Physiological and Microdevice Effects on Electric Field and Gene Delivery

Physiological and Microdevice Effects on Electric Field

Dissertation

Presented in Partial Fulfillment for Requirements for the Degree of Doctor of Philosophy in the Graduate School of The Ohio State University

Brian Earl Henslee

Graduate Program in Chemical and Biomolecular Engineering

The Ohio State University 2010

Dissertation Committee:

Dr. L. James Lee, Advisor

Dr. Gregory Lafyatis

Dr. David Tomasko

Brian E. Henslee

2010

Abstract

Gene therapy protocols have been actively seeking new gene delivery techniques for two decades. Successful gene therapy has carried our hope for a cure to thousands of diseases that currently plague mankind including Parkinson’s, Alzheimer’s,

Diabetes, and even cancer. Initially much effort and faith had been put into using viral vectors to deliver genes and drugs, and laboratory studies had shown viral vectors to be extremely efficient as a gene delivery technique. Although laboratory studies have been promising, safety concerns with viral vector gene delivery techniques have arisen since they were responsible for human death in clinical gene therapy trials.

Lately more attention and research is focused on physical and chemical gene delivery methods that have potential capability of high throughput, high cell viability, and high transfection efficiency. Chemical methods predominantly try to mimic a virus using polymer or lipid based systems with attached cell targeting molecules. Physical methods typically transfer genes into a cell by a physical force such as electrical or mechanical membrane disruption. Among the most attractive in physical gene delivery methods is electroporation.

Electroporation has predominantly been an in vitro delivery method. The main advantage of electroporation is in its ability to process mass quantities of cells in a very short time frame; however it has low cell viability and transfection efficiency. A modified

ii electroporation process called Nucleofection has a more successful transfection rate; however it is significantly more expensive and is unable to effectively process cells unless they are at least 106cells/ml. The main problems that hinder the use of electroporation in gene therapy are the use of very high electric fields (>1000V/cm) and non-uniform electric field distribution among cells leading to low cell viability as well as an incompatibility of this process with cell lines important to gene therapy protocols such as stem cells.

To overcome these problems we propose to reduce the overall electric field and make treatment conditions uniform for every cell in an electroporation process through the use of microdevices. We used a novel approach that employs an optical tweezers with a fluidic electroporation chip and propidium iodide dye to study the relationships between cell size and membrane breakdown, as well as cell-cell interactions and their effects on the electroporation process. Experimental results showed that each cell line has a characteristic critical electric field at which the cell begins to electroporate, and it is not dependent on the cell size. We were also able to show through both finite element analysis modeling and experimental results that cell interactions can significantly enhance or reduce the electric field and have shielding effects in certain orientations that may interfere with gene or drug delivery.

Next, we studied a microdevice that used a single micropore to trap and electroporate a cell through electric field focusing effects. Two non-cleanroom microfabrication techniques were studied and compared for quality. We found that

iii both a femtosecond laser and micromilling could produce membranes capable of trapping and electroporating cells, however the femtosecond laser was capable of producing smaller pore sizes with lower surface roughness needed for electroporation applications. Additionally we were able to show a relationship between pore size and electric field using both experimental and modeling techniques, as well as possible delivery mechanisms in micropore electroporation. Based on experimental results we found that micropore electroporation delivery is controlled by diffusion processes with enhancements from the electric field focusing effects.

The micropore electroporation device was expanded into an array of micropores to study its ability to deliver genes to mass numbers of cells. Both cleanroom and non- cleanroom fabrication methods were compared. We found that the micropore array was able to provide enhanced uniformity, reduced the electric field, and achieved high transfection efficiency through pGFP reporter gene transfection analysis when compared with bulk electroporation.

A new electroporation technique called membrane sandwich electroporation

(MSE) was developed and tested using reporter genes pSEAP and pGFP. This microdevice used two porous membranes to achieve a lower electric field that enhanced cell viability, and provided gene confinement near the cells to enhance gene delivery. Transfection results indicated an improvement over bulk and single membrane systems. MSE membranes are commercially available on a mass scale and can provide a more economically viable approach to gene delivery microdevices.

Dedication

I dedicate this work to Anneliesa for inspiring me to always be better.

Acknowledgements

I would first and foremost like to acknowledge and thank Dr. Jim Lee for his patience, guidance, and uplifting attitude throughout my doctorate studies. I am grateful for the help and advice of Dr. Greg Lafyatis, Dr. David Tomasko, Dr. Jeffery Chalmers, Dr.Yubing

Xie, Dr. Xin Hu, Dr. Shengnian Wang, Dr. Xulang Zhang, Dr. Sunny Wu, Dr. Sadhana

Sharma, Dr. Chee-guan Koh, Dr. Chunhe Zhang, and Dr. Pouyan Boukany. I would also like to thank my peers Andrew Morss, Dr. Zhengzheng Fei, Orin Hemminger, Yong Chae

Lim, Dr. Haewoon Choi, Bo Yu, Dr. Nick Ferrell, Kun-yeh Chiang, and Hyun-Chul Jung, and all other NSEC and IGERT fellows for our work together on various research projects throughout my doctorate studies.

I would also like to acknowledge and thank the National Science Foundation and OSU for their financial support through IGERT, NSEC, and University fellowships and research program funding.

A special thank you is due to my mother and father for financial, educational, and emotional support throughout my life. This would not have been possible without them.

I will always remember their encouragement to “do your best” as they sent me to school each morning.

I would lastly and most importantly thank Anneliesa for her love, encouragement, patience, and understanding throughout my graduate studies.

Vitae

December 2004……………………………B.S Food, Agricultural, and Biological Engineering

The Ohio State University, Columbus, Ohio

January 2005-present…………………Graduate Research Fellow,

Chemical and Biomolecular Engineering

The Ohio State University, Columbus, Ohio

Publications

Fei, Z.; Wang, S.; Xie, Y.; Henslee, B. E.; Koh, C. G.; Lee, L. J. (2007) Gene Transfection of

Mammalian Cells Using Membrane Sandwich Electroporation. Analytical Chemistry 79

(15): 5719-5721.

Fields of Study

Major Field of Study: Chemical and Biomolecular Engineering

Minor Field of Study: Food, Agricultural, and Biological Engineering

vii

Contents

Abstract ...... ii

Dedication ...... v

Acknowledgements ...... vi

Vitae ...... vii

List of Figures ...... xiii

List of Tables ...... xxi

Chapter 1: Introduction ...... 1

1.1 Background ...... 1

1.2 Objectives ...... 3

Chapter 2 : Literature review ...... 6

2.1 Gene therapy ...... 6

2.2 Genetic diseases ...... 6

2.3 Theory ...... 7

2.3.1 In vivo and ex vivo ...... 8

2.3.2 Gene delivery ...... 9

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2.3.3 Viral vs. Non-viral ...... 10

2.3.4 Genetic materials ...... 11

2.3.5 Electroporation ...... 12

2.4 Parameters ...... 24

2.5 Practical considerations ...... 28

2.6 Electroporation devices ...... 30

2.6.1 Bulk electroporation ...... 30

2.6.2 Microdevice electroporation ...... 33

Chapter 3 : Optical tweezers electroporation ...... 36

3.1 Introduction...... 36

3.2 Materials and methods ...... 37

3.2.1 Experimental setup ...... 37

3.2.2 Propidium iodide ...... 44

3.2.3 Cell culture ...... 46

3.2.4 Electroporation procedure for size experiments...... 48

3.2.5 Procedures for cell-cell interaction experiments ...... 50

3.2.6 Computer simulation ...... 53

3.2.7 Procedure for optical tweezers effects ...... 54

3.3 Results ...... 55

3.3.1 Cell size and electroporation ...... 55

3.3.2 Cell-cell interactions...... 62

3.3.3 Critical electric field ...... 75

3.3.4 Electroporation axis shift ...... 80

3.3.5 Optical tweezers effects ...... 86

3.4 Discussion and conclusions ...... 89

Chapter 4 : Membrane Based Electroporation ...... 97

4.1 Introduction...... 97

4.2 Single pore electroporation ...... 98

4.3 Single micropore materials and methods ...... 98

4.3.1 Single micropore experimental system ...... 98

4.3.2 Single micropore cell culture ...... 103

4.3.3 Delivery molecules ...... 105

4.3.4 Single micropore membrane fabrication ...... 106

4.3.5 Single micropore device procedures ...... 111

4.3.6 Single micropore modeling ...... 113

4.4 Single micropore results ...... 113

4.4.1 Single micropore fabrication...... 113

4.4.2 Micropore critical electric field ...... 119

4.4.3 Single pore delivery mechanisms ...... 128

4.5 Single pore discussion and conclusions ...... 134

4.6 Micropore array ...... 137

4.7 Micropore array materials and methods ...... 138

4.7.1 Micropore array experimental system ...... 138

4.7.2 Cell culture ...... 138

4.7.3 Green fluorescent protein plasmid (pGFP) ...... 138

4.7.4 Micropore array device procedures ...... 139

4.7.5 Micropore array membrane fabrication ...... 141

4.8 Micropore array results ...... 150

4.8.1 Micropore array fabrication results ...... 150

4.8.2 Micropore cell interactions and transfection ...... 152

4.9 Membrane sandwich electroporation (MSE) ...... 155

4.10 MSE materials and methods ...... 156

4.10.1 MSE experimental setup ...... 156

4.10.2 Reporter genes ...... 159

4.10.3 MSE procedures ...... 159

4.11 MSE results ...... 164

4.11.1 MSE transfection ...... 164

4.12 Array and MSE device discussion and conclusions ...... 168

Chapter 5 : Conclusions ...... 171

5.1 Conclusions...... 171

5.2 Recommendations ...... 175

Appendix A: Standard SEAP Activity Curve ...... 195

Appendix B: Software Interface for the Femtosecond Laser Motion Control ...... 196

Appendix C: Derivation of Electroporation Equations ...... 197

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List of Figures

Figure 2.1. In Vivo and Ex Vivo gene therapy protocol diagrams ...... 8

Figure 2.2. Diagram of a cell in an electric field ...... 15

Figure 2.3. Single-shell (A) and zero-shell (B) models ...... 16

Figure 2.4. Mechanistic steps in electroporation ...... 20

Figure 2.5. Illustration of a cell membrane before (top left) and after electroporation

(bottom left) with corresponding cryo-SEM images (top right, bottom right). www.inovio.com ...... 22

Figure 2.6. Parameters and shape of step and exponential electroporation program inputs...... 26

Figure 2.7. Cell cycle diagram. http://teachline.ls.huji.ac.il/72373/substance_x/cell- cycle2.jpe ...... 28

Figure 2.8.: Common commercial in vitro electroporation systems. (A) Bio-Rad Xcell™,

(B) Lonza Nucleofector™, (C) Invitrogen Neon™, (D) Molecular Devices Axoporator™

800A ...... 32

Figure 2.9. Microchannel for trapping and electroporating a cell (Khine, 2005) ...... 33

Figure 2.10. (a) A single constriction (Wang & Lu, 2006) or (b) multiple constrictions

(Hang, Schmidt, & Jensen, 2005) in a continuous flow through microchannel electroporation device ...... 34

xiii

Figure 2.11. a micropore electroporation device(Huang & Rubinsky, 2003) ...... 34

Figure 3.1. Schematic and picture of optical tweezers electropermeabilization system. A microscope image capture unit (dark), trapping laser unit (grey), and electropermeabilization unit (white) make up the three basic parts of this system...... 39

Figure 3.2. Optical tweezers laser guide equipment ...... 42

Figure 3.3. electropermeabilization fluidic chip (a) and cell location in the chip during experiments (b)...... 44

Figure 3.4. Chemical structure of Propidium Iodide (PI) dye ...... 45

Figure 3.5. Four basic configurations used in cell-cell interaction experiments. From the left they are oriented with respect to the electric field and then each other, perpendicular far, perpendicular close, horizontal far horizontal close...... 51

Figure 3.6: a)Schematic of the radial angle θ on a cell related to the direction of the electric field b) Schematic of permeabilization cuvette . Cells were held by the optical tweezers midway between the electrodes and about 20 um above the cover slip defining the lower sidewall. b) Sequence showing fluorescence of a permeabilized cell

(A) t=0, (B) t=5 s, (C) t=20 s, (D) t=60 s...... 57

Figure 3.7: Minimum electric field causing permeabilization vs. measured cell radius. ●-

K562 10 pulse,○- K562 single pulse, ♦-MES 10 pulse, ◊-MES single pulse, ▲-NIH-3T3 single pulse ...... 58

Figure 3.8: Transmembrane potential thresholds for permeabilization inferred using Eq.

1. The red lines define the range of threshold values found in almost all previous work.

Also shown are the “double Coulter counter” data of Zimmermann et al. On this graph,

xiv data fit by a line passing through the origin indicate the constant threshold electric field dependence on cell radius that we find. ●-K562 10 pulse,○- K562 single pulse, ♦-MES 10 pulse, ◊-MES single pulse, ▲-NIH-3T3 single pulse, X-Zimmerman Data ...... 61

Figure 3.9.PI entrance into a single cell during electroporation at time (a) 0s; (b) 7.5s; (c)

15s; (d) 22.5s; (e) 30s; (f) Schematics on how positively charged PI dye enters the cell. 63

Figure 3.10. Distribution of transmembrane potential for a single cell in bulk electric field: comparison of simulations and the analytical solution (dashed curve)...... 64

Figure 3.11. (a) Mesh, (b) electric field lines, and (c) distribution of transmembrane potential for the parallel case with gap 1 µm (symbol “O” is for the top cell, while “” for the bottom cell, and the “---“ is the single cell case...... 66

Figure 3.12. PI entrance in to two closely-separated cells parallel with the electric field at time (a) 0s; (b) 6s; (c) 12s; (d) 18s; (e) 30s; (f) 60s...... 67

Figure 3.13. PI entrance during electroporation of two far-separated cells in parallel case at time (a) 0s; (b) 7.5s; (c) 15s; (d) 22.5s; (e) 30s; (f) comparison of the distribution of transmembrane potential with a single cell in bulk electric field...... 68

Figure 3.14. Maximum value of transmembrane potential vs. gap between cells in the parallel case...... 69

Figure 3.15. (a) Mesh and (b) electric field lines for two cells perpendicular to the electric field with a 1 µm gap ...... 70

Figure 3.16. PI entrance during electroporation in two closely-separated cells perpendicular to the electric field lines at time (a) 0s; (b) 7.5s; (c) 15s; (d) 22.5s; (e) 30s, and (f) Distribution of transmembrane potential (symbol “O” is for the cell on the right,

xv while “” for the cell on the left, and the dashed curve is the single cell in bulk electric field)...... 71

Figure 3.17. PI entrance during electroporation of two far-separated cells in perpendicular case at time (a) 0s; (b) 6s; (c) 12s; (d) 18s; (e) 24s; (f) 30s; (g) Distribution of transmembrane potential (symbol “O” is for the cell on the right, while “” for the cell on the left, and the dashed curve is the single cell in bulk electric field)...... 73

Figure 3.18. Maximum transmembrane potential vs. gap for cells perpendicular to the electric field lines...... 74

Figure 3.19. Electroporation membrane breakdown threshold for two K562 cells oriented parallel or perpendicular to the electric field and a single k562 cell...... 77

Figure 3.20. Dominant cell in electroporation for two cells parallel to the electric field 80

Figure 3.21. (a) 3D distribution; (b) Projection on X-Z plane of transmembrane potential;

(c) Distribution of transmembrane potential (symbol “O” is for the cell on the right, while “” for the cell on the left, and the dashed curve is the single cell in bulk electric field) continued...... 82

Figure 3.22. Perpendicular close (a) and perpendicular far (b) polar intensity plots, perpendicular close (c) and perpendicular far (d) model predicted electric field lines, and perpendicular close (e) and perpendicular far (f) images with defined angle convention.

...... 85

Figure 3.23. A K562 cell exposed to trypan blue before (A) and after (B) electropermeabilization compared with a dead cell (C) exposed to trypan blue dye. .... 86

xvi

Figure 3.24. relative intensity values for a K562 cell exposed to trypan before and after electropermeabilization compared to a dead K562 cell...... 87

Figure 3.25. Effects of trapping laser on 3T3 electroporation. Data noted with “surface” indicate cells were attached to the surface...... 89

Figure 4.2. The confocal electroporation system ...... 99

Figure 4.3. Diagram and photo of the microfluidic chip used for membrane interfacing

...... 102

Figure 4.4. Femtosecond Laser System (Farson, et al., 2008) ...... 108

Figure 4.5. Thermal and surface effects of long wave (a) vs femtosecond laser (b) pulsing (http://www.cmxr.com/Industrial/Handbook.htm) ...... 109

Figure 4.6. SEM images for the 10µm milling bit (a) front and (b) back of the membrane

...... 115

Figure 4.7. Femtosecond laser microfabrication results. (A) large and small opening vs laser power (B) membrane cross sectional diagram (C) brightfield images (D)SEM micropore images of the front, and (E) SEM images of the back of the membranes .... 117

Figure 4.8. A calcian stained cell interacting with a single micropore ...... 119

Figure 4.9. K562 critical electric field using micropore membranes ...... 121

Figure 4.10.Comparison of micropore membrane electric field with bulk electroporation for NIH-3T3 cells ...... 122

Figure 4.11. (a) Electric field distribution and (b) transmembrane potential plot for the 4 micron pore with the cell located on the membrane surface ...... 124

xvii

Figure 4.12. (a) electric field lines and (b) transmembrane potential for the 10 micron pore with cell located on the membrane surface ...... 125

Figure 4.13. (a) electric field lines and (b) transmembrane potential for the 12 micron pore with cell located on the membrane surface ...... 126

Figure 4.14. (a) electric field lines and (b) transmembrane potential for the 4 micron pore with the a 500 nm gap between the cell membrane and the pore edge ...... 127

Figure 4.15. Bulk and micropore PI fluorescence curves after electroporation at (t=0) 129

Figure 4.16. Confocal images of PI dye delivery to a K562 cell at 10 seconds post electroporation from the a) top, b) front, and c) side and 20 seconds post electroporation from the d) top, e) front, f) side...... 130

Figure 4.17. ODN delivery through the 4 micron pore immediately after and 10 minutes after electroporation...... 132

Figure 4.18. Fluorescence images of YOYO-1 stained pGFP delivered to a cell in bulk electroporation a) before electroporation, b) after electroporation and confocal images of yoyo-1 stained pGFP delivery to a cell using a micropore 15 minutes post electroporation from the c) bottom, d) side, and e) front of the cell. The white circle indicates the approximate location of the cell, and the squares indicate the approximate location of the micropore membrane...... 133

Figure 4.19. microfluidic used for micropore array membrane interfacing ...... 140

Figure 4.20. spin speed vs. SU-8 25 photoresist thickness ...... 143

Figure 4.21. (a) Top and (b) 45 degree SEM of the developed SU-8 25 photoresist ...... 145

Figure 4.22: Soft lithography micropore array membrane fabrication diagram ...... 146

xviii

Figure 4.23. Broken micropillars in PDMS molding ...... 147

Figure 4.24. Mold replication with modified processing conditions ...... 147

Figure 4.25. Membrane thickness vs spincoat speed (rpm) ...... 148

Figure 4.26. Schematic and light microscopy images of (A) incomplete (3000rpm) and (B) complete (5000rpm) processing conditions for PCL membrane fabrication...... 149

Figure 4.27. (a) Bright field image of a femtosecond laser micropore array membrane and (b) a soft lithography micropore array membrane. (c) Bright field image of a femtosecond laser micropore array membrane with cells trapped and (d) a soft lithography micropore array membrane with cells trapped...... 151

Figure 4.28. (a)array detail and (b) pore detail SEM images of a femtosecond laser fabricated micropore array membrane and (c) array detail (45 degree) and (b) pore detail SEM images of a soft lithography fabricated micropore array membrane...... 152

Figure 4.29. Diagram of cell interaciton with micropores ...... 153

Figure 4.30. 3D images of 3T3 cells interacting with the PCL micropore array ...... 154

Figure 4.31. pGFP expression 48 (A) and 24 (B) hours after electroporation ...... 155

Figure 4.32. Experimental system (a) and fluidic platform (b) for membrane sandwich electroporation (MSE) ...... 157

Figure 4.33. Schematic of (a) MSE membrane disk and (b) DNA migration during electroporation ...... 161

Figure 4.34. Schematic of the polymer stamping technique to produce spacer membranes for MSE ...... 162

Figure 4.35. Polystyrene spacer bonded with a PET track etched membrane ...... 163

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Figure 4.36. pGFP expression comparison using (a) bulk (b) local bottom (c) local top and

(d) MSE ...... 166

Figure 4.37. Comparison of SEAP gene activity between MSE and local electroporation

...... 167

Figure 4.38.pGFP expression in spacer MSE vs standard MSE ...... 167

Figure 4.39. pGFP expression in standard MSE vs ordered array local electroporation 168

Figure 5.1: nanoporous polymer membrane from sacrificial template imprinting (Wang

S. , 2006) ...... 178

Figure 5.2. Two microchannels connected by a nanochannel (P. Boukaney, et al., 2010)

...... 178

Figure 5.3. Electrical model of single cell electroporation (Bandiera 2007) ...... 180

Figure 5.4. Impedance and phase dependence on frequency before and after electroporation ...... 182

List of Tables

Table 2.1. Commercial electroporation systems comparison ...... 31

Table 3.1. Average critical electric fields with standard deviations ...... 59

Table 3.2. Optical tweezers laser effects ...... 88

Table 4.1. Square wave generator technical specifications ...... 158

Table 4.2. Comparison of bulk, localized, and MSE electrical parameters ...... 160

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Chapter 1: Introduction

1.1 Background

Gene therapy is the process of treating disease through introduction of genetic material to cells (Klug, 2003) (Mulligan, 1993). It holds the promise of being able to treat genetic diseases as well as cancer (Roth, 1997). The ability to successfully deliver a gene to a cell has been recognized as the single most difficult part of a gene therapy process

(Mulligan, 1993) (Niidome, 2002). For us to fulfill its promises, we must better understand gene delivery mechanisms and processes.

Genetic diseases

Genetic diseases affect millions of people with no cure at the current time.

Genetic diseases are either inherited from parents and passed on through the DNA or caused by a mutagen that alters a person’s DNA (Genes and Gene Therapy: Medline

Plus, 2008). A person with a genetic disorder is incapable of producing functioning proteins needed for normal body function (Genetic Disease Information, 2008). Among common genetic diseases are color blindness, cystic fibrosis, sickle cell anemia, hemophilia, muscular dystrophy, and Tay-Sachs to name a few, however, nearly 6,000 genetic diseases currently exist. In addition to traditional genetic diseases it is now

1 believed that forms of cancer can be included in genetic diseases or treated using gene therapies (Freeman, 1993). Current treatments have only been capable of correcting symptoms of these diseases. This is primarily due to the lack of technology capable of correcting genetic defects which are the underlying cause of these diseases.

Clinically relevant cells

There are many different cell types within the human body that the medical research community has had difficulty in targeting and using in gene therapy treatments. All cells in the human body come from a primordial cell type called an embryonic stem (ES) cell. ES cells have unlimited self-renewal and pluripotent capabilities enabling them to differentiate into any specific cell types such as blood, skin cells, or neuron cells making them attractive for use in gene therapy treatments

(Turgeman, 2001). Many cancerous cells are genetically defective human cells that grow and divide rapidly to form tumors (Kikuchi, 2003) and could potentially be cured through gene therapy. Cancer and Stem cells are among the most clinically relevant cells to study in gene delivery for the severity of and range of diseases they may be able to cure using gene therapy. However, it has been very difficult to introduce corrective or regulative genes in to such cells with high efficiency and viability.

Electroporation

Electroporation is a gene delivery technology that has potential to be used in gene therapy and has recently received attention for use with ES cells (Tompers, 2004).

Traditional electroporation techniques have shown the ability to deliver large gene 2 constructs (Shent, 1995) which can be advantageous for gene therapy. However, traditional electroporation has had low transfection efficiency (<20%) coupled with low cell survival rates (<50%) which are both unacceptable for a viable gene therapy protocol. ES cells have been successfully transfected with new electroporation technology (Siemen, 2005), (Lorenz, 2004) but low cell survival rates and efficiency remain a place for improvement. In current electroporation methods each cell experiences different local process conditions due to non-uniform electric fields, and high voltages. This leads to a degree of cell death and untransfected cells in every process.

1.2 Objectives

If all cells experience the same electroporation process conditions, this process can be optimized to achieve a high transfection efficiency and cell survival rate. We propose to use novel micro and nanofabrication methods to achieve uniform process conditions for in vitro gene delivery. We will use these fabrication techniques to develop devices to achieve a uniform electric field at a relatively low level, and as a result, we can increase cell survival and overall transfection efficiency.

Fundamentals

We tested theoretical relationships between cell size and critical electric field, and how interactions between cells influence the electroporation process. We used novel techniques that employed optical tweezers and a custom electroporation device to provide consistently reproducible single cell experiments. Our experiments included

3 three different cell types representative of ES cells, cancer cells, and general animal cells. We were able to show a clear relationship between cell size and critical electric field and how bulk electroporation processes can generate non-uniform process conditions through simple interactions between cells.

Micropore

We next developed microporous membranes ranging from a single pore to organized arrays of micropores. We incorporated them into a custom electroporation microfluidic device. In the single pore work, we clearly demonstrated the effects of pore size on electroporation process conditions. In the organized array work, each cell was located on a single pore and enabled each cell to experience the same process conditions. We used high speed confocal microscopy to understand how delivery mechanisms in micropores are different from bulk electroporation. The micropores focus the electric field and confine electroporation to smaller portions of the cell membrane. The focused electric field enabled us to reduce voltage used in the process, and accelerated uptake of delivery materials.

Membrane sandwich electroporation

We have also developed an original electroporation method called Membrane

Sandwich Electroporation (MSE) to provide better control of the electric field in electroporation. MSE uses micro-porous membranes to focus the electric field and confine DNA around the cells. The focused electric field enabled a lower voltage to be used increasing cell viability while DNA confinement enhanced gene uptake. Although 4 the MSE approach was more uniform than traditional electroporation processes, incorporated membranes with a random pore distribution still creates non-uniform process conditions.

Chapter 2 : Literature review

2.1 Gene therapy

Gene therapy is the process of treating a disease through the use of genetic material. Since it has the potential to treat thousands of diseases that currently have no cure it has received intense research attention over the past 20-30 years. Researchers have been mapping the human genome since the early 1980’s and began the first clinical gene therapy trial in 1990 (Gene Therapy, 2008), however much more needs to be done.

2.2 Genetic diseases

A genetic disease in its broadest sense is an abnormal condition resulting from an abnormal genome. Variations in genes arise naturally within populations from generation to generation depending on environmental conditions and is often studied in wild drosophila (Henslee, 1966). Natural genetic variation can result in a dysfunctional protein that is essential to healthy body function causing a genetic disease to arise.

Genetic diseases come in different forms, each necessitating a different gene therapy

6 strategy. A monogenic disease is caused by a mutation in a single gene. There are over

6000 monogenic diseases including cystic fibrosis, sickle cell anemia (Genetic Disease,

2010). A polygenic disease results from mutations on two or more genes and is often coupled with environmental factors before causing health problems. Common examples of polygenic diseases include heart disease, Alzheimer’s, and cancer. Chromosomal diseases are the result of an abnormal set of chromosomes. In Down syndrome an extra chromosome is present, and in other chromosomal diseases a chromosome may be missing a part, lack normal structure. An organelle within human cells called mitochondria also has its own DNA that can be affected by mutations and cause human mitochondrial disease.

2.3 Theory

Past clinical approaches to these diseases have sought to treat the symptoms since no cure has been available. However, since researchers have developed a better understanding of the human genome coupled with the development of recombinant

DNA technology new theories on how to treat genetic diseases have given rise to gene therapy. The basic theory behind gene therapy is that one can correct genetic disease with the use of genetic material. Although this sounds straightforward, the differences and relative complexities of each genetic disease require different strategies to achieve a cure through gene therapy.

2.3.1 In vivo and ex vivo

Gene therapy can be accomplished through in vivo or ex vivo treatments (Figure

2.1). In vivo gene therapy is a technique that treats cells inside the body. In this method a doctor administers viral or engineered gene delivery particles that bind specifically with a cell in need of gene therapy (Thomas, 2003). Viral or engineered particles can cause immune response and toxicity (Niidome, 2002). Researchers have tested other gene delivery methods and have increased safety standards for viral vectors in vivo treatments.

len.epfl.ch

biochem.arizona.ed Figure 2.1. In Vivo and Ex Vivo gene therapy protocol diagrams u

Ex vivo treatment techniques remove cells from a patient for gene delivery outside the body. Genetically modified patient cells are then reintroduced back into the areas that need treatment (Mulligan, 1993). Doctors primarily use electroporation or lipofection to deliver genes to the cells (Nishi, 1997). Cells used in ex vivo treatments are not necessarily put back where they came from as they may be capable of adapting into other cell types when reintroduced (Schindhelm, 1999; Ashara, 2000). An important example of a cell of this nature is an ES cell.

2.3.2 Gene delivery

Gene delivery is the single most important area for improvement in gene therapy protocols (Mulligan, 1993; Mountain, 2000). Although, there are many technologies capable of transecting cells, none meet the rigorous demands of gene therapy. The single most important aspect of a gene delivery technology is that it does not harm a patient. It must also have a very high transfection efficiency to generate a treatment protocol that has a significant impact on patient health. Depending on the disease it must also be specifically targeted to a type of cell that needs the treatment, or delivery a relatively large piece of genetic material.

2.3.3 Viral vs. Non-viral

A genetically modified virus is the basis for viral methods of gene delivery. A virus operates as a gene delivery tool in nature and this delivery method takes advantage of their highly specialized machinery to accomplish this task. A virus can be very specific and efficient in their delivery and almost presents the perfect delivery tool for gene therapy. They have been used in clinical trials since 1990 (Blaese, 1995), however there are problems with viral gene delivery. Viruses stimulate a significant host immune response (Thomas, 2003) and have caused death in human trials (Hollon, 2000).

In addition to these treatment problems viruses are also costly to produce as a gene delivery technique and there is concern that viruses may recover their natural disease causing abilities once in the body.

Non-viral gene delivery has become more popular for its cost effectiveness and safety. Non-viral gene delivery methods fall into two categories; physical and chemical methods. Chemical methods primarily seek to mimic elements of a virus (Balicki, 2002).

Chemical methods generally coat DNA with polymeric or lipid based materials and may or may not contain specific cell binding and targeting agents (Dauty, 2001; Whitman,

2001). Researchers are primarily concerned with the high toxicity associated with the materials used to coat DNA in these chemical methods (Mehir-Humbert, 2005). Physical delivery methods include micro-injection, electroporation, gene-gun, sonoporation, and laser irradiation. These techniques directly introduce genetic material into cells and avoid secondary effects seen in viral or chemical delivery methods. The majority of

10 physical methods can be time consuming and destructive to the cells being treated and lack desired throughput compared to chemical and viral vectors. As a result many physical methods are not used in gene therapy. Of the physical delivery methods electroporation is the most practical and has been used for in vivo and ex vivo gene therapy studies (Niidome, 2002; Oshima, 1998; Miklavcic, 1998).

2.3.4 Genetic materials

There are many different genetic materials that can be used in gene therapy since it is a broad term covering any treatment using genetic material. The type and form of genetic material used in gene therapy depends on the type of treatment.

Traditionally gene therapy protocols sought to use large (1-100kbp) strands of DNA that would be delivered and incorporated into a patient’s genome somewhere to produce a required protein. These DNA strands could be linear or circular, single or double stranded (Mountain, 2000). In addition to using DNA, RNA is also a genetic material that can be used in gene therapy (Mountain, 2000; Zhang, 2004).

Genetic material can also be added to a cell to correct a genetic disorder without being incorporated into the patient’s genome. Researchers have found that one can use small pieces of interfering RNA (siRNA) to silence unwanted gene expression. siRNA is in the 10’s of base pair range and come from degradation of double stranded RNA pieces of 100-300bp length (Tuschl, 2002). siRNA affects gene expression by combining with a dicer complex to target specific mRNA effectively silencing transcription of a gene into

11 protein. Plasmids can be used to correct a genetic disorder without being incorporated into a patient’s genome. A plasmid is a circular form of DNA that acts as a stand-alone gene, outside of chromosomal DNA, capable of producing a needed protein or performing other common genetic functions (Siemen, 2005).

2.3.5 Electroporation

Electroporation has become an important gene delivery tool for gene therapy because it has the potential to offer optimal conditions for some gene therapy protocols. In particular it has great potential for ex vivo protocols, and use with ES cells

(Mountain, 2000; Siemen, 2005; Lorenz, 2004; Tompers, 2004). There is a renewed interest in electroporation research to develop more efficient transfection methods to meet the demands of gene therapy.

Theory

Electroporation, sometimes called electropermiabilization, relies on a pulsed electric field to cause transient openings on a cells membrane (Weaver J. , 1993),

(Chernomorodic, 1987; Gabriel, 1997). The cell membrane is made of a lipid bilayer and pores from electroporation are nanoscale (Chang D. C., 1992). These openings permit diffusion across the membrane that would not normally occur. Scientists regard electroporation as one of the most reliable and effective methods for mass transport of large genes and drugs into cells. Other gene delivery techniques are used in situations

12 where electroporation has been ineffective or is inapplicable. Despite its widespread use molecular mechanisms of electroporation are still not well understood (Neumann,

Kakorin, & Toensing, 1999; Teissie J. G., 2005).

Electroporation can be used in a variety of different ways in cell biology. It is most well known as a gene delivery method for animal cells, plant cells, bacteria, and fungi. Although electroporation has not been successful with all cell types in these categories, it is applicable to a majority of them. Electroporation can also be used to delivery other drugs or large macromolecules into cells in the same way it is used to deliver genes. Electroporation techniques can be used in vivo (Haas, 2001) or in vitro

(Jaroszeski, 1999); however they are most common in vitro. A more obscure use of electroporation is for the insertion of proteins into the cell membrane (Chang D. C.,

1992). Electroporation has also been used in the production of monoclonal antibodies in a process called electrofusion to join two different cell types (Wang J. L., 2006).

Electroporation pulses usually come from a DC power source and can take on a variety of wave forms including square wave inputs, bimodal square inputs, triangular inputs, positive followed by negative inputs, exponential inputs, ect. The most common among these are the exponential and square waves (Chang D. C., 1992; Bio RAD, 2008).

The electric pulse establishes an electric field around the cells through their liquid culture media. The electric field strength (E) depends on the distance between the electrodes (l) and the voltage output (V).

V E  (1) l 13

Cross sectional area changes between the electrodes can also influence the electric field in that the electric field will increase inversely proportional to the change in the cross section. Researchers have recently been using this relationship to design new electroporation devices.

E1 A1  E2 A2 (2)

When electroporating a cell, the membrane field strength ( Em ), which is dependent on the outside electric field strength, determines when a cell membrane will break down and electroporate.

  E  m (3) m d

d where m is the transmembrane potential and represents the dielectric membrane

thickness (~5nm). m can be further decomposed into its induced i , surface s 

, and natural n  components. Surfaces , and natural n  components have a

finite value dependent on the cell and it has been shown that n  commonly takes on a value around -70mV (Cevc, 1990). The induced membrane potential is most influenced by electroporation pulses. If the cell is spherical with a shape factor of 1.5, the induced membrane potential is represented by (DeBruin, 1999):

t    1.5rEf () cos() (1 e  ) (4) i

where r is the radius of the cell, E as mentioned previously is the external electric field strength, f () is a conductivity factor dependent on the media (usually 1),  is the angle between the electric field and specific point on the membrane (Figure 2.2), t is the time that E is enabled, and  is the charging or relaxation time of the membrane.

Therefore the overall transmembrane potential is represented by (Neumann E. B.,

1989):

r E θ

Anode (+) Cathode (-)

Figure 2.2. Diagram of a cell in an electric field

 t      s  n  m  1.5rEf ()(1 e )   cos() (5)  cos()   

As the transmembrane potential increases it reaches a critical transmembrane potential and the cell membrane will break down as ions will flow in and out of the cell to reduce the potential. With the proper electrode placement single cell electroporation can be achieved with less than 1 volt (Khine, 2005; Olofsen, 2003).

A B

Figure 2.3. Single-shell (A) and zero-shell (B) models

There are two common models used to analyze the transmembrane potential of a given system: the first one is called a zero shell model (Figure 2.3B) assuming the potential distribution across the cell membrane is linear since the membrane thickness is negligible, however this approach doesn’t solve the Laplace equation for the cell

16 membrane. To consider the current density flow across the membrane a single-shell model (Figure 2.3 A) was developed. Its governing equations and boundary conditions are given as follows:

( ii )  0 (6)

( ee )  0 (7)

   i   e  (  ) / d (8) i n e n e i m

where i and e are intracellular and external potentials,  i and  e are intracellular

and external conductivities and (e i ) m / d is the current density flow across the cell membrane. Once the external and inner potentials are solved, the transmembrane

potential m  e (R) i (R) on the cell membrane can be obtained. This approach has been used to solve the distribution of transmembrane potential of a single cell under an electric field through a micro-capillary (Zudans, 2007).

The other approach is to solve the single-shell model numerically. That means we need to additionally solve the electric field on the cell membrane. That is

( mm )  0 (9)

In this model, there are two interfaces: one is the external interface between the external medium and the cell membrane; the other is the inner interface between the

17 cytoplasm and the cell membrane. Thus two boundary conditions need to be considered:

   e   m ,    on the external interface (10) e n m n i m

   i   m ,    on the inner interface (11) i n m n e m

Once all the variables have been calculated, the transmembrane potential (the potential difference between the external and inner interfaces) can be obtained.

   (R)  (R  d) m m m (12)

These two approaches can be extended to any multi-cell system. Overall, the second approach is more complicated than the first one, but it is more general and can be used in the case when the cell is electroporated, e.g., there are nanopores formed on the cell membrane.

Successful electroporation optimizes parameters so that the pores are reversible. If the field strength is too high or if the duration of pulses is too long it will damage the cell beyond its ability to repair the membrane and the cell will eventually or immediately die due to loss of intracellular contents. A cell is able to seal up electropores when done successfully using its natural cellular machinery (Benz R. B.,

1979; Hapala, 1997).

Mechanisms

Although these fundamental equations allow us to begin to understand cell membrane breakdown in electroporation there are still many unanswered questions regarding mechanisms that take place during and after membrane breakdown (Farvard,

2007). The entire electroporation process can be thought of as a number of steps seen in Figure 2.4. Since electroporation mechanisms are not fully understood this is only a proposed theory of a stepwise mechanism based on the field’s best scientific studies and analysis.

Cell membrane Electropore

breakdown expansion

Membrane Membrane

restructure stabilization

(recovery) Membrane

resealing

Figure 2.4. Mechanistic steps in electroporation

At the beginning of the mechanism is membrane breakdown. The aforementioned fundamental equations (1)-(5) help describe the cell membrane breakdown step of this process. The asymmetric cos(θ) dependence in equation (5) has been demonstrated in experimental results (Hiubino, 1991). Energy from electroporation charges the cell membrane like a capacitor until it reaches a critical potential (Babakov, 1966; Wobshall, 1966). Once a critical transmembrane potential is reached the membrane begins to breakdown. Membrane breakdown includes the destabilization of the cell membrane lipid bilayer up to the point where it begins to form electropores. The shape of electropores is thought to be circular due to its minimum

20 energy state and the minimum theoretical diameter of the electropores at this point is around 1 nm (Weaver J. C., 1996). Molecular simulation of this step suggest a dipole forces between water molecules play a primary role in pore formation and that pores begin to form in a few nanoseconds after an electric field is applied (Tieleman, 2004).

Electropore expansion covers the growth in size and shape of the electropores.

Fundamental understanding of when the membrane breaks down is the most understood part of electroporation; however we lack knowledge of how and to what extent the pores expand. Some cryogenic electron microscopy experiments have attempted to visualize electropores with some success. Both flap like geometry

(Stenger, 1986) and volcano like geometry (Chang D. a., 1990) have been reported to show pores 20-120 nm (Figure 2.5). These sizes contradict smaller sizes proposed by analytical methods (Neumann, Kakorin, & Toensing, 1999) and cryogenic processes take on the order of milliseconds therefore these images are at best what would be observed through osmotic swelling of the initial pores formed from electroporation and at worst an artificial defect caused by the cryogenic process. The best evidence we have of when the pores form and grow has been through observation of increased electrical current flow through a lipid membrane (Hibino, 1993; Zimmermann, 1974).

Figure 2.5. Illustration of a cell membrane before (top left) and after electroporation (bottom left) with corresponding cryo-SEM images (top right, bottom right). www.inovio.com

After the electric field becomes sub-critical the membrane stabilizes and transmembrane diffusion takes place as pores reseal. This part of the process has traditionally been detected by a strong decrease in detected current through the membrane. Small molecule transport through the membrane during pore expansion and stabilization has been described using Fick’s laws of diffusion (Neumann, Kakorin, &

Toensing, 1999; Rols M. a., 1990; Neumann E. K., 2000). Osmotic swelling of the cell takes place during the time the pores form until they close (Tsong, Electroporation of

22 cell membranes, 1991), (Tsong, 1992). The overall length of this process can be anywhere from a fraction of a second up to a few minutes (Weaver J. , 1993).

Mechanisms of gene delivery have been proposed to follow one of two mechanisms; electrophoresis driven or diffusion driven uptake. In an electrophoretic driven uptake mechanism, genetic material is driven to the surface of the cell membrane near the pores by the electric field and enters the cytoplasm through endocytosis (Klenchin, 1991; Sukharev, 1992). In diffusion driven uptake, the genetic materials behave like other small molecules and enters the cell primarily through diffusion driven processes (Sukharev, 1992). This type of uptake mechanism is most appropriate for small pieces of genetic material like siRNA or microRNA. Uptake mechanisms have been observed in both in vivo (Mir, 1999) and in vitro studies (Golzio,

2002). DNA has been shown to enter a cell predominantly at the cathode side suggesting that electrophoretic forces do play a significant role in the uptake process

(Faurie, 2004).

The final step of the process is membrane memory. The cell membrane is not just a simple lipid bilayer, it is composed of a fluid mosaic of lipids, cholesterol, proteins, and carbohydrates. The electroporation process strongly disrupts the physiological function of common membrane processes and organization (Dressler, 1983). Although the membrane has resealed its electropores at this point it takes hours to fully reestablish membrane function (Glougaure, 1993). Once a cell has fully recovered it is capable of enduring the process again.

Primary sources of information about these processes are measurements of secondary phenomena. We have yet to visualize pore formation and resealing. We do not have definitive evidence of gene transport mechanisms or how nucleic acids or drugs interact with pores. We do not understand the processes a cell must undergo to reseal its pores or reorganize its membrane. Many of the theoretical equations and models we have used to characterize electroporation have yet to be experimentally accounted for and verified.

2.4 Parameters

To achieve electroporation one must make decisions on parameters that are under control. Typical variable parameters in electroporation include voltage, pulse shape, pulse length, pulse frequency, pulsing duration, pulsing buffer composition, cell density, and stage of cell life cycle. Electrical parameters are typically controlled using a specialized electrical power supply, while chemical parameters are dependent on formulation. Each specific cell type requires different optimized settings to cause electroporation. For instance an ES cell does not have the same critical transmembrane potential as a fibroblast cell and may require a higher or lower electric field to cause electroporation. The goal of parameter setting in electroporation is to optimize settings to provide the best transfection at the highest cell viability.

An electroporation power supply is used to control electrical pulsing parameters of the electroporation process. The pulse voltage is possibly the most critical setting

24 since it is the variable that controls the strength of the electric field. The pulse voltage must create an electric field that is over the threshold value needed to cause electroporation otherwise known as a critical transmembrane potential (Δφc). The voltage setting dictates Δφ as described in equation (5). It also must be low enough so it does not cause irreversible electroporation leading to cell death or cell rupture. An increase in voltage causes a proportional increase in electric field as described in equation (1).

Pulse shape, length, frequency and duration are also controlled with the electroporation power supply. Although the variety of possible electrical pulse wave forms is endless there are predominantly two pulse forms that are used in electroporation; step inputs and exponential decrease. A step input is simply a step change in voltage in the positive or negative direction, while an exponential decrease starts at a peak and decays exponentially from that point (Figure 2.6). Exponential inputs usually are a single input and take the shape of a capacitor discharge. The initial peak amplitude of the exponential pulse is set to cause electroporation while the subsequent voltage decay acts to keep electropores open and drive DNA or other genetic material into the cell through electrophoresis. In an exponential pulse peak amplitude and an exponential decay time constant are selected to program the pulse.

Step inputs have a pulse voltage amplitude to cause pores to form and usually have an overall pulsing duration to enlarge and keep the pores open. Throughout the pulsing duration there can be multiple pulses depending on the pulse frequency. Each pulse also has a specified pulse length corresponding to how long each individual pulse is applied.

A single long constant pulse can cause a cell to rupture before transfection takes place.

To counteract this dilemma multiple short pulses are used to allow the cells to have a

short membrane recovery period between pulses with a long total pulse exposure

needed for adequate diffusive and electrophoretic flux. Amplitude, frequency, duration

and length parameters work in concert to optimize transfection for each cell line.

Step Exponential

duration frequency length

Voltage

amplitude

Time Figure 2.6. Parameters and shape of step and exponential electroporation program inputs.

Electroporation buffer is also another factor that can influence the outcome of

electroporation. The presence of ionic salts allows current to flow through the

electroporation buffer, however a low ionic strength is preferable. High ionic strength

increases current flow and can more rapidly cause joule heating and cell death (Rols M.

a., 1990; Wang H. L., 2006). Apart from electrical considerations from ion content there

26 are ions that can promote cell health, membrane healing, and transfection. Calcium ions

(Ca2+) can be added to aid in membrane recovery, while magnesium ions (Mg2+) can be added to help cell membrane recovery as well as increase transfection rate (Tsong,

Electroporation of cell membranes, 1991; Klenchin, 1991).

Other buffer content additions have aided in cell recovery post electroporation.

Chemical imbalances between the cytoplasam of a cell and its external environment can cause osmotic swelling leading to cell rupture and loss of critical ion balances needed for cell survival. Additions of sucrose have been shown to enhance cell viability by reducing some of these effects while at the same time promote better transfection (Myers,

2003). Beyond sucrose different surfactants including polyoxyethylene glycol, poloyxmer, peptides, and dextran have been added to electroporation buffers to successfully aid in cell recovery (Kanduser, 2003; Tsung, 1999).

Transfection success in electroporation is also influenced by the physiological state of the cell. It has been shown that gene delivery is most successful when cells are rapidly growing and producing new cells in what is known as logarithmic growth

(Anderson, 1991). Cells have a natural life cycle where they grow and divide to form new cells called the cell cycle (Figure 2.7). Logarithmic growth takes place when a cell is operating in the mitotic (M) phase of the cell cycle while it is dividing, when a cell is not in the growth cycle it is in what is called G0 phase or a no growth phase. Transfection is more successful when the cell is dividing in the mitotic phase (M) since this is when genetic material is copied and divided (Golzio, 2002). In mitotic phase access to the cell

27 nucleus is at its peak and it offers the best conditions for incorporation of new genetic material through gene delivery.

Figure 2.7. Cell cycle diagram. http://teachline.ls.huji.ac.il/72373/substance_x/cell- cycle2.jpe

2.5 Practical considerations

Electrode selection and design is one aspect of electroporation that does not receive much attention, however it plays an important role in electroporation devices.

There are a variety of materials that can conceivably be used in an electroporation

28 device. Most systems are made with metal electrodes for cost, availability and manufacturability however conductive polymer and graphite materials can work as well.

In a disposable or one time use device it is possible to use metals like aluminum or copper, however if these types of material are used long term they are prone to oxidation. Another consideration when using non-noble metals in long pulse situations is that the electrodes will dissolve away over time due to electrodeposition processes

(Mantell, 1960). For long term electrode use the most cost effective material may be stainless steel, however noble metals such as gold, platinum, or palladium may work well also.

Another practical consideration that must be accounted for in electroporation processes is electrolysis. In electrolysis a DC current between electrodes can cause hydrogen gas formation (Engelhardt, 1904). Electroporation pulses often generate gas bubbles in the electroporation media near the electrode surfaces. In traditional bulk electroporation a foamy froth from hydrolysis is present at the top of an electroporation cuvette after electroporation. These bubbles cause reduction in cell viability when they rise to the surfaced and pop near surrounding cells (Garcia-Briones, 1994; Cherry, 2008).

Different pulsing programs can be used to minimize the electric field and pulse duration effectively reducing hydrolysis.

Temperature during electroporation can influence cell viability. Electroporation pulses add energy to a cell suspension and as a result cause joule heating in the cell suspension. If too much joule heating occurs the resulting high temperatures can

29 denature crucial cell proteins effectively cooking and destroying cells. A common practice in electroporation is to cool a cell suspension prior to electroporation to counteract joule heating effects. In addition to cooling it is also recommended to minimize the pulsing lengths and duration to reduce joule heating effects. After electroporation it is common to incubate animal cells at 37oC to enhance viability and transfection efficiency.

2.6 Electroporation devices

2.6.1 Bulk electroporation

Bulk electroporation is the traditional in vitro electroporation process that is in wide use throughout the world. It involves a suspension of cells, a cuvette with two electrodes and a power supply. In most commercial devices cuvette or 96 well shuttle is made of a polymer with integrated aluminum electrodes and is intended for one time use. The leading traditional electroporation systems have both single cuvette and 96 well shuttle applications. There are also some specialty systems intended for single cell experiments.

Table 2.1 summarizes the most common commercial systems and they are pictured in

Figure 2.8.

Table 2.1. Commercial electroporation systems comparison

System Company Format Advantage Disadvantage

Gene Pulsar Xcell Bio-Rad single cuvette program flexibility low efficiency

or Mxcell 96 well plate

Nucleofector Lonza single cuvette efficiency Program flexibility specialized 96 well plate reagents

Neon Invitrogen pipette tip efficiency Low throughput

program flexibility

Axoporator 800A Molecular Devices single cell micropipette single cell targeting Limited throughput

A B

C D

Figure 2.8.: Common commercial in vitro electroporation systems. (A) Bio-Rad Xcell™, (B) Lonza Nucleofector™, (C) Invitrogen Neon™, (D) Molecular Devices Axoporator™ 800A

There are a few in vivo systems on the market as this has been a recent topic of research (Liu, 2002; Heller, 2000). In Vivo electroporation is a complex process compared to in vitro. Depending on the location of delivery a new set of parameters must be accounted for. The electrode shape and design must be different depending on the site of delivery. The electroporation parameters must change depending on the density of tissue. There are still many unknown aspects of electroporation that prohibit widespread in vivo protocols.

2.6.2 Microdevice electroporation

Micro/ nanofabrication and lab on a chip (LOC) technologies have given way to new ways to study and implement electroporation. Microchannels have been used in multiple ways to enhance the electric field to cause electroporation or cell lyses. At U.C.

Berkley Luke Lee’s group has demonstrated a lab on a chip device that implements microchannels to trap cells and cause localized electroporation on the cell membrane trapped inside the microchannel (Figure 2.9). Other microchannel systems implement a single or series of constrictions to create a temporary increase in electric field while the cell is in a constricted area (Figure 2.10). A micropore (Figure 2.11) can accomplish essentially the same results as a microchannel through trapping and focusing the electric field.

Figure 2.9. Microchannel for trapping and electroporating a cell (Khine, 2005)

Figure 2.10. (a) A single constriction (Wang & Lu, 2006) or (b) multiple constrictions (Hang, Schmidt, & Jensen, 2005) in a continuous flow through microchannel electroporation device

Figure 2.11. a micropore electroporation device (Huang & Rubinsky, 2003)

These devices hold many promises for the future of electroporation technology.

Currently many of the electroporation devices are oriented toward working with small numbers of cells or a single cell to enable the study of process mechanisms or specific cell parameters. Some of the fabrication processes require highly specialized equipment and involved fabrication processes. Microdevice electroporation has not been widely commercialized and remains largely a laboratory scale device.

Chapter 3 : Optical tweezers electroporation

3.1 Introduction

Electroporation has great potential to impact health care through clinical applications of gene therapy. Before we realize its full potential for clinical applications and beyond we must develop a better understanding of electroporation mechanisms and abilities. Electroporation is a complex stepwise process that is difficult to understand from a macro process viewpoint; however when we develop a fundamental understanding of how each step works and how they impact with each other we can begin to understand and tailor electroporation protocols to a specific application.

Here we share our investigation of the electroporation process using optical tweezers. The optical tweezers allow us to observe a single or small number of cells throughout the entire electroporation process. We are also able to use the optical tweezers to recreate exact conditions between experiments enabling us to produce quantitative information in addition to our qualitative observations. The optical tweezers approaches used here are the least invasive methods available providing a minimal process interference while investigating electroporation. The studies we conducted allowed us to test theoretical models of cell size dependence on

36 electroporation parameters, as well as how interactions between cells can affect electroporation outcomes.

Our goal for cell size experiments was to understand the relationship between cell size and the critical transmembrane potential when a cell is first electropermeabilized. Specifically we wanted to test the relationship established by a commonly simplified version of the Schwan equation (Section 2.3.5 equation (4)). In cell- cell interaction experiments we aim to understand how locations of cells relative to one another impact the electric field and potential electroporation outcomes.

3.2 Materials and methods

3.2.1 Experimental setup

The optical tweezers electropermeabilization system consists of three basic units that have been combined to enable electroporation studies. Figure 3.1 shows a diagram of these units. The heart of the system is the microscope outfitted with a digital camera connected to a PC for video capture and a mercury lamp with appropriate filters for fluorescent illumination of samples. The camera has a photomultiplier in its detector capable of enhancing signals from the samples being viewed. The next unit is the optical trapping system. This consists primarily of the trapping laser and micromanipulation stage although there are peripheral mirrors, filters and prisms that polarize and guide the laser through the microscope objective lens to the sample being trapped. The

37 micromanipulation stage enables the stage to be moved in fine increments around the trapped sample. The last unit is the electroporation apparatus. This consists of an electroporation power supply and an electroporation fluidic chip capable of interfacing with the micromanipulation stage and trapping laser.

Back of system Front of system Figure 3.1. Schematic and picture of optical tweezers electropermeabilization system. A microscope image capture unit (dark), trapping laser unit (grey), and electropermeabilization unit (white) make up the three basic parts of this system.

3.2.1.1 Optical tweezers

Optical tweezers can provide control over cell location while maintaining conditions similar to bulk electroporation. It is able to accomplish its trapping functions through changes in index of refraction between the cell and its environment (Ashkin,

Dziedzic, & Yamane, 1987). In bulk electroporation cells remain in suspension throughout the process and are located randomly throughout the suspension. Optical trapping is capable of fixing a cell to a location while remaining in suspension. This enables the observation of a cell throughout an electroporation process and avoids the minority cases in bulk electropermeabilization of cell deformation and non-uniform electric fields generated at the surfaces of cuvettes. This level of control can also be useful in studying some of the electroporation microdevices being developed today since the optical tweezers can move the location of the cell relative to a microfeature or nanofeature of interest.

In our experiments here, cell manipulation was accomplished using an optical tweezers. A 2 W, 1064 nm laser (Crystal Laser) was used to trap cells. The 1064 nm laser is widely used in biological experiments due to minimal laser interaction with samples.

The power of the trapping laser can be adjusted from 0-2 W and was commonly held at

0.6 W for experiments. The optical tweezers trapping laser was guided into the microscope by a series of optical instruments as seen in Figure 3.2. Mirrors (Thor Labs) were used to guide the laser paths. The laser was passed through a series of beam modifying elements including a half wave plate, polarizing prism, and telescoping beam

40 expander. The half wave plate (Thor Labs) took in the laser beam and output a polarized wavelength in a single direction. This half wave plate provides a system for regulating the relative power of the trapping beams after they are split by the prism. A prism (Thor

Labs) was used when multiple traps were needed for an experiment. The prism split the beam according to beam phase into two beams. The split beams roughly equal to half the original laser power. A telescoping beam expander (Thor Labs) was used to adjust the beam size before going into the microscope. Once the beam was focused into the side port of the microscope it was brought to the sample. We were able to adjust the relative position of beams, thus the cells, using laser guide mirrors.

Manipulation of a trapped cell is accomplished on the microscope. The beam was held in a stationary location while the microscope stage was moved relative to the beam. The location was controlled by a micromanipulation stage (Mad City) on top of the microscope. The stage was capable of moving on the micrometer scale in the x-y plane while the vertical z-plane is controlled by the focal location of the objective lens.

Figure 3.2. Optical tweezers laser guide equipment

3.2.1.2 Microscopy

A Nikon X-81 inverted microscope in conjunction with a Photometrics Cascade II:

512 EMCCD camera was used to capture video of electropermeabilization events. We captured images and focused the trapping laser with a 60x water immersion Nikon objective (NA = 1.2) under aqueous emersion fluid. Images were taken through the glass cover slip (no.1 thickness) into a custom fluidic electroporation chip. Exposure settings were based around a 20 ms shutter speed. We achieved capture and analysis of the images using a PC equipped with Metamorph software. The microscope was equipped with a 100W mercury lamp (Nikon) and an appropriate filter block for propidium iodide

(PI) dye fluorescence detection. The microscope stage was equipped with a

42 micromanipulation control. This micromanipulation controller was used to move the stage and fluidic chip relative to the fixed position of the cell once trapped.

3.2.1.3 Electroporation device

The electropulse generation apparatus consisted of a complete Bio-RAD Gene

Pulser Xcell ™ electroporation system with custom electrode contacts. It had the ability to generate a variety of different DC pulse sequences and forms including step inputs and exponential decay. Common pulse sequences used in our experiments were either a series of 10 unipolar DC step pulses in the range of 60-110 volts with a 1 ms duration at a frequency of 10 Hz or a single DC step pulse in the range of 60-110 volts with a 1 ms duration.

A custom fluidic chip was used to carry out the electropermeabilization. The chip consists of a 1.2 mm x 10 mm x 1 mm channel lined with parallel electrodes down the side lengths of the channel. This system can provide a uniform electric field across the channel.

The fluidic chip was constructed of poly(methyl methacrylate) (PMMA) for the main channel with copper electrodes lining the sides. We machined a 1.5 mm thick sheet of

PMMA (McMaster-Carr item No. 8560K171) with a custom CNC micromilling machine

(Aerotech). Adhesive backed copper film (McMaster-Carr item No. 76555A712) was used to form the electrodes on the sides of the PMMA channel. The copper electrodes were soldered to electrical leads that connected to the power supply. The channel in the fluidic chip was sealed with a glass cover slip (no.1 thickness) and instant adhesive

(Loctite ™ 495). This fluidic chip allows images to be captured from an inverted microscope while operating the optical trapping laser through the glass cover slip. It is capable of generating electric fields of 100-10,000V/cm when combined with the Bio-

RAD power supply. The system is schematically shown in Figure 3.3.

a b

Figure 3.3. electropermeabilization fluidic chip (a) and cell location in the chip during experiments (b).

3.2.2 Propidium iodide

We used cell impermanent dye propidium iodide (PI) to detect electropermiabilization (Figure 3.4). It is commonly used to detect dead cells. Under normal circumstances a cell has an intact cell membrane and this dye can’t pass through it. As a result it exhibits a low fluorescence. PI is part of a class of chemicals know as intercalation agents. If the cell membrane is compromised or damaged this dye is able to pass into the cell and combine with nucleic acids, DNA or RNA, and as a result exhibits

44 a 30x enhancement of its fluorescence. It is more likely that the PI dye combines with messenger RNA and small RNA in our experiments to exhibit strong fluorescence in the cytoplasm. The size of PI is roughly 668 daltons and can pass through openings on the order of 20 nm. We purchased PI dye from Invitrogen (catalog No. P3566).

Figure 3.4. Chemical structure of Propidium Iodide (PI) dye

In the experiments cells (104 cell/ml) were incubated in a 100 µM PI in D-PBS solution for 10 minutes prior to electroporation. Cells were loaded into the custom microfluidic and non-fluorescent cells were then manipulated and electroporated as described. A high intensity 100 W mercury lamp (Nikon) with appropriate detection filters for this dye was used to detect fluorescence from PI dye entering the cell.

3.2.3 Cell culture

3.1.1.1. K562 cell culture

We used cell line K562 (a leukemia cell line, CCL-243) in optical tweezers experiments. This cell line is a cancer model animal cell line. Cells were procured from

American Type Culture Collection (ATCC, Manassas, VA). We used Invitrogen (Carlsbad,

CA) to supply the culture media and additives. Cells were grown in nutrient medium consisting of RPMI 1460 (D-MEM/F-12, catalog No. 21870-076), with the addition of

10% (v/v) newborn calf serum (NCS, heat-inactivated, catalog No. 26010), 2 mM L- glutamine (catalog No. 25030), 1mM sodium pyruvate (catalog No. 11360). Cells were

2 o cultured in 25 cm T-flasks incubated at 37 C under an atmosphere of 5% CO2 . We passaged cells every 2 to 3 days cells by placing a portion in a fresh media filled flask.

Leading up to an experiment we allowed the K562 cells to grow to 80% confluence and then we harvested them from suspension. We centrifuged cells into a pellet and washed them with Dulbecco's Phosphate Buffered Saline (D-PBS, catalog No.

14190). The pellet was resuspended in D-PBS containing 100 mM PI (Invitrogen, catalog

No. P3566).

3.2.3.1 NIH-3T3 cell culture

NIH 3T3 fibroblasts (a mouse fibroblast cell line, CRL-1658) were used in optical tweezers experiments. This cell line is a broad based model animal cell line. Cells averaged 15.6 µm in diameter (1.9 µm standard deviation). Cells were procured from

American Type Culture Collection (ATCC, Manassas, VA). We used Invitrogen (Carlsbad,

CA) to supply the culture media and additives. Cells were grown in nutrient medium consisting of Dulbecco’s modified Eagle’s medium nutrient mix F-12 (D-MEM/F-12, catalog No.10565), with the addition of 10% (v/v) newborn calf serum (NCS, heat- inactivated, catalog No. 26010), 2 mM L-glutamine (catalog No. 25030), 1mM sodium

o pyruvate (catalog No. 11360). Cells were cultured in 25 cm2 T-flasks incubated at 37 C under an atmosphere of 5% CO2 . Every 2 to 3 days cells were detached from the surface of the culture flask with a solution 0.05% trypsin with 1mM EDTA·4Na (Catalog No.

25300) and placed in fresh media filled flask.

Leading up to an experiment we allowed the NIH 3T3 cells to grow to 80% confluence and then we harvested them by trypsinization. We centrifuged cells into a pellet and washed them with Dulbecco's Phosphate Buffered Saline (D-PBS, catalog No.

14190). The pellet was resuspended in D-PBS containing 100mM propidium iodide

(Invitrogen, catalog No. P3566).

3.1.1.2. Mouse Embryonic Stem Cells

We procured mouse embryonic stem cells (MES cells, CCE strain) from StemCell

Technologies (Vancouver, BC, Canada). Invitrogen (Carlsbad, CA) was the source for all

MES CCE cell culture reagents unless noted. MES cells were maintained in an undifferentiated state using a high glucose Dulbecco’s Modified Eagle’s Medium (DMEM with 4500 mg D-glucose/L, StemCell Technologies, catalog No. 36250) supplemented with 15% ES-Cult fetal bovine serum (FBS, StemCell Technologies, catalog No. 06952), 2 mM L-glutamine (catalog No. 25030), 1 mM MEM sodium pyruvate (catalog No.11360),

1000 U/mL recombinant mouse leukemia inhibitory factor (rm LIF, Millipore, catalog No.

LIF2010), 100 U/ml penicillin G + 10 μg/ml streptomycin (catalog No. 15140), 0.1 mM

MEM non-essential amino acids (NEAA, catalog No. 11140), and 150 μM monothioglycerol (MTG, Sigma-Aldrich, catalog No. M6145). Mouse ES cells were cultured in 0.1% (w/v) gelatin coated t-flasks and at 37°C under an atmosphere of 5%

CO2. MES cells were passaged every 3 days by trypsinization. Experiments using Mouse

ES cells were conducted at 50-70% confluency. To suspend the colonies into single cells,

0.25% trypsin with EDTA·4Na (catalog No. 25200) was used for harvesting; for intact colony suspension, 0.05% trypsin with EDTA ·4Na (catalog No. 25300). Sizes of MESc can be as small as 6 µm in diameter and as large as 30 µm. The average cell size was 14.7 µm

(standard deviation 5.8 µm).

3.2.4 Electroporation procedure for size experiments

Our goal for cell size experiments was to understand the relationship between cell size and the critical transmembrane potential when a cell is first electropermeabilized. We set about making as controlled, and simple-to-interpret study as we could design. Experimental considerations included (a) using cell lines with spherical cells and a large range of cell sizes, (b) working with cells in suspension in contrast to plated cells, and (c) using electropermeabilization conditions that are similar to those routinely use in applications. Our basic idea was to isolate a single cell at a precise location in our apparatus, measure the cell’s radius, and then subject it to

48 increasing voltage pulses until we observed permeabilization. A major experimental difficulty is that the voltages required for electropermeabilization generate enormous numbers of bubbles due to electrolysis and these violently move cells making it ordinarily impossible to track a single suspended cell. Our solution was to hold cells under study with optical tweezers. An optical tweezers uses a tightly focus laser beam to manipulate microscope objects based on differences in the indices of refraction between the object and its environment. Our apparatus was able to hold various types of cells against the electrolysis bubbles for permeabilization pulses shorter than 3 ms.

Various diagnostic tests we conducted indicated that the optical tweezers itself has negligible impact on the permeabilization process.

Figure 3.1 and Figure 3.3 illustrate the systems used in our experimental approach. A suspension of cells in a D-PBS (catalog No. 14190) and PI solution was introduced into a custom built poration cuvette mounted on the stage of an inverted microscope. The microscope was used both for observations and to provide the tight focus required for the laser tweezers beam. About 0.6 W of laser power at 1064 nm was used to select and position a cell for study. The cell’s radius was measured from pictures made by our imaging system. A series of voltage pulses would then be applied to the cuvette’s electrodes such that successive pulses increased the field experienced by the cell in steps of either 8 V/cm or 16 V/cm. Usually a pulse length of 1 ms was used and the time between pulses was typically a half a minute. Note: we investigated whether our data were sensitive to the interval between permeabilization attempts and measured no dependence at all for intervals ranging from 15 s to 2 minutes. After each

49 pulse, the cell was watched to see if it had been permeabilized. To detect electropermeabilization, the cell impermanent dye propidium iodide (PI) was used.

Typically between 3 and 5 cycles of pulsing and observation would occur before a cell was permeabilized. Once permeabilization was observed, the last pulsing voltage was recorded and the pulsing, observation cycle stopped. To prepare for the next measurement, the poration cuvette was flushed with deionized water, dried and reloaded with a fresh suspension of cells. The custom poration cuvettes were used roughly 20 times before being discarded.

3.2.5 Procedures for cell-cell interaction experiments

Our goal for cell-cell interaction experiments was to understand how cells affect electroporation outcomes based on their relative proximity to one another. We designed our experiment to be as controlled and simple-to-interpret study as possible.

We sought to use electropermeabilization conditions that are similar to those routinely use in practical applications, and selected cells of the same or similar size within each experiment. Additional experimental consideration included using cell lines with spherical shape that can remain in suspension, in contrast to plated attached cells. Our basic experiment was to isolate two cells at a precise location in our device, line them up in one of our predesigned configurations relative to each other and then observe how they responded to electroporation followed by recording differences in critical transmembrane potential.

A suspension of cells in a D-PBS (catalog No. 14190) and PI solution was introduced into a custom built poration cuvette mounted on the stage of an inverted microscope. The microscope was used both for observation and to provide the tight focus required for the laser tweezers beam. 0.6 W of laser power at 1064 nm was used to select and position a cell for study. We conducted experiments to determine the optimum settings that were above the critical transmembrane potential without causing cell rupture and used the same settings in each experiment. The optical trap beam was split using a prism and used to trap multiple cells relative to each other.

In these experiments, we manipulate cells to create the desired situation and hold them in suspension using the optical tweezers. Once we positioned the cell or cells without any other interfering cells around them we delivered an electric pulse to the system (800 V/cm, 10 pulses, 1 ms pulse duration, 10 Hz). The surrounding media contained PI (100µM) to detect successful electroporation. A study using trypan blue

51 dye was conducted after an experiment to confirm that these parameters produced reversible electroporation.

In the single cell experiments the critical transmembrane potential was identified by increasing voltage pulses until we observed permeabilization. A series of voltage pulses would then be applied to the cuvette’s electrodes such that successive pulses increased the field experienced by the cell in steps of either 8 V/cm or 16 V/cm. Usually a pulse length of 1 ms was used and the time between pulses was typically a half a minute. Note: we investigated whether our data were sensitive to the interval between permeabilization attempts and measured no dependence at all for intervals ranging from 15 s to 2 minutes. After each pulse, the cell was watched to see if it had been permeabilized. To detect electropermeabilization, the cell impermanent dye propidium iodide (PI) was used. Typically between 3 and 5 cycles of pulsing and observation would occur before a cell was permeabilized. Once permeabilization was observed, the last pulsing voltage was recorded and the pulsing, observation cycle stopped. To prepare for the next measurement, the poration cuvette was flushed with deionized water, dried and reloaded with a fresh suspension of cells.

A major experimental difficulty is that the voltages required for electropermeabilization generate enormous numbers of bubbles due to electrolysis and these violently moved cells making it impossible to track a single suspended cell. Our solution was to hold cells under study with optical tweezers. An optical tweezers uses a tightly focused laser beam to manipulate objects based on differences in the indices of

52 refraction between the object and its environment. Our apparatus was able to hold various types of cells against the electrolysis bubbles for permeabilization pulses shorter than 3 ms. Various diagnostic tests we conducted indicated that the optical tweezers itself has negligible impact on the permeabilization process.

3.2.6 Computer simulation

In order to solve the cell-cell interaction in 3D electric field, we use both 3-layer

(single shell) and the contact resistance method with the finite element commercial software COMSOL (Mathworks Inc). The 3D simulation using the contact resistance method is used for the perpendicular case, while 2D axis-symmetric simulation using the

3-layer model is for the parallel case.

Values of conductivities of intercellular cytoplasm, external surrounding medium, and cell membrane take a very board range depending on types of cells and

7 solutions. We chose standard values  i   e  0.2 S/ m and  m  510 S/m based on other work (Kotnik, Bobanovic, & Miklavcic, 1997). The radius of a cell assumed as

R  7 m and the membrane thickness is d  5 nm . The external bulk electric field strength is set to 1000 V/cm in the 3D simulation.

3.2.7 Procedure for optical tweezers effects

Cell viability post electroporation is equally important as delivering PI in showing that we successfully electroporated a cell. Our experiment used trypan blue dye to determine cell viability post electropermeabilization. Trypan dye is unable to cross an intact cell membrane and when it does cross the cell membrane it causes the cell to turn blue in color and darkens the appearance of the cell. An electroporated cell is expected to darken as some dye will enter the cell upon electropermeabilization. A dead cell will turn a darker shade since the dye has infinite time to saturate the inside of the cell as compared to electropermeabilization where dye has milliseconds to enter. The equipment described in Figure 3.1 was used to carry out these experiments. A K562 cell was held in the optical tweezers in a 0.025% trypan solution and electroporated.

Electropermeabilization settings were at an 850 V/cm electric field, 10 pulses, 1 ms pulse duration with a 10 Hz frequency between pulses.

Crucial to this work is checking to see if the optical tweezers themselves significantly affect the permeabilization process. We carried out three experimental tests using the equipment in Figure 3.1. First using K562 cells, we reduced the laser power used to hold cells by a factor of 10 to 0.06 W and measured permeabilization thresholds as indicated in the cell size experiments. Normally we keep the trapping laser power at about 0.6 W to hold the cell through the 10 pulse experiments. For the lowered laser power trap, we were only able to keep a cell trapped through a single- pulse. As a second test, we made additional measurements similar to the cell size

54 experiments for K562 cells, however we turned off the trapping laser for the 1 ms period that the electroporation pulse was applied. We accomplished this control through the use of LabView software combined with a National Instruments A/D conversion system. Our final test was to use 3T3 cells plated out on a surface of our permeabilization cuvette. NIH-3T3 cells normally adhere to surfaces making it easy to move them while suspended to a specific location in the electroporation device with the optical tweezers and have them attach and remain in that location without the tweezers. Since the cells stick to the surface we are able to track the individual cells throughout the electroporation process. Single cells attached to the surface with and without the laser were compared along with single cells that remained trapped in suspension. We included PBS and 100 mM PI dye in the media for electroporation to detect a successful electroporation event. The Ec values were obtained in the same method as described for the K562 and MES cells in cell size experiments.

3.3 Results

3.3.1 Cell size and electroporation

Figure 3.6 displays a diagram of a cell in the electric field with our angle convention as well as a schematic of our electroporation chip and a time series of fluorescent images that represent a typical result of an electroporated cell. Figure 3.7 and Table 3.1 summarize our primary results. We initially studied K562 cells, a line of immortalized leukemia cells. In addition to the one millisecond, single pulse

55 measurements described above, we also measured permeabilization thresholds for single 100 s pulses and for a 10 pulse series of one millisecond pulses at a frequency of

10 Hz. Rols and Teissie (Rols & Teissie, 1998) claim that while the basic fact of permeabilization is determined solely by the size (voltage or field strength) of an applied pulse, the degree of permeabilization is determined by the length and number of pulses.

We explored the sensitivity of our results to pulse length by taking the Ec measurements using a 100 µs pulse length. Our ten pulse program checks if the single pulse measurements miss successful electroporation simply because the degree of permeabilization is below our detection sensitivity, a false negative. In Figure 3.7, both the 100 measurements and the ten pulse program results are indistinguishable from the single 1 ms pulse measurements. Therefore, we believe that our conclusions are robust with respect to pulse length and number in that we observe all successful electroporation attempts, with regard to PI fluorescence. Two features are clear from the K562 cell data. First, electropermeabilization is a stochastic process since identical sized cells can have different measured permeabilization thresholds and second, there is no apparent dependence of permeabilization threshold on cell size. The stochastic nature of the electroporation process was noted by Weaver (Weaver J. C., 1996), based on similar observations in studies on the breakdown of planar lipid membranes. The observed independence of cell size was unexpected and is counter to the intuition of many researchers in this area. Consequently we carried out measurements on two other lines of spherical cells that were available to us: mouse embryonic stem cells (MESc) and

NIH-3T3 cells (3T3). These additional data confirmed the essential K562 observations.

The MESc cells allowed measurements for over 3-15 µm variation in radius but showed no relationship between cell size and permeabilization threshold field. In Table 3.1 we summarize our data. We calculated the mean threshold field and standard deviation for each cell line, and include all measurements made for a cell line in the mean. The mean permeabilization threshold fields for the three lines are similar but measurably different. The variation in the mean permeabilization field for these three cell lines is about 30%.

a b

E 0

A B C 10µm D

Figure 3.7: Minimum electric field causing permeabilization vs. measured cell radius. ●-K562 10 pulse,○- K562 single pulse, ♦-MES 10 pulse, ◊-MES single pulse, ▲-NIH-3T3 single pulse

Avg. Threshold Field Std Deviation Cell line (V/cm) (V/cm)

K562 692 32.0

MESc 555 30.1

3T3 509 22.2

Table 3.1. Average critical electric fields with standard deviations

An alternatively way of understanding our data is to use each individual data point to estimate the threshold transmembrane potential needed for permeabilization of a cell. If in Eq. 5 (ch.2) it is assumed that the membrane resting potential(휑푠 + 휑푛 ) is small enough to be ignored, and the membrane charging time is long enough, threshold permeabilization occurs at the poles of a cell when:

Vperm 1.5Ecr . (1)

where Ec is the minimum applied field to cause electroporation. Equation (1) is a simplified version of what is commonly called the Schwan equation. Scientists in the field often view the situation this way because it is generally felt that the threshold transmembrane permeabilization potential is fairly constant (0.2 V- 1.5 V) for different

59 types of cells and very constant for a given cell line. Our results have just shown that this

appears not to be the case. At this point we can use our measured values of r and Ec in

Eq. 1 to construct a transmembrane permeabilization threshold for each measurement.

We have done this in Figure 3.8. Now excellent linear fits in Figure 3.8 reflect our

previous observation that Ec in Eq. 1 is nearly constant and independent of cell size. We found that each individual measurement in our results is reasonable. We have indicated on Figure 3.8 the range of threshold potentials of almost all previous work and note that all of our data lie within that range. We have additionally included data produced by

Zimmerman as one of the only comparable studies like the one we have conducted here

(Zimmerman, Groves, Schnabl, & Pilwat, 1980).

Figure 3.8: Transmembrane potential thresholds for permeabilization inferred using Eq. 1. The red lines define the range of threshold values found in almost all previous work. Also shown are the “double Coulter counter” data of Zimmermann et al. On this graph, data fit by a line passing through the origin indicate the constant threshold electric field dependence on cell radius that we find. ●-K562 10 pulse,○- K562 single pulse, ♦-MES 10 pulse, ◊-MES single pulse, ▲-NIH-3T3 single pulse, X-Zimmerman Data

3.3.2 Cell-cell interactions

3.3.2.1 Single cell in bulk electric field

Our experiment using a single cell exposed to a uniform electric field provides a reference to compare with the following multi-cell experiments. The use of optical tweezers makes it possible to manipulate and isolate a single cell while it remains in suspension providing the closest approximation to date of a system to study bulk electroporation. Figure 3.9 shows the time series entrance of PI dye into a single cell during electroporation. It can be seen in Figure 3.9 C that initially after electroporation the dye was concentrated at the side of the cell facing the positive electrode. As time went by the dye spread and caused the entire cell to become fluorescent. This typically took about 30-60 seconds with our experimental conditions. This suggests that the dye diffuses relatively quickly through the aqueous electroporation media outside the cell and is then quickly retarded by the cells cytoplasm as it diffuse and interacts with nucleic acids.

Figure 3.9(f) explains why we observe that the PI dye enters the positive electrode side first (  0 ). A PI molecule is affected by two external forces: one is the diffusive force c pointing towards the cell because the PI concentration outside of the cell is much higher than that inside the cell; the other is the electrophoretic force qE since it is positively charged. Here  is the diffusion coefficient, c is the concentration, q is the charge density of PI, and E is the electric field vector. Since

62 electrophoresis is dominant, we expect the net force for a PI particle near the positive electrode to point to the cell and thus it can enter the electropores during the electroporation. For PI near the negative electrode side (   ), the net force is pointing away from the cell; thus it cannot enter the cell through the nanopores while the electroporation pulses take place. After the electropulse we see in Figure 3.9(c,d) that some diffusive flux of PI dye is present on the negative electrode side but it is much less than the combination of diffusive and electrophoretic flux on the positive electrode side.

(f)

Figure 3.9.PI entrance into a single cell during electroporation at time (a) 0s; (b) 7.5s; (c) 15s; (d) 22.5s; (e) 30s; (f) Schematics on how positively charged PI dye enters the cell.

The calculated transmembrane potentials by the 3-layer model (in “” symbol)

63 and the “contact resistance” method (in “O” symbol) are compared with the analytical solution (dashed curve) in Figure 3.10. Due to axis-symmetric properties we only show the distribution around half of the circle passing through the positive and negative poles

(thus the facing angle varies from 0 to  ). We can see that the simulation results agree very well with the analytical solution.

Figure 3.10. Distribution of transmembrane potential for a single cell in bulk electric field: comparison of simulations and the analytical solution (dashed curve).

3.3.2.2 Two-cell systems

3.3.2.2.1 Parallel to the electric field lines

When cells are in series parallel to the electric field lines we have an axis- symmetric case using the 3 layer model, spacing between cells in simulation include seven different gaps: g = 0.25, 0.5, 1, 5, 10, 20, and 40 µm. In the physical experiments,

64 we consider two gaps, i.e., close and far cases with g = 1 and 20 µm.

For closely-separated cells (with a gap of 1 µm) in the parallel case, Figure

3.11(a) shows the finite element mesh near two cells. Here, the left boundary is the axis-symmetry line. We used a dense mesh near the cell membranes to capture the most detail in this area. The electric field lines around these two cells are shown in

Figure 3.11(b). Due to the low conductivity of cell membrane, the electric field is very small inside the cells and a majority of the electric field lines pass around the cells. We also plotted the distribution of transmembrane potential in Figure 3.11(c), where the dashed curve is the distribution for a single cell in the electric field, “O” is for the top cell, and “” for the bottom cell. The model predicts that the values of transmembrane potential for both cells are lower than that for a single cell in the bulk electric field. The total current through the cell membrane decreases or the total resistance increases since cells behave as series resistors in an electric circuit with the fixed external electric potential. Thus the simulation predicts a higher electric field and transmembrane potential is required to electroporate this two-cell system compared with the single cell system in bulk electroporation. Figure 3.12 shows a typical time series of images taken when a result indicates the Ec has been reached.

(a) (b)

Figure 3.12. PI entrance in to two closely-separated cells parallel with the electric field at time (a) 0s; (b) 6s; (c) 12s; (d) 18s; (e) 30s; (f) 60s.

We found that when cells are parallel with the electric field spaced far apart (20

µm gap) electroporation for each cell was an independent process. As shown in Figure

3.13(a)-(e), there is not much difference from a single cell electroporation in the bulk electric field. Also the simulation indicated that the transmembrane potential distribution is close to that of a single cell in the bulk electric field (as shown in Figure

3.13(f)).

(f)

We use the simulation to investigate the relation of the maximum transmembrane potential vs. the gap between two cells in the parallel case (Figure

3.14). With the gap between two cells increases, the maximum value of transmembrane potential follows a monotonic curve up to a value of 1.05 V, which is the maximum value for a single cell (radius 7 µm) in the bulk electric field of 1000 V/cm.

Figure 3.14. Maximum value of transmembrane potential vs. gap between cells in the parallel case.

3.3.2.2.2 Perpendicular to the electric field lines

When cells are lined up perpendicular to the electric field lines the geometry is no longer axis-symmetric, therefore we need to carry out a 3-dimensional simulation.

We consider 7 gaps in the simulation: g = 0.25, 0.5, 1, 5, 10, 20, and 40 µm. In the physical experiment two gaps: g = 1 and 40 µm were used.

We conducted a simulation for closely-separated cells perpendicular to the electric field lines to compare with the physical experiments. Relative mesh densities

69 near the surface and boundaries can be seen (Figure 3.15(a)). The 3D electric field lines around two cells are shown in Figure 3.15(b). We can see that the electric field lines also pass around cells due to the low conductivity or high resistance of the cell membrane.

(a) (b)

Figure 3.15. (a) Mesh and (b) electric field lines for two cells perpendicular to the electric field with a 1 µm gap

Figure 3.16 shows the electroporative PI entrance in two closely-spaced cells perpendicular to the electric field at different times. In Figure 3.16(c) and (d), it appears that the intensity of PI dye is equal at both positive and negative poles, which is similar to Figure 3.22(e). Thus the simulation and experiment show that the maximum magnitude of the transmembrane potential may be affected by the cell-cell interaction.

f E

When spaced far apart perpendicular to the electric field, we found that the cells exhibited equal fluorescence. Figure 3.17(a)-(f) shows PI entering two far-spaced cells perpendicular to the electric field at different times. Time series images show some cell rotation occurring in this experiment. Figure 13(b)-(d) from video captured during the experiment shows that the highest intensity of PI dye is moving clockwise with time due to cell rotation. The rotational motion does not affect the Ec values, but may alter our

71 understanding of delivery mechanisms between rotational and non-rotational situations. Figure 13(g) shows that the distribution of transmembrane potential is nearly identical to that of a single cell.

(g)

The relationship between the maximum transmembrane potential and the gap between two cells in the perpendicular case is shown in Figure 3.18 by simulation. We can see that the maximum transmembrane potential decreases exponentially as the gap between cells increases.

Figure 3.18. Maximum transmembrane potential vs. gap for cells perpendicular to the electric field lines.

A drawback of the “contact resistance” method is that the accuracy depends on the mesh density. As shown in Figure 3.18, when two cells are far separated in the perpendicular case, the maximum value of transmembrane potential should be very close to that of the single cell in bulk case, 1.5ER 1.05 V . However, for g = 40 µm, the maximum value is around 1.0464 V, thus the relative error is around 0.34%. This small

74 error is associated with the relative coarse mesh density near the cell membrane used in the “contact resistance” method. A denser mesh for this problem requires larger computer resources to run the mesh generation and 3D calculation. Fortunately, this tiny error does not affect the trend of the relation of maximum transmembrane potential vs. the gap of two cells in perpendicular case.

3.3.3 Critical electric field

We conducted a membrane breakdown experiment to determine transmembrane potential differences as predicted by the modeling between the parallel and perpendicular close orientations. In this experiment we held cells with the optical tweezers and successive electroporation pulses were delivered to the cells until membrane breakdown. Cells we selected for these experiments were between 12 and

14µm in diameter. Pulse length, number of pulses, and pulse frequency was held constant at 1 ms, 10, and 10Hz respectively. We adjusted the pulsed electric field strength between electroporation processes until membrane breakdown was detected by PI dye fluorescence. We used electric field steps between 8 and 16 V/cm and typically took between 2-5 electric field adjustments to produce electroporation and to arrive upon the breakdown electric field. The steps between pulses were set by the minimum step size allowed by the power supply. Measurements were taken of the pulse out of the power supply to confirm the step sizes.

As predicted in the modeling results, the cells oriented parallel to the electric field require a higher field strength to achieve electroporation, while the cells oriented perpendicular to the electric field require a lower electric field to produce successful electroporation (Figure 3.19). Single cell data required an electric field between the parallel and perpendicular cases which was also predicted by modeling results. Data were averaged between 9 separate experiments for each situation. The parallel case required an average field strength of 817 V/cm with a standard deviation of 45V/cm while the single cell case required 723V/cm with a standard deviation of 27V/cm and the perpendicular case required an average of 669V/cm with a 26V/cm standard deviation.

Figure 3.19. Electroporation membrane breakdown threshold for two K562 cells oriented parallel or perpendicular to the electric field and a single k562 cell.

The difference in electric field strength between the parallel case and the single cell case is 94 V/cm. We conducted a statistical T-test to compare the mean electric field needed to produce electroporation of these two cases. The null hypothesis (H0) is that these two cases require an equal critical field strength (Ec 1=Ec 2) to produce electroporation while the alternative hypothesis (H1) is that they do not (Ec 1≠Ec 2). This is a two tailed test assuming equal variance in the breakdown thresholds. Based on the T- test there is a less than 1% chance of observing the values we obtained in our experiments therefore we reject the null hypothesis.

We observed a difference of 148 V/cm in electric field strength between the parallel case and the perpendicular case. Again we conducted a statistical T-test to compare the mean electric field needed to produce electroporation of these two cases.

The null hypothesis is that these two cases require an equal critical field strength (Ec 1=Ec

2) to produce electroporation while the alternative hypothesis is that they do not require an equal critical field strength (Ec 1≠Ec 2). This was also a two tailed test assuming equal variance in the breakdown thresholds. Based on the T-test there is a less than 1% chance of observing the values we obtained in our experiments therefore we reject the null hypothesis and conclude that these two cases have a statistically different Ec.

The difference in electric field strength between the single cell case and the perpendicular case is 54 V/cm. We conducted T-test (Ho Ec 1=Ec 2, H1 Ec 1≠Ec 2) to compare the mean electric field needed to produce electroporation of these two cases. This was a two tailed test assuming equal variance in the breakdown thresholds. Based on the T- test there is a less than 1% chance of observing the values we obtained in our experiments therefore we reject H0 in favor of H1 meaning the Ec values are statistically different.

In this experiment we investigated the shielding effects created when cells were placed 1 micron apart oriented parallel to the electric field. We repeated the experiment 21 times, and each experiment fell into one of three categories; top dominated electroporation, bottom dominated electroporation, or equal electroporation. We required the top dominated case to be at least twice as fluorescent

78 as the bottom cell. The bottom dominated case was required the bottom cell to be at least twice as fluorescent as the top cell. The equal case is a situation where both cells exhibited an equal fluorescence and any case that did not meet the previous two cases criteria was counted here. We measured cell fluorescence intensity as the average intensity across the entire cell and we recorded measurements once the value became stable or intensity reached plateau. The background intensity was subtracted from the average intensity to produce a final value for each cell. Metamorph software was used to aid in fluorescence measurements and in determining when the cell fluorescence stabilized. It can be seen in Figure 3.20 that half of the time one of the cells would exhibit a much stronger fluorescence than the other indicating a stronger electroporation event while the other half of the time the cells were equal in fluorescence intensity. We conducted the same experiment for cells oriented perpendicular to the electric field and in all of the cases the cells exhibited equal fluorescence.

Figure 3.20. Dominant cell in electroporation for two cells parallel to the electric field

3.3.4 Electroporation axis shift

The 3D distribution of transmembrane potential on the cell membrane is shown in

Figure 3.21(a). We can see that the maximum value of transmembrane potential was shifted off of a straight line axis between 0 and 180 degrees seen for the single cell case

(Figure 3.6), and the new axis bends toward the opposite cell. We plotted the contour of transmembrane potential projected on the X-Z plane in Figure 3.21(b). It was apparent that both maximum and minimum values of transmembrane potential were not at the positive or negative pole. Here we used arrows to indicate the location of

80 maximum and minimum values of transmembrane potential. Moreover, we plotted the curve of transmembrane potential vs. the facing angle around the cell membrane in

Figure 3.21(c). We can see that the value of transmembrane potential increased and was shifted slightly away from the positive (θ=∏) or negative pole (θ=0).

(a)

(b)

Figure 3.21. (a) 3D distribution; (b) Projection on X-Z plane of transmembrane potential; (c) Distribution of transmembrane potential (symbol “O” is for the cell on the right, while “” for the cell on the left, and the dashed curve is the single cell in bulk electric field) continued.

(c)

Figure 3.21 continued. (a) 3D distribution; (b) Projection on X-Z plane of transmembrane potential; (c) Distribution of transmembrane potential (symbol “O” is for the cell on the right, while “” for the cell on the left, and the dashed curve is the single cell in bulk electric field).

Modeling results predict a shift in the electroporation axis when cells are perpendicular to the electric field as shown in Figure 3.21c. Experimental results in

Figure 3.22a and b display the fluorescence intensity around the edge of the cell membrane averaged throughout the first 40 seconds of electroporation footage taken for the perpendicular close and far situations. An average of 5 pixels defined the intensity value centered along the edge of the cell as outlined in the white circles in

Figure 3.22e and f. This 5 pixel band represents a 1.3 µm wide section. The angle at the top of the cell image is noted as zero and faces the negative electrode, while the bottom of the cell is at 180 degrees and faces the positive electrode. Intensity peak angles are defined by the midpoint of the area under the top 10 percent of a peak.

For the perpendicular far case the predicted electroporation axis takes place at 0 and 180 degrees. The experimental results for this case confirm the predicted values. It can also be seen in Figure 3.22b that the left and right cell electroporation axes overlap with each other showing no bias toward the other cell. In the horizontal close situation the simulation results show a slight bias of the electroporation axis toward the opposite cell (Figure 3.22c). Experimental results agree with this prediction. Figure 3.22 shows that the electroporation axis for the cells is shifted to 175 and 327 degrees for the right cell and 170 degrees and 5 degrees for the left cell. It is also important to note that the intensity peaks for each cell are shifted toward each other and that the shift in the cell on the right mirrors the shift in the cell on the left with no overlapping in the angle of intensity peaks.

b a

c d

3.3.5 Optical tweezers effects

Cell viability post electroporation is extremely important to show that our electroporation procedures are successful. Figure 3.23 shows the before and after images of a cell in our viability experiment and also a dead cell for comparison. Figure

3.24 quantitatively shows the same information. It can visibly and quantitatively be seen in these figures that a dead cell has a drop in intensity. The electroporated cell also experienced a 10% drop in intensity shown in Figure 3.24 This intensity drop is more difficult to detect in Figure 3.23. The cell post electropermeabilization did not exhibit the same characteristics as a dead cell and therefore the cells were viable after electropermeabilization under the optical tweezers conditions.

Figure 3.23. A K562 cell exposed to trypan blue before (A) and after (B) electropermeabilization compared with a dead cell (C) exposed to trypan blue dye.

Figure 3.24. relative intensity values for a K562 cell exposed to trypan before and after electropermeabilization compared to a dead K562 cell.

Results from the laser effects experiments are critical to validate our previous data. Six trials were carried out for various sized cells at reduced power and the mean threshold field was measured to be 685 V/cm. This is in agreement with the 692 V/cm average reported for the full power measurements in Table 3.2. As a second test, we made six additional threshold measurements for K562 cells in which we turned off the trapping laser for the 1 ms period that the permeabilization electric field pulse was applied. The average measured threshold using this procedure was 683 V/cm. Again these measurements are statistically indistinguishable from the full power measurements based on a statistical T-test (two tailed, unequal variance, α=0.4, null

87 hypothesis µ1= µ2). To summarize our results: in no case did we detect statistically significant correlations between measured threshold permeabilization thresholds and the presence or strength of the tweezers laser.

Table 3.2. Optical tweezers laser effects

Results presented in Figure 3.25 show that the surface attached cells had Ec values between 500 and 530V/cm regardless of the presence of the optical trapping laser. The Ec values of trapped and suspended cells range between 470 and 530 V/cm which is slightly larger of a range than the attached cells. The presence of the laser does not seem to cause any major differences in Ec values. Additionally it can be seen in

Figure 3.25 that Ec values are independent of the cell size. This result is the same as seen in the Ec data for K562 and MES cells.

Figure 3.25. Effects of the trapping laser on 3T3 electroporation. Data noted with “surface” indicate cells were attached to the surface.

3.4 Discussion and conclusions

Electroporation literature is vast and experimental parameters other than cell size, especially pulse length and number, vary considerably. There are claims that different physical mechanisms are at work depending on short (< < 1 ms) or long pulses

(Benz & Conti, 1981; Zimmerman & Neil, Electromanipulation of cells, 1995). However it is clear that everyone uses the same formulation of the problem, a cell membrane represented as a spherical insulating layer separating conductive interior and exterior

89 contents , as a starting point both for theoretical treatments and in analyzing experiments.

In vitro electroporation frequently leaves important experimental parameters loosely controlled. An electroporation apparatus commonly has inhomogeneous fields, cells sticking to surfaces or other cells, and bubbles from electrolysis that move cells, move fluid, and distort fields. Correlations between these factors and cell size can confound the current understanding of a relationship between electroporation and cell size.

Different criteria have been used to identify “permeabilization.” These include changes in cells’ electrical properties (Zimmerman, Groves, Schnabl, & Pilwat, 1980), observed penetration of various sized molecules into the cytosol, the successful electrofusion of two adjacent cells, (Teisse & Rols, 1993) or the observed expression of reporter genes (Golzio, 2002).

Few prior experiments study the role of cell size in electroporation and these mostly measured statistical average properties of large cell populations as opposed to statistical averages of single cells as reported here. For example, Puc et al (Puc, Kotnik,

Mir, & Miklavcic, 2003) measured electroporation yield as a function of applied electric field for populations of DC-3F cells (Chinese hamster fibroblasts). They assumed that cell size was the only source of threshold field variation and as a result found good agreement with a model based on the this theory. On the other hand, Hojo et al (Hojo,

Shimizu, Yositake, & Muraji, 2003) measured the electroporation efficiency of

Saccharomyces Cervisiae cells as a function of cell size for various fields and found that electroporation rates for small cells were significantly higher than for large cells. This is counter to expectations based on the accepted notion that large cells electroporated at lower fields than small cells.

To our knowledge, the only relevant previous single cell measurements are the

“double Coulter counter” work of Zimmermann et al (Zimmerman, Groves, Schnabl, &

Pilwat, 1980) carried out shortly after the discovery of electroporation

(electropermeabilization ) or “reversible dielectric breakdown” of cell membranes. A

Coulter counter detects and sizes single cells by measuring a reduction of electrical resistance in a volume of buffer media as a cell passes through. They observed that if high fields were used for the resistance measurement, sometimes a lower-than- expected reduction in the resistance occurred. This phenomenon was associated with the high field causing electroporation. As a result they developed a “double Coulter counter” apparatus in which a cell was first detected and sized in a low field Coulter counter and then passed to a second, high field counter that measured its breakdown voltage. This device was used to measure the size dependence of the breakdown potential of guard cell protoplasts in Vicia faba. Results (seen on Figure 3.8) are consistent with the predominant theory that the permeabilization threshold transmembrane potential is independent of cell size and the data can be fit with a horizontal line. However, the data are equally well fit by a line passing through the origin and are thus consistent with our conclusion that permeabilization occurs at a constant critical electric field (Ec) value that is independent of cell size.

To explain these observations; the description of a cell modeled as an insulating membrane surrounded by a conducting interior is true at the steady state of a cell’s transmembrane potential. Many assume that the permeabilization threshold potential is the same for all cells of a given cell line (assuming constant experimental conditions: buffer, temperature…), but maybe this is not the case. Our observations could be explained if smaller cells do have smaller threshold potentials as shown in Figure 3.8.

For example, smaller cells often have higher surface tension in their cell membranes due to size and higher surface tension has been shown to reduce the required permeabilization potential. If electroporation threshold dependence is due to cell membrane tension and is compensated the radius dependence of the Schwan equation

(Eq. 1), then our data could be explained by adding a parameter to the accepted theory.

On the other hand, the Schwan equation and the above analysis are used in most experiments that report transmembrane potential electroporation thresholds

(Zimmerman & Neil, 1995) (Teisse & Rols, 1993). Our measurements call into question the validity and relevance of how well the threshold transmembrane potential is experimentally known. The Schwan equation does not accurately describe the cell during its initial capacitive charging nor is it accurate after regions of the cell membrane have “broken down” and are electrically conductive. It is generally thought that the field induced breakdown of the electrical insulation in the cell membrane is closely related to its permeability to medium sized molecules such as PI, both are called

“electropermeabilization.” It has not been shown experimentally that these occur coincidentally. Hibino et al (Hibino, 1993) observed that after electroporation the

Schwan equation did not describe the transmembrane potential. One possibly is that electrical breakdown is described accurately by the Schwan equation with additional mechanisms that affect permeability to PI that depend on electric field found here.

Finally, cell membranes are inhomogeneous insulators. They contain channels, transporters, protein structures, carbohydrates, and variant lipid domains that have a role in determining threshold permeabilization potentials for a particular cell lines. We suggest that these membrane inhomogeneities have a broader role that is missed when a cell membrane is modeled as a first order homogenous insulator and that is what is being seen in our measurements.

The experimentally observed scaling relationship for transmembrane potential in electroporation is surprising and we can only speculate as to what it is telling us about the process. The simplicity in the scaling of the threshold permeabilization voltage with cell radius leads us to imagine that there is a fundamental feature important to the electroporation process that has yet to be uncovered. The use of optical tweezers for establishing tightly controlled and reproducible experimental conditions should enable additional fundamental experimental investigations.

The cell-cell interaction data clearly shows that cells can have varying effects on one another during electroporation. Three effects that are of most interest based on our results are that relative cell orientations:

1. have significant influence on electroporation threshold potentials,

2. have influence on electroporation location,

3. have influence on electroporation extent.

Our electroporation models agree quite well with our experimental results and as a result they validate each other and our understanding of this part of electroporation.

Understanding what may cause enhancement or reduction of the critical electric field offers an element of control over the electroporation process. We show in Figure

3.19 that the arrangement of cells themselves can adjust the critical electric field up or down. Prior work in this area has shown related results when looking at the cell density in an electroporation suspension (Teissie J. , 2004). In bulk electroporation the random arrangement of cells in suspension or varying gradients of density throughout a cell suspension is likely a key factor in why we see cell death along with transfected and untransfected healthy cells. Optimization of bulk electroporation devices is simply finding a “happy medium” where the fewest cells die and the most cells get transfected.

The previous acceptance of these results is most likely due to the assumption that it has something to do with different cell sizes within a given population of cells as mentioned in the cell size results.

The locations of electropores around the cell membrane are important due to the mechanism which molecules, such as DNA or RNA, are delivered to the cell. The polar locations of where electropores form have been observed in a number of studies, and it is clear that for a cell uninterrupted by surrounding cells or objects that the pores predominantly form on the sides of the cell facing the anode and cathode as described by the cosine factor in the Schwan equation. It has also been apparent that the side of

94 the cell facing the anode porates first and to a greater extent than the side facing the cathode as seen in various parts of Figure 3.9 in particular in parts A,B,C, and D. This comes into play when delivering charged molecules to the cell, including PI, DNA and

RNA. The electrophoretic movement exhibited by charged molecules combined with locations of electropores can influence the success of delivery. For instance a positively charged particle like PI moves from the positive to the negative electrode and predominantly encounters the more permeable anode side of the cell membrane, while

DNA is negatively charged and encounters the less permeable side.

When cells are brought close together as in Figure 3.22 the electroporation axis is shifted slightly away from the electrodes. As a result the formation of electropores takes place at locations slightly off the normally predicted locations facing the electrodes. Instead the pores form slightly facing the other cell it is closest to. Although it is unclear how this may manifest itself in the electroporation process some possibilities could be imagined. For instance, part the delivery mechanisms for certain molecules, like DNA or PI, is a convective flow in addition to electrophoretic movement.

The change in electropore location is unlikely to affect the electrophoretic motion of a molecule in the field as it will follow the electric field lines to the adjusted electropore location. However, any convective flow taking place between the electrodes would seek to carry the molecules and associated momentum strait on at the cell. This could potentially cause lower delivery rates if the locations on the cells the molecules flow to do not match up with the locations of the electropores. At this point these are only speculation as much more would need to be done to demonstrate these effects. This

95 work serves as the first marker showing that the interactions between cells in electroporation can alter the locations of electropores on the cell membrane.

Chapter 4 : Membrane Based Electroporation

4.1 Introduction

Due to the inherent randomness associated with bulk electroporation techniques it has become nearly impossible to achieve low cell death and high transfection efficiency in most practical situations. Random orientation of cells relative to each other and to the imposed electric fields created in a bulk electroporation process give rise to the disparity in electroporation outcomes. Microdevices are a realistic option to overcome randomness and realize the high transfection efficiencies and survival rates required by gene therapy and stem cell clinical protocols.

Microdevices have the ability to control the location and orientations of cells within an apparatus and can affect the electric field by varying geometries that the cells interact with. By treating each cell as uniformly as possible during electroporation it may be possible to achieve an optimal combination of cell survival and transfection. In this chapter fundamental properties of membrane based electroporation microdevices are investigated to provide a better understanding of their mechanisms and how to best take advantage of them in electroporation microdevices. In addition to mechanistic studies using a single pore membrane, two membrane based devices, an ordered

97 micropore array and a novel technique called membrane sandwich electroporation

(MSE), are tested in their ability to deliver genes to cells.

4.2 Single pore electroporation

A micropore membrane concentrates an electric field inside its pore and reduces the area of the cell membrane exposed to the high electric fields in electroporation.

When micropores are used in electroporation the process is sometimes referred to as

“localized electroporation” meaning that the electroporation location is restricted and localized to a specific area of a cell membrane in contact with the micropore. There are several ways to fabricate a micropore for use in electroporation including both cleanroom and non-cleanroom processes. Here we study both fabrication techniques and effects of a micropore on cell membrane breakdown as well as delivery of both large and small molecules into the cell.

4.3 Single micropore materials and methods

4.3.1 Single micropore experimental system

The experimental system consisted of two major groups of equipment including a confocal imaging system, and an electroporation microdevice. The confocal microscope system is composed of a PC , a spin disk system, a laser illumination device, a high sensitivity photo multiplying EMCCD camera, and an inverted microscope. The

98 confocal microscope system was used to capture and process images during electroporation events. The electroporation system was constructed of an electroporation power supply connected to our custom microdevice that contained the single pore membrane. Figure 4.1 is a diagram of these systems and how they interface with each other.

Figure 4.1. The confocal microscope electroporation system

4.3.1.1 Confocal microscope

We used a high speed spin-disk confocal microscope to visualize the processes in micropore electroporation systems. This system utilizes the Yokogawa CSU-22 spin-disk unit with maximum temporal resolution up to 1000 fps. In order to visualize our processes we used a more sensitive but slower frame rate camera with a photomultiplier (Hammamatsu) and at frame rates of up to 100 fps. Two solid state laser lines including 50 mW at 491 nm and the second supplying 25 mW at 561 nm wavelength allowed for precision fluorescent illumination. The lasers could be hardware or software controlled using an AOTF and in our experiments were exclusively controlled by software. An Olympus IX-81 inverted microscope was the base microscope equipped with a 60x, 1.42 NA oil immersion objective and a Jena piezo objective- focusing collar for submicron control of the focal plane images in the z direction. The camera, spin-disk, and other parts of the system were synchronized and controlled using

Vox Cell software from Visitech International.

4.3.1.2 Single micropore electroporation platform

We used a polymer based microfluidic device to interface with the micropore membranes and our power supply to carry out experiments (Figure 4.2). The goals of this device are to enable electroporation while visualizing on the confocal microscope.

This necessitates that the entire device be as thin as possible to remain in the focal limits of the confocal microscope objectives. The base polymer chip was fabricated from

100 a 25 mm by 60 mm blank of 500 µm thick polycarbonate. A computer controlled

Aerotech designed CNC micromilling machine equipped with a 2 fluted carbide steel endmill was used to fabricate this microfluidic interfacing chip. The main channel of the device was 100 µm deep and 30 mm long. Three holes with a diameter equivalent to the width of the channel (1.5 mm) were drilled through the microchannel to provide a fluid entrance and exit on the ends and a membrane mounting interface in the center.

Platinum wire electrodes were inserted into an end port and the reservoir above the membrane during experiments. Clear tape was used to seal the bottom of the channel.

Its transparency allows us to interface the entire system with optical microscopy, and since it is removable we are able to fully clean and reuse the microfluidic chip in multiple experiments. Reusable fluidic chips were cleaned using Alconox cleaning detergent and tap water followed by a triple rinse in deionized water and then were blown dry with pressurized air. Microfluidic chips were typically reused no more than 20 times.

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Figure 4.2. Diagram and photo of the microfluidic chip used for membrane

interfacing

A membrane was mounted on the center hole of the device using double-sided tape (Scotch permanent Cat. 137DM-2) with a 2.4 mm diameter hole removed by a hand punch (Roper Whitney Co, No5 JR hand punch). This provided a water tight seal between the membrane edge and the device. The hole punched in the tape allowed the micropore in the membrane to be the only interfacing layer between the media in the microfluidic channels and the cell containing media above. Once assembled the overall thickness of the device with membrane attached was 700 µm allowing us to use a 60x

102 long working distance objective lens (LUCPlanFLN 60x Olympus) in the confocal microscopy system.

A standard Bio-RAD electroporation unit with modified electrical leads served as the power supply for our electroporation microdevice. The specific Bio-RAD unit was the

Gene Pulser Xcell™ complete with CE module and the optional PC module. Custom electroporation leads were fabricated using a single strand polymer coated electrical wire that terminated in a banana plug on one end that plugged into the electroporation system and an alligator clip termination on the other end that connected to device electrodes. The lead that connected to the top electrode wire was attached to a platinum wire electrode and positioned using a micromanipulation tool capable of controlling the X-Y-Z locations of the electrode (World Precision Instruments model

KITE-R).

4.3.2 Single micropore cell culture

4.3.2.1 NIH-3T3 cell culture

NIH 3T3 fibroblasts (a mouse fibroblast cell line, CRL-1658) were used in single pore experiments. This cell line is a broad based model animal cell line. Cells averaged

15.6 µm in diameter (1.9 µm standard deviation). Cells were procured from American

Type Culture Collection (ATCC, Manassas, VA). The culture media and additives were supplied by Invitrogen (Carlsbad, CA). Cells were grown in nutrient medium consisting of

103

Dulbecco’s modified Eagle’s medium nutrient mix F-12 (D-MEM/F-12, catalog

No.10565), with the addition of 10% (v/v) newborn calf serum (NCS, heat-inactivated, catalog No. 26010), 2 mM L-glutamine (catalog No. 25030), 1 mM sodium pyruvate

o (catalog No. 11360). Cells were cultured in 25 cm2 T-flasks incubated at 37 C under an atmosphere of 5% CO2 . Every 2 to 3 days cells were detached from the surface of the culture flask with a 0.05% solution of trypsin with 1 mM EDTA·4Na (Catalog No. 25300) and placed in fresh media filled flask.

Leading up to an experiment we allowed the NIH 3T3 cells to grow to 90% confluence and then harvested them by trypsinization. We centrifuged cells into a pellet and washed them with Dulbecco's Phosphate Buffered Saline (D-PBS, catalog No.

14190). The pellet was resuspended in Opti-Mem media as commonly used in other experiments, although cells were concentrated by a factor of 5 to a concentration between 105-106 cells/ml. In experiments a small volume of concentrated cells is pipetted into a reservoir of Opti-Mem contained on the top of the single pore membrane attached to the device platform.

4.3.2.2 K562 cell culture

We also used cell line K562 (a leukemia cell line, CCL-243) in the single pore experiments. This cell line was derived from a patient with leukemia and has served as a model cancer animal cell line in this and many other research experiments. Cells were procured from American Type Culture Collection (ATCC, Manassas, VA). Culture media and additives were supplied by Invitrogen (Carlsbad, CA). Cells were grown in nutrient

104 medium consisting of RPMI 1460 (D-MEM/F-12, catalog No. 21870-076), with the addition of 10% (v/v) newborn calf serum (NCS, heat-inactivated, catalog No. 26010), 2 mM L-glutamine (catalog No. 25030), 1 mM sodium pyruvate (catalog No. 11360). Cells

2 o were cultured in 25 cm T-flasks incubated at 37 C under an atmosphere of 5% CO2 . We passaged cells every 2 to 3 days cells by placing a portion in a fresh media filled flask.

Leading up to an experiment we allowed the K562 cells to grow to 80% confluence and then we harvested them by from suspension. The K562 cells have down regulated surface attachment characteristics which make them easy to work with in suspension. We centrifuged cells into a pellet and washed them with Dulbecco's

Phosphate Buffered Saline (D-PBS, catalog No. 14190). The pellet was resuspended and concentrated in Opti-Mem media resulting in a cell concentration between 105-106 cells/ml. In experiments a small volume of concentrated cells is pipetted into a reservoir of Opti-Mem contained on the top of the single pore membrane attached to the device platform.

4.3.3 Delivery molecules

4.3.3.1 pGFP

Reporter plasmid pmaxGFP (3486 bp) was purchased from Lonza (Switzerland).

This reporter plasmid was stained with yoyo-1 iodide (Invitrogen, cat. No. Y-3601) according to protocols described by the manufacturer. The resulting Yoyo-1 stained

105 pGFP molecule was used in experiments to represent a typical large nucleic acid molecule commonly used in electroporation. A 100 mW 488 nm laser was used for fluorescence excitation (Cobolt laser model Calypso).

4.3.3.2 G3139 oligodeoxynucleotide ODN

G3139 ODN is a commercially available FITC stained nucleic acid with a 18 base pare length. The FITC-labeled G3139 (sequence: FITC-5’-TCT CCC AGC GTG CGC CAT-3’) was purchased from Alpha DNA (Quebec, Canada). Although ODN is DNA this nucleic acid is representative of siRNA or other small nucleic acids currently being investigated in gene therapy protocols. A 100 mW 488 nm laser was used for fluorescence excitation.

4.3.3.3 Propidium iodide

Propidium iodide dye was used in experiments conducted with this system. Details of the dye can be found in Section 3.2.2

4.3.4 Single micropore membrane fabrication

We accomplished fabrication of single micropore devices most efficiently using non-cleanroom processes because of their simplicity. The size of the micropores does not necessitate cleanroom processing including photolithography and as a result the cost and ease to fabricate the single micropore membranes is significantly reduced and easily accessible. The base material for the single micropore membranes was a 12 µm

106 thick polyester membrane known by the trade name Mylar™ (McMaster Carr). Here we show two non-cleanroom approaches for fabrication including micromilling and laser ablation.

4.3.4.1 Milling

We used a micromilling technique to produce a single microhole in the Mylar™ membrane material. The micromilling system was developed by Aerotech with computer controlled G-code programming. Milling resolution is controlled by the size of the end mill and the motion control system of the milling machine. The mill has resolution in the x-y-z plane down to 100 nm and endmills used in membrane fabrication were 7 and 10 µm diameter (PMT microtool). During milling the Mylar™ film was placed on a double-sided tape (Scotch permanent Cat. 137DM-2) and mounted to the motion control stage. A camera was used to visualize the endmill in real time as it was brought to the surface of the membrane using small precisely controlled movements using the software. Once at the surface the software was able to zero the X-

Y-Z coordinate system to prepare for CNC milling. The milling speed was set via software at 10,000 rpm and moved at a speed of 1 mm/min when in contact with the membrane.

Once the coordinate system and program was set the endmill was controlled via computer to mill through the Mylar™ and produce a single micropore.

4.3.4.2 Laser

As an alternative to micromilling techniques a femtosecond laser system with motion control stage (Figure 4.3) was used to ablate the surface of the Mylar™ film to

107 produce a single micropore. By varying the laser power we were able to produce a smaller or larger diameter pore. The benefits of a femtosecond laser pulse are its ability

16 to produce very high peak intensity ( 10 W/cm2) and rapid deposition of energy into the material. Difference between a long laser pulse and femtosecond laser pulse on material removal and heat transfer is shown in Figure 4.4.

Figure 4.3. Femtosecond Laser System (Farson, et al., 2008)

108

Figure 4.4. Thermal and surface effects of long wave (a) vs femtosecond laser (b) pulsing (http://www.cmxr.com/Industrial/Handbook.htm)

109

Membranes were mounted to the edge of a ring platform using double-sided tape (Scotch permanent Cat. 137DM-2). The laser was focused on the surface of the membrane and a computer controlled shutter was used to expose the film to the laser pulses. The membranes were exposed to the laser for 100 ms resulting in 3000 laser pulses each at a duration of 150 femtoseconds.

The femtosecond laser system produces frequency doubled pulses from a mode- locked erbium-doped fiber laser intensified in a Ti:Al2O3 regenerative amplifier laser

(CPA2161, Clark-MXR). The maximum output power of the laser is Pav=4.5 W. The laser is capable of a pulse duration is Tp=150 fs, with pulse repetition frequency was fP=3 kHz, at a wavelength λ of 775 nm, and beam diameter of 5 mm. We adjusted the laser beam power by a series of optics, including thin-film polarizing beam splitters and ½ wave plates (Figure 4.3). The attenuated laser beam was delivered to the sample by a series of beam mirrors that direct the beam into a mechanical shutter and then focused on the material using a 50x infinity corrected microscope objective lens with numerical aperture—NA= 0.42 (M plan Apo NIR 50x, Mitutoyo) beam. Attenuated laser power was measured by a power meter (PM100, Thor Labs) placed under the laser focusing lens.

The beam quality in our experiments was M2=1.2 in the horizontal Y direction and

M2=1.3 in the horizontal X direction. A computer controlled (G-code) motion system

(MX80L, Parker) with a 0.5 μm resolution in the X, Y, and Z axes was used to position the sample and focus optic. For the consistency and easy focusing purpose, a coaxial vision system was installed, allowing the material to be visually located at the focus within ±1

μm range where focal depth was 1.6 μm for the selected optics.

110

4.3.5 Single micropore device procedures

During operation electroporation media was introduced into the under-channel containing the electrode via micropipette. Opti-MEM (Gibco Cat.No. 11058 ) supplemented with PI to a total concentration of 100 mM PI dye or with 2-4 µg/100 µl of DNA, either yoyo-1 stained pGFP (3.5 kbp) or FITC coupled ODN (G3139, 18 bp), served as the electroporation media in these experiments. The dimensions of the microchannel enabled capillary forces to quickly draw the media through the channel.

Once the under-channel was filled a 20 µl drop of plain Opti-MEM solution was pipetted over the micropore. Next, 10 µl of low concentration cells in Opti-MEM media (1x104 cell/ml) were pipetted into the 20 µl plain Opti-MEM droplet on top of the micropore membrane. A 1-3 mm differential in fluid height between the top of the cell reservoir droplet and the media in the microchannel below created a pressure differential sufficient enough to cause flow through the micropore and draw and trap a cell over top of the pore. Once the cell was in place the top electrode was positioned using a micromanipulator (World Precision Instruments model KITE-R) and the cell was electroporated using a Bio-RAD gene pulser Xcell power supply with custom leads to attach to the platinum electrodes.

In membrane breakdown experiments to determine the critical electric field, the electroporation apparatus started at a low electric field and was stepped up by a 2-4 volt (8-12 V/cm) increment. Pulse parameters were set at a single 1 ms. During an electroporation event the confocal microscope Vox Cell software coordinated video

111 capture at the following settings capable of detecting PI fluorescence from electroporation: shutter speed of 100 ms, binning 1x1, gain 150, 25 mW 561 nM laser power. 60 s elapsed between an increase in the electric field to allow PI detection. Once

PI fluoresce was detected the electric field was noted and the experiment was over.

The PI diffusing curve experiment was conducted in a similar way as the membrane breakdown experiments with a few exceptions. The electroporation power supply used in these experiments was the Axoporator 800A system. In these experiments a single pulse was used and length varied depending on the experiment and was set at 1 or 100 ms to understand the relationship between the pulse length and possible delivery mechanisms. PI fluorescence was recorded and analyzed using

Metamorph software. Fluorescence data was normalized as the intensity average over the entire cell at a given time divided by the maximum fluorescence intensity achieved in the experiment.

In gene delivery experiments cells were captured on a 4 µm pore in the same system as the PI dye experiments. The electroporation media underneath contained either ODN (18 bp) or pGFP (3.5 kbp) at a concentration of 3-6 µg/100 µl.

Electroporation pulses consisted of three individual 70 V/cm pulses each a duration of

100 ms spaced at a 10 Hz frequency for pGFP and a single 20 ms 200 V/cm pulse for

ODN. The VoxCell software coordinated video capture during electroporation at the following settings: shutter speed 200 ms, 2x2 binnging, 150 gain, 50 mW 488 nm laser power. Immediately post electroporation the VoxCell software was programmed to

112 execute a Z-stack image capture to produce a 3D profile of the cell on the micropore.

The Z-stack capture was subsequently executed to produce a delayed time series of 3D images for comparison and analysis.

4.3.6 Single micropore modeling

We modeled the membrane breakdown in a single micropore using the same method described for single cell modeling in bulk electroporation (3-layer shell) with some parameter adjustments to match our experimental system. Values of conductivities of intercellular cytoplasm, external surrounding medium were both

7  i   e  0.2 S/ m, while the cell membrane was  m  510 S/m The radius was assumed R  6 m and the membrane thickness is d  5 nm . The external bulk electric field strength was adjusted based on experimental results.

4.4 Single micropore results

4.4.1 Single micropore fabrication

4.4.1.1 Micromilling

Micromilling for micropore fabrication is relatively low cost compared with clean room processing and is applicable to a wide range of materials including polymers and metal films. For the electroporation applications with K562 and NIH-3T3 cells, we

113 desired a micropore that was small enough to trap any size cell present in the population while providing significant reduction in the applied electric field. The cell size in the two cell lines grown for these experiments varied between 8-18 µm and was typically 12-14 µm. Therefore it was necessary to create a pore that was 10 µm or less for effective trapping. 7 and 10 µm 2 fluted carbide steel endmills were selected in the fabrication process to create the micropores. The 7 µm endmills were repeatedly damaged in the milling process and as a result were only able to make a 16 µm pore in the membrane using this endmill. This repeated result of a pore with larger diameter than nominal endmill indicates that it was too fragile to do this micromilling and would be damaged in the process. The 10 µm endmill was capable of repeatedly and consistently producing a 10-12 µm pore in the membrane without damage to the endmill. The perimeter shape of the micropore was nearly a perfect circle as seen in

Figure 4.5 but it had significant surface roughness and debris present around the edge of the perimeter and inside of the pore from the milling process. Front and back scanning electron microscopy images (Figure 4.5) show the same pore size indicating pore geometry was strait walled all the way through the membrane. This micropore membrane shows the limits of what is possible when fabricating micropores with traditional milling processes. Endmills smaller than 10 µm have resulted in irreparable endmill damage and larger pores indicating that we are very near if not at the limits of this fabrication methods ability to make a suitable micropore for our studies.

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a b

10µm 10µm

Figure 4.5. SEM images for the 10µm milling bit (a) front and (b) back of the membrane

4.4.1.2 Femtosecond laser

The femtosecond laser system is capable of producing various pore geometries and sizes by varying the laser power. Figure 4.6 outlines the effects of varying the laser power to produce the smallest possible micropores capable of this system. Based on

SEM images from the front and back of the membrane it is apparent that one side of the membrane has a smaller pore than the other indicating the pores are conically shaped.

A laser power of 1.5 mW was unable to produce a pore all the way through the membrane under the 1 second exposure time used here. Pores were produced with laser powers beginning at 2 mW with diameters of 3 and 4 µm and remained conical in geometry, indicated by the SEM images, all the way up to 3 mW. Laser powers of 3 mW and above made larger pores, in excess of 10 µm, that were less effective in maximizing the electric field focusing effects and cell trapping capability. A 2.5 mW laser power was optimal to fabricate membranes used in our experiments and created an average pore diameter of 4 µm. This laser power ensured repeatability and provided a larger margin of error when focusing the laser on the membrane which is a manufacturing speed and 115 scale advantage. The pore diameter it created also was small enough to trap a wide range of cell sizes and shapes. It can be seen in Figure 4.6A that pore size rapidly increased between 2.5 and 3 mW indicating that 2.5 mW provided an optimum combination of penetration power with a small pore size. Laser ablation created surface roughness on all pore sizes as seen in the scanning electron micrographs (SEM) of Figure

4.6. This figure also indicates pore shape irregularity from both jagged ablation debris and ellipsoidal laser beam shape. In comparison with milling surface roughness of the laser fabrication process produced a much less jagged and intrusive micropore.

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A B

C 2mW 2.5mW 3mW 4mW

2mW 2.5mW 3mW 4mW D

5µm 5µm 10µm 10µm

2mW 2.5mW 3mW 4mW E

5µm 5µm 10µm 10µm

Figure 4.6. Femtosecond laser microfabrication results. (A) large and small opening vs laser power (B) membrane cross sectional diagram (C) bright field images (D)SEM micropore images of the front, and (E) SEM images of the back of the membranes

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4.4.1.3 Cell interactions the micropore

We were able to visualize a cell’s interaction with the micropore membrane by first staining it with calceian AM dye and then trapping it on the membrane in our device. A z-stack image analysis was done on two areas of the device; the first was above the cell down to the surface of the membrane followed by imaging the area through the membrane. These two image stacks were reassembled in parts using Imaris image analysis software to produce the images in Figure 4.7. In the 4 µm pore (Figure

4.7(a)) we can see that there was a fluorescence cloud above the membrane indicating where the cell was located outlined by the white line. A small portion of fluorescence was seen just under the cell in the membrane layer image indicating a small portion of the cell was inside the pore. This image indicates that a cell initially seals the area around the pore with some portion of the cell inside the micropore. With a 12 µm pore

(Figure 4.7(b)) we found that a majority of the cell is located inside the membrane area with a portion of the cell still outside the membrane.

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Figure 4.7. A calcein stained cell interacting with a (a) 4 µm or (b) 12 µm micropore

4.4.2 Micropore critical electric field

The micropore critical electric field studies are a continuation and comparative study to the critical electric field studies done previously with optical tweezers in the bulk electroporation system. In these studies optical methods using PI were again used to determine and confirm the critical membrane breakdown electric field (Ec).

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4.4.2.1 Single pore PI dye delivery

In the K562 experiments using propidium iodide (PI) dye a significant reduction in the critical electric field (Ec) compared to bulk electroporation was observed to produce electroporation as indicated by PI dye entrance. A set of 6 experiments (n=6) were averaged to determine the critical electric field value of 380 V/cm with a standard deviation of 30.3 V/cm using a 12 µm membrane produced by the micromilling method.

A 10 µm pore femtosecond laser fabricated membrane had a lower critical electric field of 340 V/cm with a standard deviation of 28.2 V/cm (n=9). Comparing these two sets of experiments using a two tailed statistical T-test (equal variances) with a null hypothesis that the E c of these populations is the same (Ec1=Ec2) and an alternative hypothesis that the Ec of these two populations are different (Ec1≠Ec2) we conclude that these two membranes produced a statistically different Ec. Statistically there is less than a 4% chance (α=.036) that they produced the same Ec. When the membrane pore size is reduced to 4 µm the Ec required for electroporation was reduced to 190 V/cm with a standard deviation of 23 V/cm (n=6). A two tailed equal variance statistical T-test also indicated that the 4 µm pore produced a statistically different Ec value than the 10 or 12

µm pore sizes with a less than 1% chance that the membranes were statistically indistinguishable. Figure 4.8 summarizes the critical electric field results with K562 cells and compares them to previous results obtained using optical tweezers in bulk electroporation. It can be seen that the membranes produce a 45%-72% reduction in Ec and that there is a trend of a greater reduction with smaller pore size.

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Figure 4.8. K562 critical electric field using micropore membranes

The same experiment conducted on NIH-3T3 cells yielded similar results. The 10

µm membrane produced a 47% reduction in Ec while the 4 µm membrane produced a

67% reduction in Ec based on Figure 4.9. The Ec using the 10 µm membrane was 270

V/cm with a standard deviation of 13.5 V/cm (n=9) and with the 4 µm membrane it was

170 V/cm with a standard deviation of 19.7 V/cm (n=8). Results also indicated that the membranes produced statistically significant differences in required Ec by T-Test. T-test results indicate a less than 1% chance that these two membranes produced the same Ec using a two tailed test with unequal variances (null hypothesis µ1=µ2, alternative hypothesis µ1≠µ2). It is also noted that the critical electric field with 3T3 cells was always lower than with K 562 cells in the same conditions.

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Figure 4.9.Comparison of micropore membrane electric field with bulk electroporation for NIH-3T3 cells

4.4.2.2 Single pore models

In the first set of models (2D calculation) we assume that there is no gap between the cell membrane and the edge of the micropores. As seen in Figure 4.10(a), Figure

4.11(a) and Figure 3.13(a) the electric field distribution inside the micropore is very high compared to the bulk area above the microchannel. Using the Ec values from the K562 micropore critical transmembrane potential experiments Figure 4.10(b), Figure 4.11(b), and Figure 4.12(b) show the predicted transmembrane potential on the cells surface. As a result of the high electric field inside the micropore Figure 4.10(b), Figure 4.11(b), and

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Figure 4.12(b) show the maximum transmembrane potential on the cell membrane occurring in the areas of the cell membrane that are directly over the pore surface, while the rest of the cell membrane experienced a relatively low transmembrane potential and electric field strength. We can see that all of the results predict critical transmembrane potential values (19 V, 32 V, and 29V) that are in the same range and fairly consistent across the different micropore sizes.

In the next model (2D calculation) we used the same 4 µm pore model as in

Figure 4.10 but we assumed that there is an imperfect seal between the pore and the cell membrane creating a 500 nm gap between the pore edge and the membrane

(Figure 4.13). In Figure 4.13(a) we can see that the electric field lines passing through the gap area are fairly concentrated causing a leak current around the cell. This leak results in a significant drop (compared to Figure 4.10(b)) in the maximum transmembrane potential from 19 V to a value of 6 V as seen in Figure 4.13(b).

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10-5 a

Surface ElectricSurface FieldV/cm

Length (mm)

10-5 Length (mm)

∏

Cell

θ membrane 0

Cell membrane location

Figure 4.10. (a) Electric field distribution and (b) transmembrane potential plot for the 4 micron pore with the cell located on the membrane surface

124

10-5 a

Surface ElectricSurface FieldV/cm

Length (mm)

10-5

Length (mm)

∏

Cell

θ membrane 0

Cell membrane location

Figure 4.11. (a) electric field lines and (b) transmembrane potential for the 10 micron pore with cell located on the membrane surface

125

10-5 a

Surface ElectricSurface FieldV/cm

Length (mm)

10-5 Length (mm)

∏

Cell

θ membrane 0

Figure 4.12. (a) electric field lines and (b) transmembrane potential for the 12 micron pore with cell located on the membrane surface

126

10-5 a

Surface ElectricSurface FieldV/cm

Length (mm)

10-5 Length (mm)

b ∏

Cell

θ membrane 0

Figure 4.13. (a) Electric field lines and (b) transmembrane potential for the 4 micron pore with the a 500 nm gap between the cell membrane and the pore edge

127 a

∏

Cell

θ membrane 0

4.4.3 Single pore delivery mechanisms

4.4.3.1 PI delivery

We compared the increase in fluorescence intensity for bulk electroporation and

micropore electroporation. For bulk electroporation with a 1 ms pulse length we found

that the PI fluorescence saturated at around 50 seconds, while when using a 4 µm pore

with a 1 ms length we see the cell saturated at about 30 seconds. As we further

increased the pulse length to 100 ms using the micropore we see the micropore system

saturated in about 15 seconds. Results are summarized in Figure 4.14, and it should also

be noted that the electroporation pulse started at t=0 on each of these graphs. In

micropore electroporation the fluorescence intensity increased almost immediately,

while bulk electroporation typically exhibited a “diffusion lag” between the onset of the

pulse and increase in PI fluorescence. Apart from the “diffusion lag” the rest of the bulk

electroporation process was controlled by diffusion and it was apparent that the

micropore curves took on a similar shape to the bulk electroporation curve indicating

that they are all likely diffusion controlled processes.

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Figure 4.14. Bulk and micropore PI fluorescence curves after electroporation at (t=0)

Confocal images of PI delivery using a 4 µm pore are shown in Figure 4.15. This k562 cell was subject to an 800 V/cm pulse for 5 ms using our experimental system.

Images a, b, and c in this series provide a 3 dimensional view of the fluorescent locations during electroporation. The strongest fluorescence takes place directly above the micropore and slowly decreased throughout the cell. The cell became saturated with dye in the first 20 seconds after electroporation as shown in Figure 4.15 d),e), and f) and with the strongest fluorescence still exhibited directly above the pore inside the cell due to high concentrations of DNA and PI dye located here.

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e f

Figure 4.15. Confocal images of PI dye delivery to a K562 cell at 10 seconds post electroporation from the a) top, b) front, and c) side and 20 seconds post electroporation from the d) top, e) front, f) side.

4.4.3.2 ODN and pGFP delivery

Both small and large DNA were used in gene therapy protocols, however they behave very differently biologically and in their chemical properties in solution. Results of gene delivery using our single micropore are presented in Figure 4.16 and Figure 4.17.

They show that basic mechanisms in bulk electroporation of large DNA remains the same, while the small DNA are capable of quickly diffusing into the cell. Confocal images from ODN delivery initially show that the DNA coats the outer surfaces of the cell membrane as shown in traditional bulk electroporation. This was observed in Figure

4.16 when looking at the bottom view of the cell through the micropore the surface was

130 entirely fluorescent. From the front and back views we can see that there was no fluoresce covering either side of the cells in these locations and fluorescence was only exhibited on the edges of these locations. As time progressed 10 minutes and we looked at the same views of this cell we can see that the ODN fluorescence had permeated the cell from all sides and previously dark areas of the cell had become fluorescent on the front and back. Results indicated that the ODN was able to freely diffuse into the cell cytoplasm using the micropore to aid in delivery.

When looking at the much larger pGFP (3.5 kbp) DNA delivered in bulk electroporation it was apparent from Figure 4.17(a) and (b) that the pGFP was located on the surface of the cell membrane. This was also the case in micropore electroporation. Confocal images of the cell 15 minutes post electroporation indicate that the pGFP had not penetrated the cell membrane as indicated by the planar fluorescence on the edge of the cell in Figure 4.17c,d,e. It should also be noted in these images that the DNA coats the cell in locations around the cells edge outside the micropore. This provides evidence that there is in fact a leak from an incomplete seal between the micropore and the cell membrane where the nucleic acids (pGFP and ODN) are able to coat the cells surfaces outside the micropore.

131

Figure 4.16. ODN delivery through the 4 micron pore immediately after and 10 minutes after electroporation.

132 a b

Figure 4.17. Fluorescence images of YOYO-1 stained pGFP delivered to a cell in bulk electroporation a) before electroporation, b) after electroporation and confocal images of yoyo-1 stained pGFP delivery to a cell using a micropore 15 minutes post electroporation from the c) bottom, d) side, and e) front of the cell. The white circle indicates the approximate location of the cell, and the squares indicate the approximate location of the micropore (4 µm) membrane.

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4.5 Single pore discussion and conclusions

Non-cleanroom fabrication techniques used to make a single micropore have shown to be successful down to 3-4 µm. Between the two techniques used here it is apparent that the femtosecond laser provides greater versatility over milling techniques when making electroporation membranes for immobilizing a cell and carrying out electroporation. The laser fabrication is capable of producing a smaller pore size than the micromilling methods with reduced amounts of potentially interfering surface roughness and debris. The laser also has shorter cycle times and can produce membranes in a shorter time period. Since the laser does not have tip durability issues it lends itself to the capability to produce many micropores in a single membrane. In nearly any situation the laser fabrication technique has advantages over the micromilling technique.

When comparing micropore electroporation with bulk electroporation it is apparent that a reduction in the bulk electric field takes place when using micropores from Figure 4.8 and Figure 4.9. This phenomenon occurs regardless of cell type and is due to the electric field focusing effects caused by the micropore constriction. Electric field enhancements are achieved by reducing the area that the electric field passes through. In both situations we see on average a somewhere between a 45-50% decrease in electric field for the 10 µm membrane and a 70-72% reduction for the 4 µm diameter pore.

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The models of our micropores under experimental conditions indicate a very high transmembrane potential inside the micropores before the membrane porates. We would like to note that although these transmembrane potential values are high compared to reported values for bulk electroporation by other research groups and our own results presented in Chapter 3 (i.e. ~1 V instead of 19~32 V), they have similar values for different micropore sizes indicating that any parameters that are unaccounted for in our model seem to apply equally to all micropore experiments. For instance, the model assumes a perfect micropore and the pore is in close contact with the cell in a spherical shape. This idealized case is not what observed in the experiments where the micropore is not complete circular and flat, and the cell is somewhat deformed around the pore. The cells are not fully contacting and covering the entire gap over the pore allowing a leak current to pass around the cell. Surface roughness from laser ablation and irregular shaped pores from laser ablation and milling may contribute to a leak current. When thinking in an electrical circuit analog the cell provides a large resistance in parallel with a small resistance representing the leak current. As shown in

Figure 4.14(b), the calculated critical transmembrane potential drops from 19 V to 6 V by just considering a 500 nm gap between the pore edge and the cell. There are also possible electrolysis bubbles that could present additional resistances and soak up voltage and current throughout the system. If this and other factors are considered in the model, the calculated value could be lower. In fact, if we divide our critical transmembrane potential values by a factor of 20 we get the respective transmembrane potential values of 0.95 , 1.55 , and 1.45 V for the 4, 10, and 12 µm pores, which are

135 typical critical transmembrane potential values (0.8-1.5 V) expected to cause poration in the bulk electroporation where electric fields are very high. It is apparent that these unaccounted for resistances are uniform from experiment to experiment since results were repeatable and populations exposed to the smaller pore membrane were statistically distinguishable from the larger pore.

With micropores we are able to achieve the critical transmembrane potential in a small area on the cell membrane with the rest of the cell experiencing a much lower electric field strength and transmembrane potential.

The micropore DNA experiments show that there is some leakage happening during the electroporation process. It is common during the electric pulse for the cells to rattle or move slightly. This movement can be cause by pressure created from electrolysis bubbles, from cell expansion that takes place during electroporation, or from an electrophoretic force on the cell caused by the electric field. Figure 4.16 and

Figure 4.17 show that fluorescence from the nucleic acids cover surfaces of the cell that are outside the micropore which is most likely due to a small leakage of fluid between the pore and the cells surface.

Beyond a reduction in critical electric field the micropores enhance the diffusion of materials into the cell. The diffusion time of PI dye into an electroporated cell is reduced when using a micropore. The enhanced electric field inside the pore exerts a greater electrophoretic force on charged particles like PI or nucleic acids resulting in acceleration of particles through the micropore into the electroporated cell membrane.

136

The time series diffusion curves done with PI dye show that the diffusion lag period is eliminated using a micropore. This can be explained by forces the electric field exerts on the charged particles like PI or nucleic acids. PI molecules near the pore get drawn and directed to the micropore area during the electroporation pulse resulting in an enhanced concentration of PI near the electroporated area on the cell membrane. After the pulse diffusion takes over and the fluorescence curves look similar to bulk electroporation without the diffusion lag period in the beginning. These effects can also be seen in Figure 4.15 with two consecutive 3D confocal images of a cell that received PI dye through a 4 µm pore. It is apparent that dye concentration is highest around the micropore itself and that within 20 seconds the dye has saturated the cell with fluorescence as opposed to 30-90 seconds needed in bulk electroporation. These results typically indicate a 1.5-3x enhancement over bulk electroporation and should likely be characterized as an enhanced diffusion. The micropore enhances the electric field near the cell surface causing concentration and acceleration of dye or DNA toward the cell membrane much like a funnel, however once the pulse is gone diffusion takes over. The increased concentrations lead to a diffusion enhanced delivery in micropore situations.

4.6 Micropore array

Moving away from more fundamental single cell micropore studies we sought to test practical implementation of micropore systems to make the electric field more uniform. The most obvious implementation of micropore technology is to use porous membrane to trap cells onto and carry out electroporation. We developed a simple

137 microdevice that implements a uniform ordered array of micropores to carry out electroporation on a mass number of cells at once. The goals of this device were to demonstrate the potential for mass delivery of genes using a micropore technology and to make this process as simple and uniform as possible in order to ensure all of the cells receive the most uniform treatment to result in higher transfection efficiency and lower cell death. We also aim to compare cleanroom and non-cleanroom fabrication techniques capable of producing these membranes.

4.7 Micropore array materials and methods

4.7.1 Micropore array experimental system

The experimental setup for to study the micropore array gene delivery system was the same as used in single micropore studies as described in Section 4.3.1.

4.7.2 Cell culture

We used NIH-3T3 cells in these studies and cell culture practices were previously described in Section 4.3.2.

4.7.3 Green fluorescent protein plasmid (pGFP)

We purchased gWizTM green fluorescence protein vector (GFP, 5757 bp) from

Aldevron (Fargo, ND), and purified it with an EndoFree Plasmid Maxi Kit from Qiagen

(Valencia,CA, USA) according to the manufacturer’s instructions. pGFP is a reporter

138 plasmids commonly used to indicate successful transfection. It is sometimes coupled with other genes and used to indicate if a cell was genetically transformed, however here we are simply using it to indicate that the cells were successfully transfected and are healthy enough to produce a green fluorescent protein from the pGFP we delivered.

The plasmid GFP is not itself fluorescent but it is genetic material that is used by the cell like an instruction book to build a green fluorescent protein molecule.

4.7.4 Micropore array device procedures

The fluidic device for ordered array membrane experiments was as described for the single pore experiments with the following vacuum capabilities as seen in Figure

4.18. The vacuum was connected through flexible PVC tubing to a hose barb fitting that was glued onto the top of the microfluidic chip.

139

Figure 4.18. The microfluidic device used for micropore array membrane interfacing

During operation electroporation media is introduced into the under-channel containing the electrode. Opti-MEM (Gibco Cat.No. 11058 ) supplemented with

2µg/100µl of DNA, either yoyo-1 stained pGFP or plain pGFP served as the electroporation media in these experiments. 30µl of cells in Opti-MEM media (1x105 cell/ml) are placed on top of the micropore array membrane and a differential pressure is created between the top of the cell bubble and the media in the microchannel below by pulling a vacuum while plugging the electrode channel opening. This is sufficient enough to cause flow through the micropore and draw a cells over each pore. The vacuum was controlled between 1-30 inHg using a vacuum regulator gauge and valve and was typically set at 10 inHg vacuum pressure. Once the cells are in place the top

140 electrode is brought in place and the cells are electroporated using an Axoporator 800A power supply with custom leads to attach to the platinum electrodes. The electroporator was set to deliver five DC pulses of 300 V/cm pulses for duration of 500 ms each at a 10 Hz frequency.

4.7.5 Micropore array membrane fabrication

Beyond understanding mechanisms using a single pore membrane we demonstrated how to implement a micropore gene delivery technique. There are major fabrication differences to consider when fabricating a micropore array as opposed to a single pore that we address here. Micromilling becomes impractical to directly produce large arrays, or even molds for large arrays of micropores due to development time and the delicate nature of small milling bits. Here we work with cleanroom and non- cleanroom techniques to produce large arrays of micropores.

4.7.5.1 Cleanroom

The cleanroom membrane fabrication techniques involve standard soft lithography and are based on using NANOTM SU-8-25 photoresist and SU-8 developer from Microhem Corporation. (Newton, Massachusetts). This photoresist is an octa- functional epoxy resin and the SU-8 developer is propylene glycol methyl ether acetate

(PGMEA). The resist is coated on 100mm diameter (100) P type silicon wafers and exposed to UV light through a photomask to create a master mold. The photomask is an

141 array pattern of 5 um diameter of dark-field with circles that fill a 5 mm × 5 mm area of the 120x120 mm chrome on quartz mask. Circles are spaced 16 µm on center. The pore density is designed to accommodate tight packing of a monolayer of cells without cells accessing two pores at once or interrupting one another.

142

Silicon wafers (100 mm diameter P-type) served as the base media to pattern

SU-8 upon and are cleaned by immersing them in acetone for 3 minutes, and rinsed in isopropyl alcohol for 3 minutes. We dried the cleaned wafers with filtered nitrogen gas and placed them on a hot plate at 200◦C for 10 minutes to dehydrate. Next we spin- coated SU-8 25 onto the cleaned wafers at varying revolutions per minute (rpm) to optimization the SU-8 thickness (Figure 4.19). A 15 um thickness is ideal for the purpose of our fabrication to enable resulting membranes to have sufficient thickness and flexibility while allowing more obtainable photolithography results due to difficulty of high aspect ratio features.

Figure 4.19. spin speed vs. SU-8 25 photoresist thickness

Once coated, wafers were placed in the oven for soft baking. Pre-exposure soft baking was done at 65 ◦C for 1 hour and 95 ◦C for 3 hours on a well-leveled surface in the

143 oven. We slowly cooled wafers to room temperature after soft baking and cleaned the backside of the wafers a small amount of acetone on a cotton swab in order to obtain a flat surface capable of providing good contact between the mask and wafer.

SU-8 exposure was controlled using a EV Group EVG 620 Aligner with the circle array photomask using 365nm light (15mW/ cm2). UV exposed photoresist areas cross- link and become insoluble to the photoresist developer, while the unexposed portion of the photoresist is dissolved by the developer. We varied UV exposure from 11s to 21s based on source intensity measurements taken with an i-line (365nm) radiometer and probe device caused exposure energy to range from 165 to 315 mJ/cm2. We evaluated the solubility of the exposed resist in the developer by checking the pattern structures with a microscope and checking the thickness with DekTak profilometer. Exposure energy of 250 to 300 (mJ/ cm2) produces a thickness of 15um. A standard light microscope was used to check for over-exposure cases. The result showed exposure energy of 285 (mJ/ cm2) had the best pattern structures the required thickness.

After exposure, post exposure bake was done at 65◦C for 1 minute and 95◦C for 3 minutes. The samples were slowly cooled to room temperature for 5 minutes to release the residual stress. Next, the samples were immersed in SU-8 developer (MicroChem) for 6 minutes at room temperature. Following development, the patterned wafers were rinsed briefly with isopropyl alcohol (IPA) and then dried with a gentle stream of compressed nitrogen. Next we loaded the samples in an oven for a one hour at 150 ◦C hard-baked. The developed SU-8 microstructures were platinum coated for imaging using scanning electron microscopy (SEM) shown in Figure 4.20.

144 a

Figure 4.20. (a) Top and (b) 45 degree SEM of the developed SU-8 25 photoresist

The microstructures created on the silicon wafer via SU-8 served as a mold for

replication using polydimethyl siloxane (PDMS) (Sylgard 184, Dow Corning). The general

membrane fabrication steps are illustrated in Figure 4.21. We mixed a curing agent and

prepolymer at a 1:10 weight ratio then poured mixed but uncured polymer over the

silicon/SU-8 microstructure mold inside a Petri dish to a 10mm depth. The sample cured

at room temperature for 24 hours. Initial molding attempts were unsuccessful at

145 precisely replicating the micropillars in the PDMS. SEM was used to examine the PDMS mold. Figure 4.22 details complete and incomplete PDMS micropillars. The molding process was modified to identify potential microstructure filling problems. The uncured

PDMS mixture of 10:1 prepolymer with curing agent was degassed at 4 Pa for 30 minutes to remove air bubbles poured onto the silicon/SU-8 master. The PDMS prepolymer mix was degassed for another 30 minutes at 4 Pa while in contact with the master mold in a Petri dish to allow the PDMS mixture to fill the microstructures. The

PDMS prepolymer was cured at 65◦C for 16 hours and the cross-linked PDMS was peeled from the silicon/SU-8 mold. Figure 4.23 details the new processes molding results.

Figure 4.21: Soft lithography micropore array membrane fabrication diagram

146

Figure 4.22. Broken micropillars in PDMS molding

Figure 4.23. Mold replication with modified processing conditions

147

We used the finished PDMS mold to fabricate micropore array membranes using a polymer dewetting process. A 16% by weight solution of polycaprolactone (PCL)

(sigma) was dissolved in anisol (fluka) solvent. This solution was spun across the PDMS mold with varying spin speeds from 2500 to 5000 rpm for 60 seconds to achieve a desired thickness and sufficient dewetting (Figure 4.24) for resulting membranes to have durability in our electroporation processes.

Figure 4.24. Membrane thickness vs spincoat speed (rpm)

The resulting PCL membranes were removed from PDMS mold with a tweezers and measured for thickness by a profilometer (Detek). A schematic of the PCL membrane fabrication process and 2-dimensional light microscopy images of PCL replication processes are shown in Figure 4.25. Figure 4.25 (A) shows incomplete dewetting of a 16 wt.-% polymer solution (at 3000 rpm) around the pillars with. As the 148 spin speed increased to 5000rpm the dewetting is complete. Figure 4.25B shows that the polymer is only covering holes under the 5000rpm processing conditions.

Figure 4.25. Schematic and light microscopy images of (A) incomplete (3000rpm) and (B) complete (5000rpm) processing conditions for PCL membrane fabrication.

4.7.5.2 Femtosecond laser

The femtosecond laser can also be used to fabricate a micropore array. They system used is identical to the single pore femtosecond laser system. In this new fabrication setup a more advanced G-code program was created to guide the motion control system to create an array of micropores. Mylar film was used for the base membrane substrate to create the array on. For each pore the membrane was exposed to a 2.5mW pulse for 1 second from the femtosecond laser. This laser power was chosen

149 based on results from Figure 4.6 to enable a complete micropore to be made with some leeway in the beam focusing location in the case that the focal location would shift.

4.8 Micropore array results

4.8.1 Micropore array fabrication results

4.8.1.1 Micropore array surface quality and cell trapping

The laser fabricated and soft lithography fabricated membranes are pictured in

Figure 4.26. Pore sizes on both membranes were fairly similar, however pore spacing on the laser me membrane was set at 30 µm on center, while the soft lithography membrane was set to 20 µm on center. Cell trapping done with NIH-3T3 cells can be seen in Figure 4.26 (c) and (d) and it was apparent that both membranes produced a fairly uniform monolayer of cells trapped on the surface; however the cell packing density was much higher in the soft lithography membrane using the 20 µm pore spacing.

Surface roughness of the area around the micropores is important to ensure uniform process conditions between cells on different micropores in the array. SEM

Detail of the micropores is pictured in Figure 4.27. We can see that the membrane surface of the laser fabricated membranes was smooth except outside of the pore area, however around the edges of the pore there are random jagged defects from ablated material that deposited near the pore edge. The soft lithography fabricated membrane

150 was almost the opposite. The membrane area between the pores had some slight dimpling while the area around the edge of the pore was extremely smooth and had little roughness present.

a b

50 µm 40 µm

c d

60 µm 100 µm

Figure 4.26. (a) Bright field image of a femtosecond laser micropore array membrane and (b) a soft lithography micropore array membrane. (c) Bright field image of a femtosecond laser micropore array membrane with cells trapped and (d) a soft lithography micropore array membrane with cells trapped.

151

30 µm 5 µm

Figure 4.27. (a)array detail and (b) pore detail SEM images of a femtosecond laser fabricated micropore array membrane and (c) array detail (45 degree) and (b) pore detail SEM images of a soft lithography fabricated micropore array membrane.

4.8.2 Micropore cell interactions and transfection

Cells were dyed with calceian AM dye to visualize their interaction with the membrane. A standard dose of Calcian per manufacturer’s instructions was given to

NIH-3T3 cells to enable them to fluoresce under 561nm laser excitation. A PCL micropore array membrane was mounted on the device platform (Figure 4.18) and cells

152 were trapped using suction pressure (Figure 4.28). Figure 4.29 shows 3D confocal images taken of the cells interacting with the membrane once trapped. We can see that the cells line up on the array and fill the membrane in an orderly uniform monolayer as needed to understand how increasing treatment uniformity can impact electroporation results. One thing that is apparent from this study is that all of the cells do not interact with the micropores in a uniform manner. It is apparent that some cells have longer portions that protrude into the membrane pores deeper than other cells.

Figure 4.28. Diagram of cell interaction with micropores

153

Figure 4.29. 3D images of 3T3 cells interacting with the PCL micropore array

In the transfection study pGFP was used to indicate a successful electroporation event. Cells were loaded onto the PCL membranes using a suction pressure to bring cells in contact with the membrane pores. Figure 4.30 shows positive pGFP expression as indicated by green fluorescence at 24 and 48 hours after electroporation. Cells

154 expressing pGFP have received the gene and are alive and using it. It is apparent that the cells are lined up in the array and the image reveals that over 80% of the cells on the pores have been successfully transfected after 48 hours.

A B

300 µ푚 300 µ푚

Figure 4.30. pGFP expression 48 (A) and 24 (B) hours after electroporation

4.9 Membrane sandwich electroporation (MSE)

Here we continued our study of practical implementation of methods to make the electric field more uniform. In MSE we used commercially available membranes to surround cells and carry out electroporation. In MSE there was some inherent non- uniformity since membranes had microscale pores but in a random order although it is not certain how this would impact our results. Our goals here were to confine genetic material around the cells using the membranes and carry out electroporation using the focusing effects of the micropores, we also aimed to demonstrate a system that does

155 not require as in depth of membrane fabrication techniques compared to the micropore array.

4.10 MSE materials and methods

4.10.1 MSE experimental setup

The membrane sandwich electroporation (MSE) experimental system is shown in

Figure 4.31(a). We used a high precision computer numerically controlled (CNC) machine (AeroTech Inc, Pittsburgh, PA) to fabricate the MSE device and platform. The

MSE platform is connected with a custom electronic square wave pulse generator (Table

4.1).

156

Figure 4.31. Experimental system (a) and fluidic platform (b) for membrane sandwich electroporation (MSE)

157

Table 4.1. Square wave generator technical specifications

The fluidic device shown in Figure 4.31 (b) contains a pair of cross channels (500

μm in width and depth) is connected by a center hole. One cross channel is on the top of the device and the other runs along the bottom. A 1 cm diameter reservoir located at the top center of the device is where membranes are fixed.

We fabricated the MSE device from an Acrylite® acrylic plastic sheet (thickness:

1/16", US Plastic Corporation, Lima, OH) using the CNC machine (AeroTech Inc,

Pittsburgh, PA). A 50-μm thick polymethylmethacrylate (PMMA) film (Fisher

ScientificInc., USA) was heat welded on the backside of the device using a thermal film laminator(Catena 35, GBC, Addison, IL), enclosing the bottom channel but allowing top access via reservoirs at the ends.

Before heat welding the film to the backside the devices is cleaned in an acetone ultra-sonication bath for 10 min and successively rinsed with isopropyl alcohol (IPA) and deionized water (DI water) to remove contaminants and debris. The average MSE device is reused more than 20 times before being discarded.

158

4.10.2 Reporter genes

Reporter Plasmid gWizTM green fluorescence protein vector (GFP, 5757 bp) was used and described in Section 4.7.3 In addition to pGFP we used a secreted alkaline phosphatase vector (pSEAP, 6569 bp). pSEAP was purchased from Aldevron(Fargo, ND), and purified with an EndoFree Plasmid Maxi Kit from Qiagen (Valencia,CA, USA) according to the manufacturer’s instructions. pSEAP encodes for a gene that enhances a cells ability to secrete alkaline phosphatase. This secreted molecule can be detected and quantified in the culture media through optical methods. Although it does not indicate which cells exactly received the gene, the levels of secreted phosphatase can be compared between experiments using similar numbers of cells to indicate the relative success of a transfection protocol.

4.10.3 MSE procedures

We used NIH 3T3 fibroblasts in our bulk electroporation study with the Bio-Rad

Gene Pulser XcellTM power supply(Catalog No. 165). The Bio-Rad system was equipped with CE module and ShockPod. In each experiment we pipette a 100 μL suspension cells

(1 × 105 cells) and 5μg DNA sample into a 2-mm gap electroporation cuvette. The bulk electroporation process parameters were chosen based on Tekle’s work (Tekle, 1991),

(Table 4.2). Post-electroporation, each 10 μL of the cell suspension is transferred to an individual well in a 24-well plate and covered with 240 μL culture media. The

159 transfection efficiency and cell viability were measured at 24 or 48 hours after electroporation.

Table 4.2. Comparison of bulk, localized, and MSE electrical parameters

For localized cell electroporation and MSE, an underneath support membrane with an average pore size of 400 nm was constructed from 3-mm square patch of track- etched polyethylene terephthalate (PET) membrane (BD Biosciences, San Jose, CA). The support membrane is fixed at the center reservoir of the fluidic device using sealing tape

(shown in Figure 4.31 (a)). Once the support membrane was in place, a 10-μL drop of suspended cells (1104 cells) was loaded into the reservoir using a micropipette, and a vacuum of 34 3 kPa was applied to the underneath channel to trap the cells on the support membrane. Next, a 10 μm thick solid spacer membrane with 2.4-mm diameter hole is placed around the support membrane and a 3mm square patch of track-etched

PET membrane with an average pore size of 3 μm is added over the top of the immobilized cells. Opti-MEM reduced-serum medium is then pipetted into the channels

160 and the center reservoir. We connected two platinum wire electrodes (0.010” diameter) to fluid in the inlet and outlet reservoirs, and pipette 0.5 μg DNA into the cathode reservoir. Once loaded with cells and DNA a two step electroporation process is carried out. First a DNA attraction step is executed and then electroporation is done (Table 4.2).

The DNA molecules migrate from the cathode side to the anode side as shown in Figure

4.32 (b). After 15 to 20 minutes, the support membrane with the cells is transferred to a

24-well plate loaded with 250 μL culture media in each well. The transfection efficiency and cell viability were measured at 24 or 48 hours after electroporation.

Figure 4.32. Schematic of (a) MSE membrane disk and (b) DNA migration during electroporation

161

In spacer assisted MSE normal MSE support membranes were modified using a polymer stamping microfabrication technique. A PDMS stamp with 300µm diameter pillars that are 10 µm tall was used to stamp a polystyrene spacer layer on top of a 3- mm diameter track-etched polyethylene terephthalate (PET) membrane (BD

Biosciences, San Jose, CA). A 6% by weight polystyrene in anisol solvent was spin coated on to the PDMS stamp at a speed of 3000rpm for 60 seconds to produce the stamp with polymer “ink”. A hot plate was heated to 85 oC with a glass plate cover on top of its surface. The track-etched PET membrane was placed on the glass plate atop the hot plate. When the stamp was loaded and ready it is pressed by hand onto the heated track etched PET membrane for 5 seconds. Once pressed the PDMS stamp is peeled away leaving behind the polystyrene spacer bonded with the track etched PET Figure

4.34. Loaded “Stamp” Stamp contact

Figure 4.33. Schematic of the polymer stamping technique to produce spacer membranes for MSE

162

Figure 4.34. Polystyrene spacer bonded with a PET track etched membrane

The transfection efficiency of plasmid GFP (pGFP) was evaluated by the percentage of the cells with green fluorescence among the cells observed by phase contrast with the same visual area. Our inverted digital microscope (Eclipse TS100,

Nikon, USA) was equipped with X-Cite 120 fluorescence illumination system (EXFO Life

Sciences Division, Canada) capable of detecting GFP expression and cell viability after electroporation. For each visual field we first observed cells under phase contrast microscopy, then by fluorescence. B-2E/C fluorescence filter set (Excitation filter wavelengths: 465 – 495 nm, Dichromatic mirror cut-on wavelength: 505 nm, and barrier filter wavelengths: 515 – 555 nm; Nikon, USA) were used for green fluorescence detection. A digital camera (SPOT Insight 2MP Firewire Color Mosaic, Diagnostic

Instruments, Inc., Sterling Heights, MI) captured phase contrast and fluorescence images.

The transfection efficiency of plasmid SEAP (pSEAP) was expressed as the total

SEAP activity per ten thousand initial input cells. We conducted a colorimetric assay for each sample of culture media collected 48 hours after electroporation based on the

163 hydrolysis of p-Nitrophenyl phosphate (pNPP). pNPP substrate solution was prepared using SIGMAFAST™ pNPP tablets (Sigma-Aldrich, Catalog No. N1891, St. Louis, MO).

We added 100 μL of culture media and 25 μL of pNPP substrate solution to each well of a 96-well plate. The plate was dark incubated for 15 minutes at room temperature, and a 405 nm wavelength was used to read signals in the multi-well plate reader (GENios

Pro, Tecan, Durham, NC, USA). We plotted a standard curve of absorbance value at

405nm versus total SEAP activity (mU), and experimental readings at 405nm were normalized to total SEAP activity per ten thousand input cells.

To test cell viability NIH 3T3 cells were collected and mixed with equal volume of trypan blue stain (Invitrogen, Catalog No. 15250). 10 μL of the mixture was loaded onto a hemocytometer (a counting chamber covered with a cover slide), and counted. The trypan blue stained cells indicated by a dark blue coloration were dead cells allowing us to calculate the viability in as the percentage of the cells excluded from staining.

4.11 MSE results

4.11.1 MSE transfection

Figure 4.35 highlights transfection results using pGFP in MSE against bulk and local electroporation. Green fluorescence indicates a successfully transfected cell since it is alive and expressing the pGFP to produce a green fluorescent protein. Using local electroporation from either the top(Figure 4.35c) or bottom(Figure 4.35b) produces

164 better results than bulk electroporation(Figure 4.35a), while MSE (Figure 4.35c) produced the best results of all. Using another plasmid SEAP, the levels of transgene expression mediated by localized cell electroporation and MSE were quantified. The amount of secreted alkaline phosphatase (SEAP) expression mediated by MSE was improved about 40% over other localized cell electroporation Figure 4.36. When adding a spacer to the MSE system an additional benefit can be seen in pGFP fluorescence inside the wells of the spacer in Figure 4.37. A higher density of successfully transfected cells was present. Figure 4.38 also shows the previous ordered array pGFP transfection results compared with the MSE system and it also indicates that a slight improvement of transfection is seen in the ordered array system over standard MSE.

165

150µ푚

Figure 4.35. pGFP expression comparison using (a) bulk (b) local bottom (c) local top and (d) MSE

166

Figure 4.36. Comparison of SEAP gene activity between MSE and local electroporation

Figure 4.37.pGFP expression in spacer MSE vs standard MSE

167

250µm

125µm

Figure 4.38. pGFP expression in standard MSE vs ordered array local electroporation

4.12 Array and MSE device discussion and conclusions

This work demonstrates some effective techniques to fabricate membrane based electroporation microdevices. Both cleanroom and non-cleanroom approaches can be used successfully to create micropore systems capable of enhancing electroporation beyond standard bulk electroporation protocols. Since debris around the edge of the micropore can contribute to a reduction in process uniformity it is best to use the processing conditions that create micropores with little or no surface roughness.

Cleanroom processing is by far the superior fabrication technique and it is capable of producing very uniform pores with little surface debris. In addition to surface roughness disadvantages, the laser ablation technique is also a slower process for creating micropore arrays over a large surface area (1mm2).

A new approach to electroporation was developed. MSE has shown that it is capable of delivering genes pGFP and pSEAP to NIH-3T3 cells with better transfection

168 efficiency than bulk electroporation. It was able to increase cell viability and transfection results through the use of microporous membranes that focus the electric field and confine genetic material around the cells. This technique provides a new alternative gene delivery tool to be used on cells relevant to gene therapy including cancer and stem cells.

Bothe MSE and a single membrane micropore array are very capable of high efficiency transfection using simple animal cell models such as NIH-3T3 and common plasmids such as pGFP. The transfection results are the best indication that these processes provide better outcomes compared with bulk electroporation. From a process standpoint the use of a single membrane system is much simpler than the multiple steps are needed to carry out MSE, however from a device fabrication standpoint the MSE system makes use of membrane materials that are already mass produced and commercially available.

It is clear from results using both systems that process uniformity drives higher transfection efficiency. Using just using the random micro pores in the MSE process we get improvements in transfection over bulk electroporation. Additionally we can make slight improvements to make the MSE process more uniform by the addition of the microfabricated spacer resulting in gains in transfection efficiency. The first generation spacer system used in MSE was only present on the edges and the membranes spanned a 2.4mm diameter area. It was apparent that some variation in spacing could be taking place across the two membranes. The addition of the microfabricated spacer membrane

169 ensured that a 10 µm gap was maintained across the membrane and that nearly each cell experienced the same electroporation process conditions.

170

Chapter 5 : Conclusions

5.1 Conclusions

We have developed a new method to study electroporation using optical tweezers that enables us to observe a cell in suspension throughout the electroporation process. We used this system to provide a statistically meaningful understanding of the relationship between individual cell size and the critical electric field for cell poration.

Although it has been assumed that a larger cell requires a smaller critical electric field by some researchers, our results using three cell lines, K562, NIH-3T3, and mESCs, across cell radius sizes ranging from 3-15 µm indicate that there is no relationship between cell size and the critical electric field. Instead, the critical transmembrane potential is cell size dependent. This method and results are an important step toward fully understanding the mechanisms and sub-processes behind electroporation.

Previous work in support of the theory that the critical eclectic field was cell size dependent used different cell types instead of cells within the same population. They used the average cell size in the population of cells instead of the actual size of each individual cell. Part of the reason behind this was a lack of tools to quantitatively study individual cells. In some studies they concluded that the critical electric field should

171 increase with a decreasing cell size while others concluded that it should decrease with increasing cell size. One approach to study individual cells size in relationship to the critical electric field is to carry out a bulk electroporation event and then measure the size ranges of cells that were or were not electroporated. The problem with this technique, as we have shown in our work here, is that cell-cell interactions in bulk electroporation can cause variations in the critical electric field between cells within a process and a molecular delivery shielding effect can cause some cells to remain undetected although they may have been electroporated. For these reasons we believe that our optical tweezers method provides the best framework for individual cell experiments in electroporation.

Possibly one of the largest problems in fundamental research in this field is the lack of validation between theoretical models, experimental models and experimental reality. The complex interactions between the molecules to be delivered, the cells, the surrounding environment, and the electric field present a daunting array of variables to consider when approaching a research project in this field. In conducting fundamental research in electroporation one must consider theoretical models since in many cases the information is not able to be validated experimentally. In the case of our critical electric field experiments we were able to bring together new equipment to systematically and quantitatively test the validity of the Schwan equation in a way that was previously impossible. It is only a matter of time until we find the right combination of tools to understand the complex mechanisms involved.

172

In addition to the cell size study using the optical tweezers we also were able to show how interactions between cells can cause non-uniform electric fields in bulk electroporation. We showed that when cells come very close to each other, an enhancement or reduction in the transmembrane potential takes place depending on relative orientations to the electric field. Non-uniform electric fields created in bulk electroporation are a contributing factor to low cell viability and transfection efficiency and can be improved through process modifications.

It is likely impossible to remove all factors that cause non-uniform electric fields.

Clearly the simple interactions between how cells are positioned relative to each other can cause differences in the electroporation conditions experienced by different cells.

However, it is very possible to create a situation where process conditions applied to each cell are very close to being the same and any minor differences in treatment are not amplified to lethal levels. The use of high electric fields in bulk electroporation presents a situation where any differences (physiological or location) between the cells can either cause cell lysis or for it to not porate. By taking steps to reduce the overall electric field we eliminate the potential for lethal amplification of the electric field therefore increase cell viability. Since we have shown that there should be an average critical electric field for a given cell population we should be able to create a process environment where each cell experiences an electric field above the critical value but still below any lethal level.

173

We developed several microdevice based approaches toward making electroporation more uniform. These approaches were based around using micropores to locally focus and enhance the electric field strength on a portion of the cell membrane thereby reducing the overall electric field and increasing cell viability and transfection results. Our single micropore study made use of a microfluidic device that contained a single micropore membrane. We were able to demonstrate that the micropore reduced the critical electric field for cell poration and the critical electric field became smaller with smaller pore sizes. Additionally, we were able to show that the PI dye delivery occurred on a shorter time scale using the micropore compared to bulk electroporation because of the micropores ability to enhance diffusion processes.

We demonstrated several important microfabrication processes to prepare microporous membranes including micromilling, laser ablation, and soft lithography.

Soft lithography provided the best method to make the microporous membrane with a uniformly smooth surface. Soft lithography is also best implemented when making very large arrays since lithography can pattern large areas in a short amount of time. When only a single micropore is needed we demonstrated that the femtosecond laser ablation process provided the best combination of pore size uniformity and surface roughness.

Laser ablation is best suited to rapidly manufacture a single or small array of micropores.

Two microdevices including a micropore array and MSE demonstrated the advantages of making electroporation conditions uniform across each cell. MSE used

174

“off the shelf” microporous membranes with randomly spaced pores to focus the electric field and provided gene confinement around the cell and as a result we observed better pGFP and pSEAP transfection over bulk electroporation. As the MSE process is made more uniform with the addition of a microfabricated spacer, pGFP transfection results were better. The micropore array provided additional order in the electroporation process since the pore locations are regularly spaced and can provide a

1:1 cell to pore ratio and was also more successful than bulk electroporation. Results showed that both MSE and micropore arrays required a lower electric field and as a result had both transfection and cell viability advantages over bulk electroporation.

This work has made it apparent that increasing the process uniformity by exposing each cell to the same electric field and environmental conditions can enhance cell viability and transfection outcomes in electroporation. As electroporation outcomes become more efficient through the use of techniques presented here we move closer to a gene delivery method capable of meeting the demands of today’s gene therapy protocols and eventually toward the reduction of incurable genetic diseases.

5.2 Recommendations

5.2.1 Micropore optical tweezers

In our work we presented a novel method combining optical tweezers with electroporation to better understand some fundamental mechanisms behind electroporation. This is only the beginning for what we can learn from this approach.

175

We primarily studied bulk electroporation with our optical tweezers, however, this system is equally valuable to study micro/nanodevices such as our micropore systems.

The optical tweezers can be used to locate a cell relative to a micro/nanopore or micro/nanochannel and capture information about delivery mechanisms. To do this with the current system a microdevice would need to have a thin profile under 200 µm to be able to interface with the optical tweezers focal distance. A soft lithography or other photolithography based microfluidic device would provide the best opportunity to achieve the dimensions required for this study and the optical tweezers equipment.

5.2.2 Confocal microscopy with optical tweezers

Confocal microscopy was used in part of the studies here involving microdevices, however, it was not combined with our optical tweezers system. The confocal microscopy system has certain advantages over the plain optical microscopy system currently used with our optical tweezers including higher sensitivity detection, laser illumination, and 3-D image capture. The combination of an optical tweezers with a confocal microscope system has potential to provide an extremely powerful and unique tool to study biomedical microdevices. Our confocal microscopy system provides excellent conditions for detecting and observing stained nucleic acids that were simply unable to see using the optical tweezers system. This combination could provide an ideal system to study mechanisms behind gene delivery in electroporation and could

176 provide revealing footage of genetic material entering a cell since the tweezers can hold a cell in a specific location throughout a process.

5.2.3 Nanopore electroporation

As nanotechnology matures we are beginning to have access to more nanofabrication techniques. Our research labs have developed two different nanofabrication techniques that have the ability to be applied in electroporation. The first technique is a sacrificial template imprinting technique capable of producing a polymer membrane with conically shaped pores with the small end around 80 nm

(Figure 5.). The second technique is a combined lithography with DNA stretching technique to produce nanochannels on the order of 100 nm (Figure 5.2). Using these structures we could begin to investigate nanopores or nanochannel enhanced electroporation and to determine whether nanoscale features can yield additional or unique benefits beyond microscale features in cell electroporation.

177

80nm

2 m

Figure 5.1. nanoporous polymer membrane from sacrificial template imprinting (Wang S. , 2006)

nanochannel

Figure 5.2. Two microchannels connected by a nanochannel (P. Boukaney, et al., 2010)

178

5.2.4 Electrical impedance spectroscopy

Electrical impedance spectroscopy (EIS) measurements can provide electrical parameters regarding the cell membrane capacitance and resistance. EIS can also detect fluctuations in current through a cell membrane. By incorporating EIS with our electroporation systems we could have additional information that would help develop electrical models (e.g. Figure 5.3) in our system. Electrical models could be used to better understand the cell electroporation mechanism and to predict membrane breakdown while accounting for electrical losses in the channels and electroporation fluid and optimize process conditions in bulk and micro/nanodevice based electroporation. In particular by combining the EIS with our micropore electroporation system we can differentiate between cells that have been electroporated, lysed, or unaffected by the electroporation pulse. This could also provide a complimentary system to study the critical electric field without using optical methods such as dyes.

179

Figure 5.3. Electrical model of single cell electroporation (Bandiera 2007)

We conducted a preliminary EIS test using our single micropore experimental system. A potentiostat ( G300, Gamry ) was connected to electroporation electrode leads to measure impedance of a single cell in a micropore. This equipment has a maximum current output of 300 mA, with a current resolution of 1 fA, and a voltage resolution capability of 1 µV. It can measure up to 1013 ohms over a frequency range of

10 µHz to 300 kHz. This EIS data was used to detect electrical characteristics before and after electroporation. The EIS system was connected directly to the single micropore system at the electrodes via alligator style clips and interfaces with its own PC and software package to measure, control, and plot results.

180

The curves in Figure 5.4 were generated in a critical electric field experiment with a single 3T3 cell on a 4 µm diameter pore. The electric field upon the influx of PI dye and shift in impedance value was 186 V/cm and was within a single standard deviation of the average critical field indicated in PI dye studies (170V +/- 19V/c). As seen in Figure 5.4 there was an increase in impedance from the electroporation device with only media when we add the micropore membrane. A further increase in impedance from the device with micropore membrane happened when a cell is added over the micropore. A drop in impedance occurred after the cell was electroporated and the impedance value was between an unelectroporated cell and a device without a cell on the micropore. Although this data is interesting, the detailed mechanism of EIS and how it can be used as an analytical tool for cell electroporation need to be further investigated.

181

Figure 5.4. Impedance and phase dependence on frequency before and after electroporation

182

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Appendix A: Standard SEAP Activity Curve

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Appendix B: Software Interface for the Femtosecond Laser Motion Control

Figure B.1: Windows Software Interface for G-Code Programming

Developed by Hae Woon Choi, December 2007

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Appendix C: Derivation of Electroporation Equations

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