged analyte enters the nanopore, it must bring its charge-balancing counterions with it. As a result, there is a transient introduction of additional charge carriers when the analyte is in the nanopore, and this accounts for the upward current-pulses. It is possible that a similar phenomenon is occurring here, as the protein analyte first encounters the entrance of the nanopore tip, which is more pronounced with bare PET nanopores, due to the presence of a surface charge. One concern when working with bare PET nanopore sensors is nonspecific adsorption of protein molecules to the nanopore walls. Current-voltage curves taken of a bare PET nanopore before and after sensing BSA showed that nonspecific adsorption was not a significant problem. From the current-voltage curves shown in Figure 4-5, the decrease in nanopore tip diameter after sensing was found to be only ~4 nm. This small change in nanopore tip diameter is not expected to affect the protein current-pulse events. Effect of Potential on Protein Translocation The protein current pulse events were characterized by the current-pulse amplitude (i) and the current-pulse duration (). The duration of the pulse is defined as the time interval between the precipitous increase in current and the time when the current returns to the baseline value. The current-pulse amplitude is defined here as the difference in current between the 96

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highest current within the initial upward pulse and the lowest current within the subsequent downward pulse. To examine the effect of applied potential on i and current pulses from BSA were examined at 2, 3, and 4 V applied transmembrane potential. Figure 4-6 shows a scatter plot of i vs. for BSA current pulses from 2-4 V applied potential. The scatter plot shows that as the applied transmembrane potential is increased, the average value of decreases as does the spread in values. The average value of i on the other hand increases. The spread in current-pulse amplitude, however, also increases from 2 to 3 volts and then remains approximately the same at 4 V. This effect is easier seen by examining the corresponding histograms of current-pulse duration and amplitude at each potential (Figure 4-7 and 4-8). The average values and standard deviations of and i were obtained by fitting the histograms with a Gaussian distribution (solid curve). 27,142,158-160 Previous reports with DNA sensing have shown a similar trend in current-pulse duration. In a study conducted by Golovchenko and coworkers, the effect of applied potential on DNA current-pulse duration was investigated. 24 They found that as the applied potential during translocation was increased from 60 mV to 120 mV, the average value of decreased as did the standard deviation in The current-pulses from -galactosidase exhibited a similar trend to that of BSA. The -galactosidase events, however, had a larger spread in current-pulse duration at each applied potential, relative to those of BSA (Table 4-1). This is to be expected since -galactosidase is a much larger protein than BSA. It was previously shown that as the size of the analyte increases so does the standard deviation in current-pulse duration. 92 Others have also reported similar results with DNA strands of increasing chain length. 24,27 The trends observed here at increasing potentials suggest that current-pulse duration could be useful in discriminating proteins in a 97

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mixture. A summary of the current-pulse duration data obtained at high potentials from bare PET nanopore sensors is shown in Table 4-1. Current-Pulse Events for a Mixture of BSA and -Galactosidase A 50 nm bare PET nanopore was first used to detect 100 nM BSA and 100 nM -galactosidase solutions individually. The histograms of current-pulse duration for BSA and -galactosidase at 2 V and 4 V applied transmembrane potential are overlaid in Figure 4-9. At both potentials there is overlap between however, the overlap at 2 V is much more severe. Even though there is still some overlap at 4 V, two peaks can be easily distinguished. Therefore, resistive pulse sensing of the mixture of the two proteins was carried out at 4 V applied potential. A solution 50 nM in BSA and 50 nM in -galactosidase was added to the half-cell in contract with the tip side of the nanopore membrane. When a 4 V transmembrane potential was applied current-pulse events from the protein mixture were observed (Figure 4-10). From the i-t trace it is obvious that there is a decrease in current-pulse frequency. The reasons for this decrease are still under investigation. It could be possible that the two proteins are aggregating in solution making translocation of the much larger aggregates impossible. It could also simply be due to the lower concentrations of each analyte used. Even though the total concentration of analyte (100 nM) is the same, the proteins each have very different current-pulse frequencies, due to differences in size and charge. Figure 4-11 shows a histogram of the current-pulse durations for the protein mixture. Two peaks with little overlap can be clearly distinguished. The peaks were fit with a Gaussian distribution. The first peak has a avg = 60 ms and corresponds to the current-pulse events from BSA. While the second peak has a avg = 230 ms and corresponds to the current-pulse events from -galactosidase. The average and standard deviation in fits extremely well with 98

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the values obtained when the proteins were sensed individually (Table 4-1). These results show that by decreasing the spread in current-pulse duration, discrimination of proteins within a mixture is a possibility. Conclusions Increasing the applied potential used for resistive-pulse sensing could offer several advantages. Here one of these advantages was demonstrated. It was shown that high potential sensing of protein analytes results in a significant decrease in the spread in current-pulse durations. It was also shown that high potential sensing of a mixture of two proteins allows for the two proteins to be discriminated. Thus, applied potential is an important factor in the optimization of protein resistive-pulse sensors. While the effect of high potential on the limit of detection and current-pulse frequency was not examined here, this could also be another significant advantage to result from high potentials. The advantages of the synthetic nanopore resistive-pulse sensors have also been demonstrated here. These advantages include chemical and mechanical stability of the synthetic polymer membrane, the ability to tune the size of the nanopore tip opening, and the ability to apply much larger transmembrane potentials than have been previously reported with similar work. 24-53 99

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Current (nA) -25-20-15-10-5051015-1.5-1-0.500.511.5Potential (V) Figure 4-1. Current-voltage curves of a PEG functionalized nanotube sensors before and after application of 5 V applied transmembrane potential. Before application of 5 V transmembrane potential, black : tip diameter = 22 nm. After application of 5 V transmembrane potential, red : tip diameter = 43 nm. BCDAIm Scaled(pA) 1550 1600 1650 Im Scaled(pA) 2850 2900 2950 Im Scaled(pA) 4150 4200 4250 4300 Im Scaled(pA) 5500 5550 5600 5650 50 pA5 sec1600 pA2900 pA4250 pA5580 pAIm Scaled(pA) 1550 1600 1650 Im Scaled(pA) 2850 2900 2950 Im Scaled(pA) 4150 4200 4250 4300 Im Scaled(pA) 5500 5550 5600 5650 Im Scaled(pA) 1550 1600 1650 Im Scaled(pA) 2850 2900 2950 Im Scaled(pA) 4150 4200 4250 4300 Im Scaled(pA) 5500 5550 5600 5650 50 pA5 sec 50 pA5 sec1600 pA2900 pA4250 pA5580 pA Figure 4-2. Current-time transients for a bare PET conical nanopore sensor with buffer only. A) 1 V, B) 2 V, C) 3 V, and D) 4 V applied transmembrane potential. Tip diameter = 40 nm. 100

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CBA 500 pA5 sec 500 pA5 sec Figure 4-3. Current-time transients for a bare PET conical nanopore sensor with BSA. BSA current-pulse events at A) 1 V, B) 2 V, and C) 4 V applied transmembrane potential. BSA concentration = 100 nM. Tip diameter = 40 nm. 200 pA100 ms 200 pA100 ms 200 pA100 ms 200 pA100 ms AB Figure 4-4. Expanded views of typical current pulses associated with tip-to-base translocation of BSA and -galactosidase. A) BSA (100 nM) and B) -galactosidase (100 nM). Tip diameter = 50 nm. Transmembrane potential = 4 V. 101

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Current (nA) -20-15-10-505101520-1.5-1-0.500.511.5Potential (V) Figure 4-5. Current-voltage curves of a bare PET conical nanopore sensor taken before and after sensing a 100 nM BSA solution. Before sensing BSA, black : tip diameter = 40 nm. After sensing BSA, red : tip diameter = 36 nm. 02004006008001000120014001600200400600800100012001400 i(pA) (ms) Figure 4-6. Scatter plot of current-pulse magnitude (i) versus current-pulse duration () for BSA at various applied transmembrane potentials. Transmembrane potential of 2 V (black ), 3 V (red ), and 4 V (blue ). BSA concentration = 100 nM. Tip diameter = 40 nm. 102

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0501001502002500510152025 Counts(ms) 02004006008001000010203040 Counts (ms) 02004006008001000120014001600010203040 Counts(ms)ABC Figure 4-7. Histograms of BSA current-pulse duration data. A) 100 nM BSA at an applied potential = 2 V, = 260 pA. B) 100 nM BSA at an applied potential = 3 V, = 160 pA. C) 100 nM BSA at an applied potential = 4 V, = 60 pA. Solid curves are Gaussian fits. Tip diameter = 40 nm. 103

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9001000110012001300140002468101214 Countsi (pA) 5006007008009001000010203040 i (pA)Counts 20030040050060070001020304050 Countsi (pA)ABC Figure 4-8. Histograms of BSA current-pulse amplitude data. A) 100 nM BSA at an applied potential = 2 V, i = 340 pA. B) 100 nM BSA at an applied potential = 3 V, i = 750 pA. C) 100 nM BSA at an applied potential = 4 V, i = 1160 pA. Solid curves are Gaussian fits. Tip diameter = 40 nm. 104

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0400800120016000246810 Counts(ms) 01002003004000246810 Counts(ms)AB Figure 4-9. Histograms of BSA and -galactosidase current-pulse duration data. A) 100 nM BSA (gray), 100 nM -galactosidase (white with black lines). Applied potential = 2 V. B) 100 nM BSA (gray), 100 nM -galactosidase (white with black lines). Applied potential = 4 V. Tip diameter = 50 nm. BSA and -galactosidase solutions sensed separately. 100 pA5 sec 100 pA5 sec Figure 4-10. Current-time transient for a bare PET conical nanopore sensor with a solution of BSA and -galactosidase on the tip side of the membrane. Concentration of BSA = 50 nM. Concentration of -galactosidase = 50 nM. Tip diameter = 50 nm. Transmembrane potential = 4 V. 105

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(ms) Counts 010020030040050060005101520253035 Figure 4-11. Histogram of current-pulse duration for a solution containing a mixture of BSA and -galactosidase. Solid curves are Gaussian fits. Concentration of BSA = 50 nM. Concentration of -galactosidase = 50 nM. Tip diameter = 50 nm. Transmembrane potential = 4 V. 106

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Table 4-1. Summary of current-pulse duration data obtained with three different bare PET conical nanopore sensors. Nanopore Tip Diameter (nm) Applied Potential (V) 100 nM BSA avg (ms) 100 nM -Galactosidase avg (ms) 2 260 n/a 3 160 40 4 60 2 300 n/a 3 120 45 4 50 2 170 890 3 100 410 50 4 50 220 107

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CHAPTER 5 DETERMINATION OF THE SURFACE ISOELECTRIC POINT OF PROTEINS IMMOBILIZED ON POLYMER NANOPORE MEMBRANES BY CURRENT-VOLTAGE CURVES Introduction There is increasing interest in using nanopores in synthetic 24-94 or biological 138-165 membranes as biosensors. Perhaps the most popular nanopore-based sensing paradigm is the well-known resistive-pulse, or stochastic-sensing, method which entails counting individual analyte molecules as they are driven through the nanopore sensor element. 102,103,167,168 Prototype sensors for analyte species as diverse as proteins, 39,92,149 DNA 24,26,27,30-34,36,46,89,90,140,157,158,160-162 and small molecules 88,147 have been described. While the Martin group too is exploring resistive-pulse sensors, 88-92 they have also been investigating other sensing paradigms based on artificial nanopores. 93,94 The sensing paradigm used here makes use of the well-known ion current rectification phenomenon displayed by conically shaped nanopores 69-76 to determine the isoelectric point (pI) of protein molecules immobilized on the nanopore walls. The sensor element in this case is a single, conically shaped nanopore, track-etched in a poly(ethylene terephthalate) (PET) membrane. 59,63 The sensing paradigm entails placing electrolyte solutions on either side of the membrane and using electrodes in each solution to scan the applied transmembrane potential and measure the resulting ion current flowing through the nanopore. As has been discussed in detail by our group 73-75 and others, 62,69-73,195,196 conically shaped nanopores with excess surface charge on the pore walls, and sufficiently small tip openings, show non-linear current-voltage curves (i.e., such pores are ion-current rectifiers). Because the PET nanopores used here have excess anionic surface charge, rectification is observed at positive applied transmembrane potentials (anode facing the base side of the nanopore). 108

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It was found that when proteins are immobilized on the nanopore walls, the rectification behavior of the nanopore changes due to the charge from the protein molecules. At pHs above the pI of the protein-modified surface, there will be a net negative charge on the protein and current-voltage curves will show rectification at positive applied transmembrane potentials. Below the pI of the protein-modified surface, there will be a net positive charge on the protein and current-voltage curves will show rectification at negative applied transmembrane potentials. At the pH corresponding to the pI of the protein-modified nanopore surface, the surface will be neutral and therefore current-voltage curves should show no rectification and appear linear. Therefore, the extent of rectification of protein-modified nanopores depends on the pH at which the current-voltage curves are taken. The pI of the immobilized proteins can be determined by examining current-voltage curves and ion current rectification over a range of pHs. The pI of proteins in solution can be easily determined through methods such as isoelectric focusing (IEF), 197,198 however, there are no known studies that have determined the pI of immobilized proteins. Here the effect of immobilization on the pI of proteins is investigated. The effect of the method of protein immobilization on pI determination has also been studied. This new nanopore-based sensing paradigm is described here using three different proteins with known pIs of the free proteins ranging from ~3-6. The proteins used were bovine serum albumin (BSA), amyloglucosidase, and phosphorylase B. Immobilization of the proteins on the nanopore walls and membrane faces was carried out using two different techniques. The first technique utilized the well know EDC chemistry to attach the proteins via formation of an amide bond. 90,131-136 The second technique was a two step method where ethanolamine was first attached to the nanopore membrane via EDC chemistry. 90 This was done to neutralize some of the surface charge of the polymer membrane. The protein molecules were then nonspecifically 109

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adsorbed onto the PET surface. Control studies were also carried out with nanopores modified with only ethanolamine. Experimental Materials BSA (MW ~66 kDa, pI ~4.8 173 ), anti-BSA-Fab (MW ~50 kDa), amyloglucosidase (MW ~97 kDa, pI ~3.6 199 ), and phosphorylase B, rabbit (MW ~97.4 kDa, pI ~5.8-6.3 189,190 ) were obtained from Sigma Aldrich, as was the control molecule, ethanolamine hydrochloride. Poly (ethylene terephthalate) (PET) membranes, 12 m thick, which contained a single heavy-ion induced damage track, were obtained from GSI (Darmstadt, Germany). 1-Ethyl-3-[3-dimethylaminopropyl] carbodiimide hydrochloride (EDC), N-hydroxysulfosuccinimide (Sulfo-NHS), and 2-(N-morpholino) ethanesulfonic acid buffered saline (MES) were obtained from Pierce. Isoelectric focusing (IEF) gels, buffers and standards were obtained from Invitrogen. All other chemicals were of reagent grade and used as received. Purified water (obtained by passing house-distilled water through a Barnstead, E-pure water purification system) was used to prepare all solutions. Pore Etching The damage track in the PET membrane was chemically etched into a conically shaped pore using the two-step etching method described in detail previously. 63 Conically shaped nanopores have two openings: the large-diameter (or base) opening at one face of the membrane and the small-diameter (or tip) opening at the opposite face. It has been shown that the two-step etching method provides for excellent reproducibility in both the tip and the base diameters. 63 The base diameter of the pores used for these studies was 520 nm, as determined by electron microscopy. 63 110

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The diameter of the tip opening was determined using an electrochemical method 59 described in detail in prior work. 88,89 Briefly, the membrane containing the single conical nanopore was mounted between two halves of an electrochemical cell, and an electrolyte solution of measured conductivity was placed on either side of the membrane. For these studies, this solution was 1 M KCl, pH 6 with a measured conductivity of 10 S/m. A current voltage curve was obtained, the slope of which is the ionic conductance of the electrolyte-filled nanopore. The conductance is used to calculate the diameter of the tip opening. 59,88,89 The nanopores used for these studies had tip diameters of 40-50 nm before protein modification (vide infra). PET Surface Modification The PET nanopore walls and membrane faces were modified with proteins using two different immobilization techniques; (i) EDC/sulfo-NHS chemistry and (ii) nonspecific protein adsorption over membranes modified with ethanolamine via EDC/sulfo-NHS chemistry. Nanopores modified with only ethanolamine via EDC/sulfo-NHS chemistry were also used for control studies. The pore-etching procedure generates carboxylate groups on the pore walls and membrane faces. 62 The well known EDC chemistry was used to attach proteins and the control molecule ethanolamine to these -COO groups via amide bond formation. 134-136 This was accomplished by first immersing the membrane into a solution that was 1 mM in EDC and 1.4 mM in sulfo-NHS dissolved in 10 mM MES-buffered saline, pH 6, for 2 hours. The membrane was then rinsed and immersed into a protein solution (2 mg/7 mL) dissolved in 10 mM phosphate buffered saline (PBS), pH 7.4, for 2 hours at 4 o C. The membranes that were used as controls were placed in a solution 1 mg/mL in ethanolamine dissolved in 10 mM PBS, pH 7.4, for 2 hours after being 111

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removed from the EDC/sulfo-NHS solution. The EDC coupling chemistry is shown schematically in Figure 5-1. Protein immobilization via nonspecific adsorption was accomplished by first modifying the pore walls and membrane faces with ethanolamine using the EDC/sulfo-NHS method described above for the control membranes. Ethanolamine was used to neutralize some of the surface charge resulting from the -COO groups produced during etching. 90 The ethanolamine modified membranes were then immersed in a protein solution (2 mg/7 mL) dissolved in 10 mM phosphate buffered saline (PBS), pH 7.4, overnight (~15 hours) at 4 o C. The modified nanopore tip diameters were measured using the same electrochemical method described previously. The nanopore tips showed a decrease in tip diameter after protein modification. The amount of decrease varied depending on the protein and modification technique used. The control membranes modified with ethanolamine showed little to no change after modification. Attachment of the protein BSA was also confirmed using X-ray photoelectron spectroscopy (XPS, Perkin-Elmer PHI 5100). Current-Voltage Measurements and Isoelectric Point Determination To obtain the current-voltage curve measurements used for pI determination the membrane sample was mounted between the two halves of the conductivity cell, 63 and each half-cell was filled with 3.5 mL of 10 mM KCl in 10 mM PBS. A Ag/AgCl electrode was inserted into each half-cell solution and a Keithley 6487 picoammeter/voltage source (Keithley Instruments Inc.) was used to scan the transmembrane potential from -1 to +1 V and measure the resulting ion current flowing through the nanopore. The electrodes were arranged so that at -1 V the anode was in the solution facing the tip of the pore and at +1 V the anode was in the solution facing the base of the pore. Current-voltage curves were obtained before and after protein modification at pHs between 1.5 and 7. The rectification ratios (vide infra) obtained from the current-voltage 112

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curves at each pH were used to determine the pI of the immobilized proteins on the polymer surface. Isoelectric Focusing IEF was performed on the three proteins investigated here in order to confirm the pI of the proteins in solution. This was done using Novex precast pH 3-10 gels, IEF buffers, and IEF markers. The gels were run at 100 V for 1 hour, 200 V for 1 hour and then 300 V for 30 min. The gels were then stained for 1 hour in SimplyBlue SafeStain. The concentration of all protein samples was 1mg/mL. Results and Discussion Surface Protein Determination Attachment of one of the proteins used in these studies, BSA, was confirmed using XPS. The unmodified PET membrane exhibits strong signals for C and O (Figure 5-2A). The XPS spectra of PET membranes modified with BSA via EDC/sulfo-NHS chemistry and nonspecific adsorption both show the addition of a N signal, confirming the presence of the protein on the surface (Figure 5-2B and 5-2C). The peaks at 720 eV and 990 eV are Auger peaks for oxygen and carbon, respectively. The surface atomic percentages obtained from the XPS spectra are shown in Table 5-1. Considering the bare (not reacted with BSA or ethanolamine) membrane first, we see that the percents of C and O are in perfect agreement with values calculated from the empirical formula of the PET monomer, C 10 O 4 H 8 It is worth mentioning, however, that since XPS does not detect H, the atomic percentages calculated from the monomer are 71.4% and 28.6% for C and O, respectively. The PET membranes modified with BSA via EDC/sulfo-NHS chemistry show a decrease in C and O content of 5% and 0.6%, respectively, and an increase in N content from zero to 113

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5.6%. The membranes modified with BSA via nonspecific adsorption over ethanolamine modified membranes show a decrease in C and O content of 2.7% and 2.4%, respectively, and an increase in N content from zero to 5.2%. It should be mentioned that the presence of ethanolamine will also contribute to the N signal. XPS spectra of PET membranes modified with only ethanolamine via EDC/sulfo-NHS chemistry (Figure 5-2D) show a N content of only 0.9% (Table 5-1). Therefore, the increase in N content seen in the XPS spectra, upon adsorption of BSA, results from the presence of the protein. Previous studies have shown that XPS and elemental analysis can be used to characterize and quantify the surface content of proteins immobilized on polymer surfaces. 200 XPS provides surface atomic percentages within the first 500 nm of the outer surface. 201 Combining XPS results with the calculated elemental contents of each component allows one to calculate the percentage of these components present on the surface of the sample. The elemental contents of the polymer membrane and BSA can be calculated from the empirical formula of the monomer and amino acid composition of the protein, 202 respectively. The bare PET polymer membrane contains no N and a C content of ~71%. The C and N content of BSA is ~56% and ~15%, respectively. For the BSA modified nanopore membranes the surface weight fractions of the PET polymer (P) and BSA (B) must follow Equation 5-1: P + B = 100 (5-1) The percentages of C and N present on the surface of the modified membranes can be calculated from the following two equations, C = 71%P + 56%B (5-2) N = 15%B (5-3) 114

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From the XPS data (Table 5-1), and Equations 5-1 to 5-3, the ratio of N/C can be used to solve for P and B. The PET membrane modified with BSA via EDC/sulfo-NHS chemistry has a surface weight fraction of BSA equal to ~37%. As previously mentioned, the membranes that were modified with BSA via nonspecific adsorption over ethanolamine-modified PET nanopores will also have a contribution of N from the ethanolamine. Therefore, in order to determine the surface weight fraction from BSA with these membranes it is first necessary to determine the contribution from ethanolamine. This can be done by first changing Equations 5-1 to 5-3 to fit a membrane modified with only ethanolamine (E). The content of C and N in ethanolamine, calculated from the empirical formula, is ~50% and ~25%, respectively. Using the XPS data from the ethanolamine modified membrane, and the modified equations, gives a surface weight fraction of ethanolamine equal to ~3%. The surface weight fraction of BSA for membranes modified via nonspecific adsorption over ethanolamine membranes can now be calculated, assuming that the contribution from ethanolamine will be the same as when no protein is present. For these membranes, Equation 5-1 now becomes, P + B + E = 100 (5-4) Because E was determined to be ~3% from the membranes modified with only ethanolamine, Equation 5-4 can be rewritten as, P + B = 97 (5-5) The equations for the percentages of C and N present on the surface must also be rewritten to include the contribution from E as follows, C = 71%P + 56%B + 50%E (5-6) 115

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N = 15%B + 25%E (5-7) From the XPS data (Table 5-1), and Equations 5-5 to 5-7, the ratio of N/C can again be used to solve for P and B. The PET membrane modified with BSA via nonspecific adsorption over ethanolamine-modified membranes has a surface weight fraction of BSA equal to ~29%. These calculations indicate that nonspecific adsorption of the proteins yields slightly less surface coverage than immobilization via EDC/sulfo-NHS chemistry. To demonstrate that the proteins were present on the nanopore walls and still active after EDC/sulfo-NHS modification, current-voltage curves were taken in 1M KCl, pH 6, before and after modification of the PET surface, and after addition of a binding and non-binding antibody to a solution in contact with the nanopore membrane. Current-voltage curves of single conical nanopores modified with BSA and phosphorylase B via EDC/sulfo-NHS chemistry (Figure 5-3) both show a decrease in ionic current after modification, which corresponds to a decrease in nanopore tip diameter. The decrease in tip diameter indicates that the protein is not only present on the membrane surface (as shown by XPS) but the nanopore walls as well. To show that the proteins remained active after immobilization, the BSA and phosphorylase B modified nanopore membranes were soaked in a solution 1 M in anti-BSA-Fab for 1 hour. The membranes were then rinsed and the current-voltage curves repeated. The BSA modified nanopore showed a further decrease in ionic current, corresponding to a further decrease in nanopore tip diameter, after soaking in the anti-BSA-Fab (Figure 5-3A). This further decrease in tip diameter is a result of the anti-BSA-Fab binding to the BSA immobilized on the nanopore walls. In contrast, the phosphorylase B modified nanopore showed no change in the current-voltage curves (Figure 5-3B). This is because the anti-BSA-Fab does not bind to the phosphorylase B and therefore results in no change in the nanopore tip diameter. These results 116

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indicate that the proteins present on the nanopore membrane are still active and show selectivity after immobilization. Determination of Isoelectric Points from Current-Voltage Curves Current-voltages can also provide information about the surface charge on the pore walls. 73 It is well known that when the radius of the tip opening of the conical nanopore is comparable to the thickness of the electrical double layer associated with the fixed surface charge on the pore wall, the nanopore will rectify ion current (i.e., show a non-linear current voltage curve). The ion-current rectification phenomenon can be described by defining on and off states for the nanopore rectifier. 74 The reasons for this ion-current rectification have been discussed in detail in the literature. 73,169 If however, the surface charge is removed, then the pore will show a linear current-voltage curve. 73 PET nanopores carry a negative surface charge at neutral pH due to the carboxylate groups produced during etching. 62 Current-voltage curves of bare PET nanopores will show a non-linear current-voltage curve with the on state at negative applied potentials and the off state at positive potentials. 73 The extent of ion current rectification can be quantified via the rectification ratio, r.r., 74 VViirr11.. (5-8) where i -1V is the current at an applied transmembrane potential of -1V and i +1V is the current at an applied transmembrane potential at +1 V. When the nanopore carries a negative surface charge, as described above with bare PET, and the r.r. will be greater than one. If the nanopore carries a positive surface charge, the direction of rectification will then be reversed, with the off state at negative potentials and the on state at positive potentials, and the r.r. will be less than 1. 117

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Proteins also carry charge, and when immobilized on the nanopore walls it is expected that these molecules will contribute to the majority of the surface charge on the pore. The immobilized proteins will have a certain pI, where all charges on the protein are neutralized. When a current-voltage curve is taken at the pH that corresponds to the pI the pore will show a linear current-voltage curve. At this pH the r.r. value will be equal to one since no rectification is observed. Above the pI, the nanopore surface will be negative, and below the pI, positive. At these pH values current-voltage curves will show rectification and r.r. greater than or less than one, respectively. Therefore by obtaining current-voltage curves at various pH values and examining the rectification ratios relative to the pH, one can determine the pI of protein films immobilized on the nanopore surface. PET and Ethanolamine Modified Nanopores Before modifying the PET single nanopores, current-voltage curves were taken in 10 mM KCl with 10 mM PBS at pHs of 1.5, 3, 5 and 7. The rectification ratios at each pH were obtained from the current-voltage curves and plotted vs. pH. The rectification ratios were plotted on a log scale and a linear fit was applied to the data. The pH at which the rectification ratio equaled one was then extracted from the linear fit. This pH was taken as the pI of the surface. The data from three PET nanopore membranes is shown in Figure 5-4. From all PET nanopore data obtained (17 pores), the average pI value was 2.57.63. The reported pI of PET is ~3. 62 Control membranes were modified with ethanolamine via EDC/sulfo-NHS chemistry. Current-voltage curves taken in 1 M KCl, pH 6, were taken to measure the tip diameter after ethanolamine modification (Figure 5-5). The nanopore tip diameters showed little to no change after modifying with ethanolamine, giving an average change in tip diameter of 1 nm. Current-voltage curves were also taken in 10 mM KCl with 10 mM PBS at pHs of 1.5, 3, 5 and 7, in order to monitor the change in rectification and determine the pI of the surface. The plots 118

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of rectification ratio vs. pH for the ethanolamine modified nanopore membranes gave an average pI of 3.74.70. The data from three ethanolamine modified nanopore membranes is shown in Figure 5-6. These results indicate that there is still surface charge present after modification with ethanolamine. 90 Attachment of ethanolamine to the carboxylate groups on the PET membrane lowers the anionic charge density on the surface, however, does not remove 100% of the carboxylate sites on the PET surface. 90 BSA, Phosphorylase B and Amyloglucosidase Nanopores Modified via EDC/Sulfo-NHS Chemistry In contrast to the ethanolamine-modified control membranes, the nanopores modified with proteins via EDC/sulfo-NHS chemistry all showed a significant decrease in nanopore tip diameter. Nanopores modified with BSA showed an average decrease in nanopore tip diameter of ~17 nm. The amyloglucosidase modified pores showed a decrease of ~12 nm and the phosphorylase B modified pores decreased and average of ~24 nm (Figure 5-7). The average change in tip diameter for all the modified nanopore membranes is summarized in Table 5-2. After protein modification, current-voltage curves were again taken in 10 mM KCl with 10 mM PBS from pHs 1.5-7. The changes in ion current rectification can be seen by comparing these current-voltage curves with the ones obtained before modification (bare PET nanopore). The typical current-voltage curves obtain before and after modification, at various pHs, are shown in Figure 5-8 for each of the three protein-modified pores. The current-voltage curves for the amyloglucosidase modified nanopore showed the least noticeable change in rectification, while those from the phosphorylase B modified nanopore showed the most. This is because the known pI of free amyloglucosidase (pI ~3.6) 199 is similar to that of the bare PET surface (pI ~3). 62 Immobilization of this protein, therefore, does not greatly alter the surface charge of the nanopore. Phosphorylase B, however, has a reported pI value between 5.8 and 6.3. 189,190 Of all 119

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the proteins examined here, this protein has an pI value that is farthest away from that of the bare PET surface. Therefore, immobilization of this protein is expected to cause the greatest change in the surface charge of the pore, and indeed, the greatest change in rectification behavior is observed with this protein. The plots of rectification ratio vs. pH for each of the protein-modified nanopore membranes are shown in Figure 5-9. As with the bare PET membranes, the data was plotted on a semi-log plot and fit linearly. From the linear fit, the pH corresponding to the pI was extracted. The experimentally determined pIs of the BSA, amyloglucosidase, and phosphorylase B modified nanopores were 4.05.07, 2.92.10, and 5.33.29, respectively. The proteins pI values obtained from current-voltage curves, IEF, and the literature are summarized in Table 4-3. The experimentally determined pI values are slightly lower than the reported pI values for the free proteins. There are two possible explanations for this result. The first could be that some COO groups on the PET surface, left unfunctionalized after protein immobilization, are interfering with the pI determination. Another reason for the lower pIs could be a result of the immobilization process. Amine groups on the proteins are utilized during immobilization with EDC/sulfo-NHS, which could also effect the pI of the protein. BSA, Phosphorylase B and Amyloglucosidase Nanopores Modified via Nonspecific Adsorption In order to determine if the method of protein immobilization affected the surface pI, proteins were also immobilized via nonspecific adsorption onto the polymer surface. As previously mentioned these nanopore membranes were modified with ethanolamine via EDC/sulfo-NHS chemistry prior to protein immobilization. This was done in order to neutralize some of the surface charge on the bare PET membrane. Nanopores modified with this technique also showed a significant decrease in tip diameter after protein immobilization. Nanopores 120

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modified with BSA decreased and average of ~14 nm, amyloglucosidase decreased ~14 nm, and phosphorylase B decreased ~39 nm (Table 5-2). The plots of rectification ratio vs. pH for each of the nanopores modified with this technique are shown in Figure 5-10. Nanopores modified with this technique tended to show a greater variation in rectification ratio at high and low pHs. However, the rectification ratios around the pI value were very reproducible. From these plots, the experimentally determined pIs of the BSA, amyloglucosidase, and phosphorylase B modified nanopores were 4.50.15, 3.99.13, and 5.47.05, respectively. These results do show a slight variation from the pores modified via EDC/sulfo-NHS chemistry, and are actually closer to the reported values of the free proteins (Table 5-3). The reasons for this variation still needs further exploration, since a number of factors could be contributing to the results. It is possible that functionalization with ethanolamine prior to protein immobilization eliminates some interference from any exposed PET surface. Nonspecific adsorption also results in a variety of orientations of the immobilized proteins, 203 whereas with EDC/sulfo-NHS immobilization the proteins are all attached to the surface via an amide bond. This variation in protein orientation could lead to and pI closer to that of the free proteins. Conclusion In this work it was demonstrated that single conical nanopores in PET membranes can be used as pI sensors. Determination of the surface isoelectric point of proteins immobilized on the PET nanopore sensors can be easily achieved from current-voltage curves. It has also been shown that proteins can be immobilized on the nanopore walls using both EDC/sulfo-NHS chemistry and nonspecific adsorption. Furthermore, it was shown that the rectification behavior 121

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and the pI of the nanopore surface changes as a result of protein immobilization. The amount of change depended on the pI of the protein immobilized as well as the immobilization technique. The reasons we obtain different pI values for the immobilized and free proteins are still being investigated. These differences could be from interference from any exposed PET surface or a result of the immobilization of the proteins. However, previous studies of the surface isoelectric point of native air-formed oxide films found that the pI of the surface films differed between one to three pH units of the reported pIs of the bulk oxide powders. 204 These results were obtained by contact angle titration. Although further study is needed, this device shows promise as a useful sensor for surface isoelectric point determination. The technique is relatively straightforward and fast, and could also be applied to any type of modified or unmodified conical nanopore to determine the pI of the surface. 122

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Figure 5-1. Schematic of the EDC/sulfo-NHS chemistry. The scheme shown is for modification with ethanolamine. For protein immobilization, ethanolamine is replaced with the desired protein. [adapted Kececi, K.; Sexton, L. T.; Buyukserin, F.; Martin, C. R. Nanomedicine 2008, 3, 787-796.] 123

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Binding Energy (eV)N(E)Min: 910Max: 160510 1000900 800 700 600 500 400 300 200 100 0 ABinding Energy (eV)N(E)Min: 910Max: 160510 1000900 800 700 600 500 400 300 200 100 0 Binding Energy (eV)N(E)Min: 910Max: 160510 1000900 800 700 600 500 400 300 200 100 0 AO 1sC 1s Binding Energy (eV)N(E)Min: 720Max: 258867 1100990 880 770 660 550 440 330 220 110 0 BO 1sC 1sN 1sBinding Energy (eV)N(E)Min: 720Max: 258867 1100990 880 770 660 550 440 330 220 110 0 BBinding Energy (eV)N(E)Min: 720Max: 258867 1100990 880 770 660 550 440 330 220 110 0 Binding Energy (eV)N(E)Min: 720Max: 258867 1100990 880 770 660 550 440 330 220 110 0 BO 1sC 1sN 1s Figure 5-2. XPS spectra before and after membrane modification. A) Bare PET membrane. B) PET membrane modified with BSA via EDC/sulfo-NHS. C) PET membrane modified with BSA via nonspecific adsorption over an ethanolamine modified surface. D) PET membrane modified with ethanolamine. 124

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Binding Energy (eV)N(E)Min: 24360Max: 147820 10009008007006005004003002001000 O 1sC 1sN 1sCBinding Energy (eV)N(E)Min: 24360Max: 147820 10009008007006005004003002001000 O 1sC 1sN 1sC Binding Energy (eV)N(E) 10009008007006005004003002001000 N 1s DO 1sN 1sC 1s Figure 5-2. Continued 125

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-25-20-15-10-505101520-1.5-1-0.500.511.5Potential (V)Current (nA)A -25-20-15-10-505101520-1.5-1-0.500.511.5Current (nA)Potential (V)B Figure 5-3. Current-voltage curves taken in 1 M KCl before and after modification with a protein, and after further modification with an antibody. A) PET (black : d~55 nm), after modification with BSA via EDC/sulfo-NHS chemistry (red : d~37 nm), and after soaking in a solution 1 m in anti-BSA-Fab for 1 hour (blue : d~27 nm). B) PET (black : d~50 nm), after modification with phosphorylase B via EDC/sulfo-NHS chemistry (red : d~26 nm), and after soaking in a solution 1 m in anti-BSA-Fab for 1 hour (blue : d~25 nm). 123456780.1110 Rectification RatiopH Figure 5-4. Plot of rectification ratio versus pH for three PET single nanopore membranes. 126

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-25-20-15-10-50510152025-1.5-1-0.500.511.5Potential (V)Current (nA) Figure 5-5. Current-voltage curves taken after chemical etching of PET (black : d~55 nm) and after modification with ethanolamine via EDC/sulfo-NHS chemistry (red : d~52 nm). 12345670.010.11 Rectification RatiopH Figure 5-6. Plot of rectification ratio versus pH for three ethanolamine modified single nanopore membranes. 127

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-25-20-15-10-50510152025-1.5-1-0.500.511.5Potential (V)Current (nA)A -25-20-15-10-50510152025-1.5-1-0.500.511.5Potential (V)Current (nA)B Figure 5-7. Current-voltage curves taken in 1 M KCl before and after modification to measure the change in tip diameter. A) PET (black : d~54 nm), after modification with BSA via EDC/sulfo-NHS (red : d~39 nm), B) PET (black : d~44 nm), after modification with amyloglucosidase via EDC/sulfo-NHS (red : d~33 nm), and C) PET (black : d~55 nm), after modification with phosphorylase B via EDC/sulfo-NHS (red : d~35 nm). 128

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C -25-20-15-10-50510152025-1.5-1-0.500.511.5Potential (V)Current (nA)C Figure 5-7. Continued 129

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-3-2-1012345-1.5-1-0.500.511.5 -1.2-1-0.8-0.6-0.4-0.200.20.40.6-1.5-1-0.500.511.5pH 1.6pH 5Current (nA)Potential (V)Potential (V)A -1-0.8-0.6-0.4-0.200.20.40.60.8-1.5-1-0.500.511.5 -2-1.5-1-0.500.511.5-1.5-1-0.500.511.5pH 1.6pH 5Current (nA)Potential (V)Potential (V)B Figure 5-8. Current-voltage curves taken in 10 mM KCl, 10 mM PBS buffer before and after modification with proteins at pH 1.6 and 5. Black is before modification, A) red is after modification with BSA, B) red is after modification with amyloglucosidase, and C) red is after modification with phosphorylase B. 130

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-2-101234567-1.5-1-0.500.511.5 -1.2-1-0.8-0.6-0.4-0.200.20.4-1.5-1-0.500.511.5Current (nA)Potential (V)Potential (V)pH 1.6pH 5C Figure 5-8. Continued 131

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12345670.1110 Rectification RatiopH1.52.02.53.03.54.04.55.05.51 Rectification RatiopH5.05.25.45.65.86.06.26.46.60.1110 Rectification RatiopHABC12345670.1110 Rectification RatiopH1.52.02.53.03.54.04.55.05.51 Rectification RatiopH5.05.25.45.65.86.06.26.46.60.1110 Rectification RatiopHABC Figure 5-9. Plot of rectification ratio versus pH for protein modified nanopores. A) BSA, B) amyloglucosidase, and C) phosphorylase B modified single nanopore membranes. The proteins were immobilized via EDC/sulfo-NHS chemistry. 132

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12345670.1110 Rectification RatiopH12345670.010.11 Rectification RatiopH4.55.05.56.06.57.07.50.1110100 Rectification RatiopHABC12345670.1110 Rectification RatiopH12345670.010.11 Rectification RatiopH4.55.05.56.06.57.07.50.1110100 Rectification RatiopHABC Figure 5-10. Plot of rectification ratio versus pH for protein modified nanopores. A) BSA, B) amyloglucosidase, and C) phosphorylase B modified single nanopore membranes. The proteins were immobilized via nonspecific adsorption over ethanolamine-modified PET membranes. 133

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Table 5-1. Percent atomic composition of PET membrane before (bare) and after modification with ethanolamine and BSA. Atom Percent Bare PET Ethanolamine modified via EDC/sulfo-NHS BSA modified via EDC/sulfoNHS BSA modified via nonspecific adsorption (modified with ethanolamine prior to adsorption) C 71.4 79.7 66.4 68.7 O 28.6 19.4 28 26.2 N 0 0.9 5.6 5.2 Table 5-2. Average change in nanopore tip diameter with type of molecule and method of immobilization. Molecule Average Decrease in Tip Diameter (nm) Immobilized via EDC/sulfoNHS Immobilized via nonspecific adsorption Ethanolamine 1 n/a BSA 17 14 Amyloglucosidase 12 14 Phosphorylase B 24 39 Table 5-3. Comparison of protein pI values determined from current-voltage curves, IEF and literature. Protein pI Value Literature* IEF* Modified via EDC/sulfoNHS Modified via nonspecific adsorption Amyloglucosidase 3.6 3.5 2.92.10 3.99.13 BSA 4.8 4.5 4.05.15 4.50.15 Phosphorylase B 5.8-6.3 6.0-6.9 5.33.29 5.47.05 Free protein 173,189,190,199 Immobilized on polymer surface 134

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CHAPTER 6 CONCLUSION The goals of this research were to develop single conical nanopore and nanotubes by the track-etch method and to investigate the potential use of these devices as biological sensors. Chapter 1 introduced and reviewed all pertinent background information for this dissertation, including the track-etch method, characteristics and properties of conical nanopores, strategies for tailoring the surface chemistry of conical nanopores, biological nanopores, and the resistive-pulse sensing method. In Chapter 2, a strategy for adding selectivity to synthetic nanopore resistive-pulse sensing devices was introduced. In this work a PEG functionalized conical nanotube was used as the sensing element. It was shown that current-pulse events from a free protein (BSA) could be distinguished from those of a protein/antibody complex (BSA/anti-BSA-Fab). Furthermore, it was shown that the BSA/anti-BSA-Fab pulses could be easily distinguished from the pulses obtained for free anti-BSA-Fab and from pulses obtained for a control protein that did not bind to the anti-BSA-Fab. Finally, it was demonstrated that the current-pulse signature for the BSA/anti-BSA-Fab complex could provide information about the size and stoichiometry of the complex. In Chapter 3, the resistive-pulse sensing of protein analytes with PEG functionalized nanotube sensors was continued. In this work three protein analytes were examined individually. The proteins were BSA (MW ~66 kDa), phosphorylase B (MW ~97.4 kDa), and -galactosidase (MW ~116 kDa). The effect of the protein size on current-pulse frequency and the current-pulse signature was investigated. The current-pulse frequency reveals information about the analyte size. Adjustment of the nanotube tip diameter was proven to be a useful tool for optimization of protein resistive-pulse sensing. The protein size was found to effect the current-pulse duration 135

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much more dramatically than the current-pulse amplitude. Current-pulse duration could prove to be a useful tool for discrimination of protein analytes, however, further optimization is needed. The spread in current-pulse duration was found to increase with protein size which could lead to significant problems with overlap of the durations of different proteins. In Chapter 4, resistive-pulse sensing of proteins at higher applied transmembrane potentials (> 1 V) was investigated as a means to optimize protein sensing and decrease the spread seen in current-pulse duration. In this work a bare PET conical nanopore was used as the sensing element. PEG functionalized nanotubes could not be used at high potentials because the PEG and underlying gold layer were removed. BSA and -galactosidase were sensed between 2-4 V applied potential. As the potential was increased, a significant decrease in the standard deviation in current-pulse duration was observed. This allowed for a solution composed of a mixture of the two proteins to be sensed and discriminated. In Chapter 5, a new type of protein sensing devices was developed from single conical nanopores in bare PET membranes. This device utilized the ion current rectification phenomenon exhibited by conical nanopores to determine the isoelectric point (pI) of proteins immobilized on the nanopore surface. Current-voltage curves of bare PET nanopores were first taken at various pHs and the extent of ion current rectification was examined. When proteins were immobilized on the nanopore surface the rectification behavior of the nanopore changed and was seen in the current-voltage curves. The change in rectification can be attributed to the charge from the immobilized protein molecules. Current-voltage curves taken above and below the pI of the immobilized protein continued to show ion current rectification due to the charge from the protein molecules. However, current-voltage curves taken at the pH corresponding to the pI of the immobilized proteins showed no rectification. This is because at this pH the surface 136

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charge on the proteins is neutralized. The protein pI values determined from the current-voltage curves differed slightly from the reported values of free proteins, and depending on the method of immobilization used. Control studies were also carried out using ethanolamine. The work presented here has demonstrated that single conical nanopores can be used in protein sensing devices. Resistive-pulse sensing of proteins using synthetic nanopores is a relatively new field. Although, further studies will be needed to continue to optimize these devices, the work presented here may hopefully serve as a foundation for this work. In this work, we have also demonstrated that it is possible to make use of other sensing paradigms with single conical nanopores. Utilizing new sensing paradigms with these devices could allow for a diverse set of sensing applications to be developed. 137

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BIOGRAPHICAL SKETCH Lindsay Taylor Sexton was born in Columbia, SC. She spent four years at the University of South Carolina, and earned a B.S. in chemistry in the fall of 2003. After graduation, she continued working as an undergraduate research assistant in the lab of Prof. Michael Myrick, at the University of South Carolina, until entering graduate school. In August of 2004, Lindsay started graduate school in the analytical chemistry department at the University of Florida, and joined the research group of Prof. Charles R. Martin. She completed her research in the spring of 2009, and obtained a Doctor of Philosophy in analytical chemistry. 149