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Membrane Sandwich Electroporation for In Vitro Gene Delivery
Dissertation
Presented in Partial Fulfillment of the Requirements for the Degree Doctor of Philosophy in the Graduate School of The Ohio State University
By
Zhengzheng Fei, M.S.
Graduate Program in Chemical Engineering
The Ohio State University
2009
Dissertation Committee:
Professor L. James Lee. Yang, Advisor
Professor Robert J. Lee
Professor Jessica Winter
Copyright by
Zhengzheng Fei
2009
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ABSTRACT
Gene therapy is the delivery of therapeutic genes into cells and tissues with the
aim of treating and curing a disease. As an enhanced understanding of the roles of genes
in health and disease, gene therapy is showing promise against various diseases such as
cancer, diabetes, Parkinson's disease, and several inherited physiological defects. Viral
transduction is very efficient, but safety issues, such as immune and inflammatory
responses, have hampered their clinical uses in humans. Non-viral methods, including
either chemical transfection with cationic lipids/polymers or physical transfection using
electroporation/microinjection, are becoming attractive approaches.
Electroporation is one of the most popular non-viral gene transfer methods for in
vitro cell transfection. Initial studies with electroporation experienced very low
transfection efficiencies and cell viability, severely limiting the development of this
technology. The emergence of nucleofection (a modified electroporation technology)
provided an efficient means for transfecting cells in vitro. However, nucleofection still
encounters many limitations such as the large number of cells required (>106) and high
cost involved. Moreover, cell viability is still an issue due to the high electric voltage
used and the non-uniform electric field strength distribution generated during the process.
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To address these problems, we propose to develop an electroporation system
based on an innovative micro-/nanoengineering technology for in vitro gene delivery. In
our approach, electroporation is carried out in a mild and uniform electric field, with
potential for a wider process window that can be generated to cover a wide range of cell
lines and even primary cells.
A new membrane sandwich electroporation (MSE) approach was demonstrated
using plasmids GFP and SEAP as model materials. NIH 3T3 fibroblasts were tested and a
significant improvement in transgene expression was observed compared to current
electroporation techniques. In the MSE method, the focused electric field enhances cell
permeabilization at a low electric voltage, leading to high cell viability; more important,
the sandwich membrane configuration is able to provide better gene confinement near the
cell surface, facilitating gene delivery into the cells.
Next, we demonstrated the use of femtosecond laser fabricated micro-nozzle
arrays on a gelatin-coated PET membrane for MSE. Using micro-nozzle array enhanced
MSE, we observed high and uniform gene transfection, and good cell viability of mouse
embryonic stem (ES) cells compared to the bulk electroporation. Since typically cells or
tissues from the patients are very limited and therapeutic materials such as plasmids and
oligonucleotides are very expensive, the ability to treat a small number of cells (i.e. a
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hundred) offers great potential to work with hard-to-harvest patient cells for patient-
specific ex vivo gene therapy and in vitro pharmaceutical kinetic studies.
Numerical calculation of transmembrane potential qualitatively explains the
observed differences between MSE and bulk electroporation. Since there’s a good
correlation between transfection results and transmembrane potential calculations, the
simulation process with the threshold experiments can be used to predict the transfection
results, and thus largely reduced the trial-and-error window size.
Furthermore, we successfully integrated an electrospun nanofiber scaffold as a
cell-binding substrate into MSE, called nanofiber based MSE. With a micro-well spacer,
the uniform size of mouse ES cell colonies were obtained, and plasmid transfection by
electroporation were performed during colony formation. In addition, repeated plasmid
SEAP transfection of NIH 3T3 fibroblasts was tested and better cell survival and
recovery rate was observed using the electrospun nanofiber scaffold as compared to using
micro-porous membrane. Due to its capacity of extend the exposure time with
reprogramming factors, nanofiber based MSE demonstrated the potential for efficient
induced pluripotent stem (iPS) cell generation by repeated plasmid transfection.
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ACKNOWLEDGMENTS
I would like to express my sincere gratitude to my advisor Dr. L. James Lee for
his patient guidance, constructive advice and continuous support during my PhD study at
the Ohio State University. I am indebted to Dr. Yubing Xie, Dr. Shengnian Wang, Dr.
Xin Hu, Dr. Hae Woon Choi, Dr. Sadhana Sharma, Mr. Brian Henslee, Mr. Bo Yu, Dr.
Yun Wu, and Dr. Weixiong Wang for their technical support, insightful suggestions, and
encouragements. I would like to acknowledge Dr. Jingjiao Guan, Dr. Xulang Zhang, Dr.
Chee Guan Koh, Dr. Yong Yang, and all former and current group members for their
valuable discussions, and helpful comments. Thanks also go to Mr. Shi-Chiung Yu, Dr.
Chunghe Zhang, Mr. Daniel Gallego, Ms. Natalia Higuita, Mr. Yong Chae Lim, Mr. Chi
Yen, and all the students and staffs at the center for their warm help on my research
project.
The financial support and technical directions from NSF sponsored Nanoscale
Science and Engineering Center for Affordable Polymeric Biological Devices (NSEC-
CAPBD) is appreciated.
Finally, I would like to thank my parents for their love and dedications for raising,
supporting, and educating me. Great appreciations to my husband, Mr. Ziru Zhang, for
his love, support, accompany and understanding through all these years.
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VITA
June, 2001…………………………………B.S. Chemical and Biochemical Engineering,
Zhejiang University, Hangzhou, China
March, 2004……………………………… M.S. Chemical and Biochemical Engineering,
Zhejiang University, Hangzhou, China
October 2004 to present.…………………. Graduate Research Fellow,
Chemical and Biomolecular Engineering,
The Ohio State University, Columbus, OH
PUBLICATIONS
1. Fei, Zhengzheng; Wang, Shengnian; Xie, Yubing; Henslee, Brian E.; Koh, Chee
Guan; Lee, L. James. Gene transfection of mammalian cells using membrane
sandwich electroporation. Analytical Chemistry (2007), 79, 5719.
2. Guan, Yixing; Fei, Zhengzheng; Lou, Man; Yao, Shanjing. Choromatographic
refolding of recombinant human interferon gamma by an immobilized sht GroEL191-
345 column. Journal of Chouromatography A (2006), 1107, 192.
3. Guan, Yixing; Fei, Zhengzheng; Lou, Man; Yao, Shanjing. Production of
minichaperone (sht GroEL191-345) and its function in the refolding of recombinant
human interferon gamma. Protein & Peptide Letters (2005), 12, 85.
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4. Jin, Ting; Guan, Yixing; Fei, Zhengzheng; Yao, Shanjing. A combined refolding
technique for recombinant human interferon-gamma inclusion bodies by ion-
exchange chouromatography with a urea gradient. World Journal of Microbiology &
Biotechnology (2005), 21, 797.
5. Fei, Zhengzheng; Guan, Yixing; Yao, Shanjing. A colorimetric method to assay
biological activity of recombinant human IFN-γ. Weishengwuxue Tongbao (Chinese
Edition) (2004), 31, 65.
6. Guan, Yixing; Fei, Zhengzheng; Lou, Man; Yao, Shanjing. Minichaperone
(GroEL191-345)-mediated in vitro refolding of recombinant human interferon
gamma inclusion body. Shengwu Huaxue Yu Shengwu Wuli Jinzhan (Chinese
Edition) (2004), 31, 907.
7. Jin, Ting; Guan, Yixing; Fei, Zhengzheng; Lou, Man; Yao, Shanjing. Renaturation of
recombinant human interferon gamma inclusion body by dilution. Huagong Xuebao
(Chinese Edition) (2004), 55, 770.
FIELDS OF STUDY
Major Field: Chemical Engineering
Minor Field: Biochemical Engineering
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TABLE OF CONTENTS
Page
Abstract ............................................................................................................................... ii
Acknowledgments............................................................................................................... v
Table of contents.............................................................................................................. viii
List of Tables .................................................................................................................... xv
List of Figures .................................................................................................................. xvi
Chapter 1: Introduction................................................................................................. 1
1.1 Background ............................................................................................................. 1
1.2 Objectives ............................................................................................................... 3
1.2.1 Membrane sandwich electroporation (MSE) .................................................... 3
1.2.2 Micro-nozzle array enhanced MSE .................................................................. 3
1.2.3 Nanofiber based MSE....................................................................................... 4
Chapter 2: Literature review......................................................................................... 6
2.1 Gene delivery.......................................................................................................... 6
2.1.1 Viral versus non-viral ....................................................................................... 6
2.1.2 In vivo versus in vitro........................................................................................ 7
2.2 Electroporation........................................................................................................ 8
2.2.1 Electroporation theory ...................................................................................... 8
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2.2.1.1 Pore formation during electroporation.................................................... 9
2.2.1.2 Pore resealing after electroporation ...................................................... 11
2.2.1.3 Possible mechanisms of gene delivery by electroporation ................... 11
2.2.2 Factors to determine cell suspension electroporation ..................................... 12
2.2.2.1 Electric field strength with pulse duration and number ........................ 13
2.2.2.2 Cell density and properties.................................................................... 15
2.2.2.3 Concentration and properties of genetic materials................................ 16
2.2.2.4 Buffer composition ............................................................................... 17
2.2.2.5 Temperature .......................................................................................... 18
2.2.3 In vivo vs in vitro electroporation ................................................................... 19
2.2.4 Commercially available in vitro electroporation systems............................... 20
2.2.5 Microfluidic electroporation ........................................................................... 23
2.3 Genetically modified embryonic stem cells by electroporation ........................... 25
2.3.1 Embryonic stem cells and their properties...................................................... 25
2.3.2 Gene delivery to embryonic stem cells ........................................................... 26
2.3.3 Electroporation of embryonic stem cells ........................................................ 28
2.4 Generation of transgene-free induced pluripotent stem cells by electroporation . 29
2.4.1 Induced pluripotent stem cells ........................................................................ 29
2.4.2 Strategies to generate induced pluripotent stem cells ..................................... 31
2.4.3 Electroporation for transgene-free induced pluripotent stem cells ................. 32
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Chapter 3: Membrane Sandwich Electroporation....................................................... 34
3.1 Introduction........................................................................................................... 34
3.2 Materials and methods .......................................................................................... 35
3.2.1 DNA preparation............................................................................................. 35
3.2.2 NIH 3T3 fibroblast culture and preparation.................................................... 36
3.2.3 Experimental set-up ........................................................................................ 37
3.2.4 Fabrication and assembly of microfluidic device ........................................... 39
3.2.5 Electroporation procedure............................................................................... 40
3.2.5.1 Bulk electroporation.............................................................................. 40
3.2.5.2 Localized cell electroporation and MSE............................................... 41
3.2.6 Detection of green fluorescence protein (GFP) expression ............................ 43
3.2.7 Assay for secreted alkaline phosphatase (SEAP) Activity ............................. 44
3.2.8 Cell viability.................................................................................................... 45
3.2.9 DNA distribution study by spin-disk confocal microscopy............................ 45
3.3 Results and discussions......................................................................................... 47
3.3.1 Comparison of MSE with bulk electroporation .............................................. 47
3.3.2 Comparison of MSE with localized electroporation....................................... 47
3.3.3 Mechanism analysis by a spin-disk confocal microscope .............................. 50
3.4 Conclusion ............................................................................................................ 52
Chapter 4: Micro-nozzle array enhanced membrane sandwich electroporation ........ 55
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4.1 Introduction........................................................................................................... 55
4.2 Fabrication of micro-pore arrays on gelatin-treated polyethylene terephthalate
(PET) track-etched membrane by femtosecond laser ablation ...................................... 56
4.2.1 Micro-patterning of pores by femtosecond pulsed laser ablation................... 56
4.2.2 Femtosecond laser system used in this study.................................................. 58
4.2.3 Thermal effect of femtosecond laser fabrication on gelatin-coated
polyethylene terephthalate surface.............................................................................. 61
4.2.4 Femtosecond laser drilling of gelatin-treated polyethylene terephthalate track-
etched membrane with micro-pore arrays................................................................... 63
4.3 Micro-nozzle enhanced sandwich electroporation................................................ 66
4.3.1 Experimental ................................................................................................... 66
4.3.1.1 Reporter plasmids ................................................................................. 66
4.3.1.2 Culture of mouse embryonic stem cells................................................ 68
4.3.1.3 Experimental set-up .............................................................................. 69
4.3.1.4 Electroporation procedure..................................................................... 70
4.3.1.5 Assay for transfection efficiency and cell proliferation........................ 73
4.3.1.6 Statistical analysis................................................................................. 74
4.3.2 System optimization........................................................................................ 75
4.3.4.1 Converging micro-nozzle vs straight micro-channel............................ 75
4.3.4.2 Effect of porosity and micro-pore shape of top membrane .................. 76
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4.3.4.3 Effect of top membrane location........................................................... 80
4.3.3 Comparison of MSE with bulk electroporation and nucleofection ................ 83
4.4 Simulation of transmembrane potential distribution............................................. 86
4.4.1 Three-layer model........................................................................................... 87
4.4.2 Two-dimensional (2-D) simulation process.................................................... 89
4.4.3 Simulation results............................................................................................ 91
4.4.3.1 Effect of cell shape................................................................................ 91
4.4.3.2 Effect of porosity and pore shape of top membrane ............................. 93
4.4.3.3 Converging micro-nozzle vs straight micro-channel............................ 95
4.4.3.4 Comparison of micro-nozzle enhanced sandwich electroporation with
bulk electroporation .............................................................................................. 97
4.5 Conclusion ............................................................................................................ 97
Chapter 5: Nanofiber based membrane sandwich electroporation ............................. 99
5.1 Introduction........................................................................................................... 99
5.2 Materials and methods ........................................................................................ 101
5.2.1 Cell culture.................................................................................................... 101
5.2.2 Fabrication and characterization of nanofiber scaffolds with micro-well
spacers ....................................................................................................................... 101
5.2.2.1 Preparation of electrospun poly (ε-caprolactone) (PCL) /gelatin
nanofiber scaffolds.............................................................................................. 101
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5.2.2.2 Fabrication of PCL /gelatin nanofiber scaffolds with polystyrene (PS)
micro-well arrays ................................................................................................ 103
5.2.2.3 Structure characterization by scanning electron microscopy (SEM).. 104
5.2.3 Experimental set-up for nanofiber based MSE............................................. 105
5.2.4 Electric Resistance Measurements................................................................ 106
5.2.5 Electroporation procedure............................................................................. 107
5.2.5.1 Single cell electroporation .................................................................. 107
5.2.5.2 Cell colony electroporation................................................................. 108
5.2.6 Assays for transfection efficiency and cell proliferation .............................. 109
5.2.7 Cell morphology characterization by confocal microscopy ......................... 110
5.3 Optimization of nanofiber based membrane sandwich electroporation ............. 110
5.3.1 Effect of support membrane.......................................................................... 110
5.3.2 Effect of nanofiber thickness ........................................................................ 114
5.4 Nanofiber based MSE of mouse embryonic stem (ES) cell colony.................... 116
5.4.1 Mouse ES cell colony formation with controlled size .................................. 116
5.4.2 Nanofiber based MSE with vs without micro-well spacer ........................... 118
5.4.3 Nanofiber based MSE vs Bulk electroporation of cell colony ..................... 121
5.5 Nanofiber based MSE of NIH 3T3 fibroblasts ................................................... 122
5.5.1 NIH 3T3 fibroblasts with micro-well spacer ................................................ 122
5.5.2 Repeated SEAP transfection of NIH 3T3 fibroblasts ................................... 123
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5.6 Conclusions......................................................................................................... 126
Chapter 6: Conclusions and recommendations......................................................... 127
6.1 Conclusions......................................................................................................... 127
6.2 Recommendations............................................................................................... 129
6.2.1 Individual cell array trapping........................................................................ 129
6.2.2 Cell membrane permeability experiments .................................................... 130
6.3 Possible ways of in vivo applications.................................................................. 135
References....................................................................................................................... 137
Appendix A: Standard Curve.......................................................................................... 148
Appendix B: Optimization of bulk electroporation and nucleofection of mouse embryonic
stem (ES) cells ................................................................................................................ 149
Appendix C: Analytical solution of transmembrane potential for a two-dimentional (2-D)
cell in bulk....................................................................................................................... 155
Appendix D: G-code generation for fabricating micro-pore arrays by femtosecond laser
......................................................................................................................................... 159
Appendix E: Electroporation of mouse embryoid bodies............................................... 160
Appendix F: Membrane permeability experiment.......................................................... 162
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LIST OF TABLES
Page
Table 2.1: Commercially available electroporation systems. .......................................... 21
Table 3.1: Technical specifications of the square wave pulse generator. ........................ 39
Table 3.2: Comparison of conventional bulk electroporation (BE), localized cell electroporation (LCE), and membrane sandwich electroporation (MSE). ....................... 41
Table 3.3: Optical set-up of spin-disk confocal system. (Hemminger et al., 2007) ........ 46
Table 4.1: Technical specifications of the multi-functional pulse generator. .................. 70
Table 4.2: Comparison of nucleofection, conventional bulk electroporation by Bio-Rad Gene Pulser XCell system, and membrane sandwich electroporation (MSE).................. 71
Table 4.3: Top membranes with different pore size, pore density, and pore shape......... 77
Table 4.4: Top membranes with different distance to cell binding membrane................ 81
Table 4.5: Parameters of the three-layer model (Kotnik et al., 1997) ............................. 90
Table 5.1: Properties of three different types of membranes used as support membrane.......................................................................................................................................... 113
Table 5.2: Thickness and corresponding resistance of nanofiber layer controlled by electrospinning time........................................................................................................ 115
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LIST OF FIGURES
Page
Figure 2.1: Theoretical drawing and cryo scanning electronen microscope (cryo-SEM; cryo =cold) images (the upper right insert) of cell membrane before electroporation (top) and pore formation by electric breakdown of the lipid bilayer after electroporation (bottom). (http://www.inovio.com)................................................................................... 10
Figure 2.2: Schematic of cell cycle. Outer ring: I = Interphase, M = Mitosis; inner ring: M = Mitosis, G1 = Gap 1, G2 = Gap 2, S = Synthesis; not in ring: G0 = Gap 0/Resting. (http://en.wikipedia.org/wiki/Cell_cycle) ......................................................................... 16
Figure 2.3: (a) Amaxa’s Nucleofection Biosystem. (www.amaxa.com); (b) Bio-Rad Gene Pulser XCell system. (www.biorad.com) ................................................................ 22
Figure 2.4: Microfluid electroporation devices: (a) single cell electroporation (Khine et
al., 2005); (b) electroporation microchip (Lin et al., 2001).............................................. 24
Figure 2.5: Three pathways to possibly reprogram multi-potent stem cells for treatment of human disorders. (Fuchs and Segre, 2000)................................................................... 27
Figure 2.6: Schematic of iPS-cell-based treatment. (Passier et al., 2008)....................... 30
Figure 2.7: Three strategies to generate induced pluripotent stem cells: (a) retroviral or lentiviral transduction, (b) adenoviral transduction, and (c) plasmid transfection. (Lowry and Plath, 2008) ................................................................................................................ 32
Figure 3.1: Plasmid maps of gWizTM green flurescence protein vector (GFP, 5757 bp) and secreted alkaline phosphatase vector (SEAP, 6569 bp). ............................................ 35
Figure 3.2: Experimental set-up (a) and fluidic device (b) of membrane sandwich electroporation (MSE). (Designed and fabricated by Mr. Shi-Chiung Yu, Dr. Weixiong Wang, and Dr. Chuhe Zhang, 2006) ................................................................................. 38
Figure 3.3: Schematic drawing of (a) cell-binding substrate in MSE disk and (b) DNA migration path during electroporation. ............................................................................. 43
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Figure 3.4: Comparison of membrane sandwich electroporation (MSE) with conventional bulk electroporation and localized cell electroporation (LCE) using plasmid GFP. The green fluorescence indicated green fluorescence protein (GFP) expression 24 hours after bulk electroporation (a), MSE (b), and LCE with genes and cells on (c) opposite sides and (d) the same side of the support membrane. ....................................... 48
Figure 3.5: Comparison of membrane sandwich electroporation (MSE) with localized cell electroporation (LCE) using plasmids SEAP. The bars indicated the activity levels of secreted alkaline phosphatase (SEAP) expressed by NIH 3T3 cells 48 hours after electroporation. Data were plotted with the standard deviation from the mean (n=3). .... 49
Figure 3.6: (a) DNA distribution in the gap between two membranes in the observed domain. The zero position is set at the surface of the top membrane. 3 sets of consecutive images were analyzed at each z slice. (b, c) Confocal images of the slices near the top membrane (x = 0) and in the middle of the two membranes (x = 5.2µm)........................ 53
Figure 4.1: Physical phenomena that are present when machining with a long laser pulse (a) and ultrafast laser pulses (b). (http://www.cmxr.com/Industrial/Handbook.htm)....... 57
Figure 4.2: Femtosecond laser CPA system (Model 2161, Clark-MXR) with micro-station. The arrow indicates the laser pathway. ................................................................ 58
Figure 4.3: Schematic drawing of regenerative amplifier, including High Reflective (HR) mirror, Faraday Rotator (FR), Pockels cell (PC), Dichouroic mirror (DM), and Radiofrequency (RF) unit.(Clark-MXR, CPA 2110 User manual. 2nd Edition, 2004) ... 59
Figure 4.4: Block diagram of the beam delivery system set-up. ..................................... 60
Figure 4.5: Heat effect of various laser beam power on the surrounding gelatin-coated polyethylene terephthalate (PET) surface. ........................................................................ 63
Figure 4.6: SEM image of PET track-etched membrane with average pore size of 400 nm after coating with gelatin. White arrows point out the pores blocked with gelatin. ... 64
Figure 4.7: (a) shape and size of the micro-pores produced under various laser beam power up to 4 mW; (b) SEM images of micro-pores on the gelatin coating side produced at the average laser beam power of 2.5 (upper) and 3.5 mW (lower)............................... 65
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Figure 4.8: Plasmid map of pmaxGFP, encoding the new green fluorescent protein from Pontellina sp. (http://www.lonzabio.com/uploads/tx_mwaxmarketingmaterial/ amaxa_Newsletter_amaxa-news-03.pdf).......................................................................... 67
Figure 4.9: The second generation of MSE system, including a multi-functional pulse generator, and a platform, which is able to handle three fluidic devices in parallel. (Designed and fabricated by Mr. Mr. Shi-Chiung Yu and Dr. Shengnian Wang, 2008) . 69
Figure 4.10: (a) Schematic of MSE disk set-up; (b) Schematic of DNA migration path in the MSE device. DNA molecules migrate from cathode to anode. .................................. 73
Figure 4.11: Effect of different pore shapes, micro-channel () and micro-nozzle (), on mouse ES cell transfection by MSE. The bars indicate total activity of SEAP expression 24 hours after MSE under the optimized electrical field (Appendix B). ......................... 76
Figure 4.12: Comparison of top membrane with different micro-pore size and micro-pore density in MSE: (a) transfection efficiency and (b) cell viability of mouse ES cells 24 hours after MSE. ............................................................................................................... 79
Figure 4.13: Effect of top membrane location in MSE on cell transfection: (a) Transfection efficiency and (b) cell viability of mouse ES cells 24 hours after electroporation. ................................................................................................................. 82
Figure 4.14: Comparison of mouse ES cell transfection by micro-nozzle array enhanced MSE, bulk electroporation by Bio-Rad Gene Pulser, and nucleofection. (a) Transfection efficiency and (b) cell viability 24 hours after electroporation. From left to right, bulk
electroporation with initial input cell number of 6101× and 5101× ; micro-nozzle
enhanced MSE with initial input cell number of 4101× ; and nucleofection with initial
input cell number of 6101× . ............................................................................................. 84
Figure 4.15: GFP transfection of mouse ES cell by micro-nozzle enhanced MSE. A
hundred of cells were trapped on a 1010 × micro-nozzle array, and (a) phase contrast and (b) fluorescent images were taken 24 hours after electroporation. ................................... 86
Figure 4.16: Simulation comparison of top membranes with different pore size, pore density, and pore shape. (a) Schematic diagram of Cases I to IV, from left to right, with electric field lines across/around a single cell; (b) calculated transmembrane potential distribution. θ is the angle around cell surface. ................................................................ 92
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Figure 4.17: Simulation comparison of transmembrane potential distribution of top membranes with different pore size, pore density, and pore shape. ................................. 94
Figure 4.18: Simulation comparison between support membranes with micro-nozzles and micro-channels (a) electric potential distribution and electric field lines across/around a single cell near a micro-nozzle (left) and micro-channel (right); (b) calculated transmembrane potential distribution. θ is the angle around cell surface. ........................ 96
Figure 5.1: Schematic diagram of fiber formation by electrospining process where a droplet of a polymer solution is elongated by a high electrical field. (http://nano.mtu.edu/Electrospinning_start.html)........................................................... 102
Figure 5.2: Schematic of fabricating electrospun nanofiber scaffold with polystyrene (PS) micro-well arrays. (1) PDMS stamp with micro-pillar arrays; (2) Drop-cast PS solution; (3) PS solution is spin-coated and it de-wets on the surface of the PDMS stamp; (4) PS in-between the features is removed; (5) PS micro-well arrays were bonded to electrospun nanofiber scaffold by thermal bonding. (Gallego et al., In preparation)..... 104
Figure 5.3: SEM image of PCL/gelatin nanofiber scaffolds with 300 µm PS micro-wells.......................................................................................................................................... 105
Figure 5.4: Schematic drawing of electrospun nanofiber based MSE........................... 106
Figure 5.5: Comparison of different cell-binding substrates used for membrane sandwich electroporation. The transfection efficiency (a) and cell viability (b) of mouse embryonic stem cells, and the resistance (R) of MSE disk (c) were presented using PET membrane only, aluminum oxide membrane only, PET membrane with nanofibers, and aluminum oxide membrane with nanofibers.................................................................................... 111
Figure 5.6: Effect of electrospun nanofiber thickness on the transfection efficiency (a) and cell viability (b) of mouse embryonic stem cells. The thickness of nanofiber layer corresponds to electrospinning time as shown in Table 5.2........................................... 115
Figure 5.7: Confocal images of mouse ES cell colonies after cultured 24 (a, c) and 48 (b, d) hours on randomly distributed PCL/gelatin nanofiber scaffolds without (a, b) and with (c, d) 100 µm PS micro-wells. The cell seeding density was 5,000 / mm2. Cells were fixed with 70% ethanol and stained with PI dye. The length of the standard bars is 100 µm. .................................................................................................................................. 117
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Figure 5.8: SEAP transfection of mouse embryonic stem cells by nanofiber based MSE without and with micro-well spacer, bulk electroporation by Bio-Rad Gene Pulser X-Cell system, and nucleofection: (a) transfection efficiency (b) cell viability 24 hours after electroporation. ............................................................................................................... 119
Figure 5.9: Confocal images of mouse ES cells 6 (a, c) and 30 (b, d) hours after nanofiber based MSE without (a, b) and with (c, d) 100 µm micro-well spacer. Mouse ES cells were fixed with 70% ethanol and stained with PI dye. The length of the standard bars is 100 µm................................................................................................................. 120
Figure 5.10: Morphology of NIH 3T3 fibroblasts with (a) and without (b) 300 µm micro-well spacer after 48 hours. .............................................................................................. 123
Figure 5.11: Repeated SEAP transfection of NIH 3T3 fibroblasts using PET micro-porous membrane () and electrospun PCL/gelatin nanofiber scaffolds (): (a) transfection efficiency (b) cell viability at day 1 and 2 post-electroporation. Both cell-binding substrates had the micro-well spacer. ................................................................ 125
Figure 6.1: Schematic illustration of individual cell array trapping by an optical tweezer array created by focused laser beam through a micro-lens array.................................... 130
Figure 6.2: Electroporation of a cell. The electroporation mediated gene transfection process includes two parts: (1) cell membrane break-down and reseal, (2) genes bounding to the cell membrane during the electroporation, and entering cell plasma by endocytosis. (http://www.inovo.com).................................................................................................. 131
Figure 6.3: The third generation of MSE system, including (a) a MSE stage with two MSE disks and (b) an electroporation box, which is able to connect with one AC pulse generator and one DC power supply. Each MSE disk consists of a bottom and a top piece. (Designed and fabricated by Mr. Shi-Chiung Yu and Dr. Weixiong Wang, 2009) ....... 133
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CHAPTER 1: INTRODUCTION
1.1 Background
Gene therapy is the delivery of therapeutic genes into cells and tissues with the
aim of treating and curing a disease. (Mulligan, 1993) As an enhanced understanding of
the roles of genes in health and disease, gene therapy is showing promise against various
diseases such as cancer, diabetes, Parkinson's disease, and several inherited physiological
defects. (Izumikawa et al., 2005; Le Meur et al., 2007) For instance, gene therapy has
already successfully restored the health of two children with severe combined
immunodeficiency in 1990. (Blaese et al., 1995) Since then, many clinical trials in human
are paving their ways towards treating human diseases.
Over the past decades, the use of genetically modified primary embryonic stem
(ES) cells is becoming an attractive tool for fundamental studies as well as clinical
applications. (Ben-Nuna and Benvenisty, 2006; O’Connor and Crystal1, 2006;
Strulovici1 et al., 2007) ES cells are pluripotent cells derived from the inner cell mass of
an in vitro fertilized embryo grown to the early stage, know as a blastocyst. ES cells are
good candidates for cell-based therapies due to their unlimited self-renewal capacity and
differentiation potential into various cell types that can function as neurons, muscles,
bone, or blood.
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Genetic manipulation of cultured ES cells provides great potential for ex vivo
gene therapy. (Schindhelm and Nordon, 1999) However, little progress has been made
due to the difficulties involved in successful transfection. Viral transduction of ES cells is
very efficient, but safety issues, such as immune and inflammatory responses, have
hampered their clinical uses in humans. (Niidome and Huang, 2002; Thomas et al., 2003)
Non-viral methods, including either chemical transfection with cationic lipids/polymers
or physical transfection using electroporation/microinjection, are becoming attractive
approaches. (Mehier-Humbert and Guy, 2005; Wells, 2004)
Electroporation is one of the most popular non-viral gene transfer methods for ES
cell transfection. (Tompers and Labosky, 2004) Initial studies with electroporation of ES
cells experienced very low transfection efficiencies (<20%) and cell viability (<50%),
(Mohour et al., 2006) severely limiting the development of this technology. The
emergence of nucleofection (a modified electroporation technology) provided an efficient
means for transfecting ES cells in vitro. (Lorenz and Harnack, 2004; Siemen et al., 2005)
However, nucleofection of ES cells still encounters many limitations such as the large
number of cells required (>106) and high cost involved. Moreover, cell viability is still an
issue due to the high electric voltage used and the non-uniform electric field strength
distribution generated during the process.
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1.2 Objectives
To address these problems, we propose to develop an electroporation system
based on an innovative micro-/nanoengineering technology for in vitro gene delivery. In
our approach, electroporation is carried out in a mild and uniform electric field, with
potential for a wider process window that can be generated to cover a wide range of cell
lines and even primary cells.
1.2.1 Membrane sandwich electroporation (MSE)
A novel electro-transfection method, called membrane sandwich electroporation
(MSE) was developed. NIH 3T3 fibroblasts were used as cell models and were tested
using reporter genes (plasmid pGFP and pSEAP). In the MSE set-up, the focused electric
field enhances cell permeabilization at a low electric voltage, leading to high cell
viability; at the same time, the sandwich membrane configuration is able to provide better
gene confinement near the cell surface, facilitating gene delivery into the cells. Compared
to current bulk electroporation techniques, the MSE method increased transfection
efficiency and cell viability.
1.2.2 Micro-nozzle array enhanced MSE
The MSE design could not provide a uniform electric field distribution to each
cell because of randomly distributed pores on the track-etched membrane. To address this
-4-
limitation, we recently fabricated polyethylene terephthalate (PET) membranes with well-
defined micro-hole arrays by using femtosecond pulsed laser ablation. By adjusting the
laser output powers and laser beam focus points, we were able to produce both
converging micro-nozzle and straight micro-channel arrays on the membrane. This new
design was used for gene transfection of mouse embryonic stem (ES) cells. The observed
good transfection results are explained by numerical calculations of the transmembrane
potential distribution on the cell surface.
1.2.3 Nanofiber based MSE
We further integrated the MSE method with a three-dimensional (3-D)
electrospun nanofiber cell culture system for genetic modification of ES cells in a gentle
manner without breaking the colonies. Electrospun fibers are known to provide an in
vivo-like environment required for healthy and viable cells. A combination of MSE and
fibers allows for the cells to be cultured on the same substrate before and after
transfection, and reduces the number of harsh instances, such as repeated cell
trypsinization for getting single cell suspension, and longer time duration outside the
incubator, which cells have to undergo during conventional electroporation. Furthermore,
this method provides the flexibility of using appropriate type of support membrane
substrates specific for an intended application and cell type. The applicability of this
method was demonstrated using mouse ES cells and NIH 3T3 fibroblasts as a model
system.
-5-
It should be stated that the results and the descriptions of membrane sandwich
electroporation presented in Chapter 3 have already been included in a published journal
paper: Fei et al., 2007. The experimental procedure and many results presented in the
Chapter 4 and 5 are being included in two manuscripts under preparation.
-6-
CHAPTER 2: LITERATURE REVIEW
2.1 Gene delivery
A major challenge in gene therapy is to deliver therapeutic genes into the
designated target cells with a high transfection efficiency and minimal cell damage.
There are two basic criteria used to distinct the field of gene delivery: viral versus non-
viral and in vivo versus in vitro.
2.1.1 Viral versus non-viral
Viral gene delivery uses genetically modified viruses to introduce genes into cells
or tissues by infection. These recombinant viruses are highly efficient, and have been
used in clinical trials since 1990. (Blaese et al., 1995) However, safety concerns, such as
immune and inflammatory responses, are limiting their clinical applications, (Thomas et
al., 2003) especially after the first gene therapy death caused by the adenovirus-based
treatment reported in 1999. (Hollon, 2002) Another challenge relates to virus-based gene
delivery is the high cost of viral vector production. In addition, there is always the fear
that viral vectors, once inside the patient, may recover their ability to cause disease.
-7-
Non-viral systems have become widespread for gene delivery because of their
relative safety and low manufacturing cost. Non-viral methods can be divided into two
groups: chemical and physical (or mechanical). (Niidome and Huang, 2002) Chemical
methods use reagents such as cationic lipids and polymers, or proteins that will complex
with DNA or RNA, condensing it into particles and directing it to the cells. Physical
methods involve introducing plasmids into cells by mechanical ways, including
microinjection, particle bombardment (gene gun), electroporation, sonoporation, and
laser irradiation. As compared to chemical methods, physical methods can transfer naked
DNA into cells directly and avoid the harmful side effects associated with synthetic
vectors, such as lipoplexes and polyplexes. (Mehier-Humbert and Guy, 2005)
2.1.2 In vivo versus in vitro
In vivo gene delivery refers to introducing the genes directly into the affected
tissue inside a living organism, and requires that the vector be targeted specifically and at
sufficiently high frequencies to the desired cell types. Viruses and liposomes have been
widely investigated as in vivo carriers, but safety issues such as immune response and
cytotoxicity have limited their clinical applications. (Niidome and Huang, 2002; Thomas
et al., 2003) Physical methods are more benign, because they can directly transfer naked
DNA into cells and avoid the risks associated with introducing a secondary agent.
(Mehier-Humbert and Guy, 2005; Wells, 2004)
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In vitro gene delivery involves performing the experiments in a controlled
environment outside a living organism. This type of research aims at describing the
effects of an experimental variable on a subset of an organism's constituent parts.
Currently, electroporation is one of the most popular research tools used for gene transfer
into mammalian cells in vitro.
2.2 Electroporation
2.2.1 Electroporation theory
Electroporation (or electropermeabilization) is a process that can induce transient
openings in cell plasma membrane by executing external electric field on cells, and thus
increased the permeability of cell membrane. (Chernomordik et al., 1987; Chang et al.,
1992; Weaver and Chizmadzhev, 1996; Gabriel and Teissie, 1997 and 1999) Since it was
developed in the early 1980’s (Neumann et al., 1982), electroporaiton has been widely
used to deliver exogenous macromolecules into cytoplasm, ranging from ions, drugs,
dyes, tracers, and antibodies, to DNA, RNA and oligonucleotides. (Coulberson et al.,
2003)
Normally, one or multiple short and high-voltage pulses are imposed on cells, and
cell membranes are permeable at the locations of highest transmembrane potential
gradient, typically the areas closest to the electrodes (the anode side opens first, followed
-9-
by the cathode side). (Gehl, 2003) Since the applied pulses are short, the lipid bilayer
structure can be restored and, therefore, the cell survives.
2.2.1.1 Pore formation during electroporation
The experimental observations of electroporating planar lipid bilayers
(Zimmermann and Vienken, 1982; Chernomordik et al., 1983; Glaser et al., 1988) and
the molecular dynamic (MD) simulations (Mounir, 2005) demonstrated that the kinetics
of pore formation by electric breakdown of the lipid bilayer (as shown in Figure 2.1)
includes three steps:
(1) Induction: Water fingers penetrate the hydrophobic core of the lipid bilayer
from both sides at the beginning of applying the electric field, and extend to form the
water wires.
(2) Stabilization: Hydrophobic polar heads migrate toward the interior of the
bilayer surrounding with hydrophilic polar heads, and thus stable large water pores are
formed.
(3) Resealing: water channels disappear and polar lipid head-groups go back to
the lipid-water interface after the electric field is turned off.
Chang and Reese (1990) observed pore-like crater structures or volcano funnels
of 20 to 120 nm diameters in electroporated red blood cell membrane by rapid-freezing
electron microscopy. The analytical methods indicated that the initial electropores should
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be much smaller. (Neumann et al., 1999) The observation of Chang and Reese (1990)
most probably results from the enlargement of smaller primary pores by osmotic or
hydrostatic pressure due to Maxwell stress.
Figure 2.1: Theoretical drawing and cryo scanning electronen microscope (cryo-SEM;
cryo =cold) images (the upper right insert) of cell membrane before electroporation (top)
and pore formation by electric breakdown of the lipid bilayer after electroporation
(bottom). (http://www.inovio.com)
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Besides leading to electric breakdown of the lipid bilayer, the transmembrane
potential gradient can also cause the opening of many protein channels in cell membrane.
(Tsong, 1991) Therefore, electric field induced pore formation occurs in both lipid
domain and protein channels.
2.2.1.2 Pore resealing after electroporation
The kinetics of pore resealing after electroporation is mainly measured by
studying the time course of the decrease in membrane conductivity (Chen and Lee, 1994;
Chernomordik et al., 1987), membrane permeability to small inorganic ions (Bier et al.,
2002), and the fraction of cells permeable to certain membrane-impermeant compounds
(Gabriel and Teissie, 1995; Rols and Teissie, 1989; Zimmermann et al., 1980). The
resealing process after cell electroporation was also analyzed theoretically, and compared
with experimental data available in the literature. (Saulis, 1997; Bier et al., 2002) It is
generally agreed that the resealing of the membrane requires seconds to minutes, and it is
a random process.
2.2.1.3 Possible mechanisms of gene delivery by electroporation
Two possible mechanisms for gene delivery by electroporation have been
proposed and verified by both in vitro and in vivo studies (Golzio et al., 1998; Mir et al.,
1999): diffusion-controlled and electrophoresis-driven uptake, both generally occurring
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near the temporary openings facing the cathode. Diffusion-controlled uptake involves
DNA molecules binding to the cell surface and diffusing into the cytoplasm leading to the
cell transfection. (Sukharev et al., 1992) Electrophoresis-driven uptake follows two steps:
(1) Induced by an applied electric field, cell membrane breaks down and results in
formation of lipid vesicles containing DNA; and (2) DNA molecules are loaded into cells
by endocytosis after electroporation. (Klenchin et al., 1991; Sukharev et al., 1992;
Glogauer et al., 1993)
2.2.2 Factors to determine cell suspension electroporation
The degree of transfection by electroporation is highly cell-dependent, and
normally assessed by two aspects: the transfection rate of therapeutic materials
(transfection efficiency) and the survival rate of electropermeabilized cells (cell
viability). (Chu et al., 1987)
The transfection efficiency of electroporation depends on various factors,
including the status of cells (e.g. growth phase, density, size, orientation) (Tsong, 1991),
the physical and chemical properties of therapeutic materials (e.g. DNA size,
configuration and concentration) (Tsong and Xie, 1997), and applied electric conditions
(e.g. pulse amplitude, duration and number) (Kotnik et al., 2003; Gehl, 2003). Other
issues that should be considered when performing electroporation include temperature,
post-pulse manipulation and composition of electrode and pulsing medium.
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Generally, cell survival is not a concern when working with bacterial cells, but
can be a major issue with mammalian cells. The viability of mammalian cells depends on
the resealing of electric-field-induced pores in cell membrane after electroporation and
the extent of excess leakage of intracellular molecules during electroporation. (Rols and
Teissie, 1990)
2.2.2.1 Electric field strength with pulse duration and number
For a certain cell line, extent of cell permeabilization is dependent on the
amplitude of electric pulses, and degree of molecule transportation is dependent on the
duration and number of electric pulses. (Gehl, 2003)
The transmembrane potential difference can be described by the following
equation (Teissie and Rols, 1993):
)]exp(1[cos τθ textm rfgEV −−=∆
(2.1)
where:
∆Vm: transmembrane potential difference, V/cm;
f : shape factor of impacted cells under an external field;
g: relative electric permeability;
Eext: external electric field strength, V/cm;
r: radius of the cell, µm;
θ : angle between Eext and the point on the cell membrane;
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t: elapsed time after the field is turned on, s;
τ : charging or relaxation time of membrane, s.
Typically, for a pure dielectric pulse, g =1. Transient and reversible breakdown of
the membrane can be achieved only when the transmembrane potential surpasses the cell
membrane capacitance, expressed as a threshold potential, ∆Vs. Considering the same
lipid bilayer feature of cellular membrane for eukaryotic cells, ∆Vs is reported to be 200
mV. (Gehl, 2003) As shown in Eq. (2.1), transmembrane potential difference is
dependent on the external field and cell properties. (Kotnik et al., 1997) When ∆Vm >∆Vs,
it is believed that transient hydrophilic pores are formed.
It is generally agreed that the formation of electropores takes place on the order of
micro- to miliseconds, whereas resealing of the membrane requires seconds to minutes.
(Rols and Teissie, 1990) If extensive electroporation is applied, there will be a high
probability for slow recovery of cells and the loss of intracellular components, leading to
irreversible permeabilization and eventually cell death. In addition, cell lysis occurs as
the phenomena of the electroporation-induced apoptosis, if the electric field strength is
too high or the pulse duration is too long. Typical electric field strengths used in cell
suspension electroporation are 0.5 ~ 1.0 kV/cm for mammalian cells. (Rols and Teissie,
1990)
Pulse shape has also played different roles in cell electroporation. (Kotnik et al.,
2003) There are two popular types of pulse shapes: square wave (rectangular) or
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exponentially decay. (Chang et al., 1992) An exponentially decay pulse is generated by
discharging a capacitor, which is relatively simple but cannot provide precisely control of
pulse parameters. A square wave pulse is generated by a more sophisticated instrument to
accurately control both the intensity and the duration of each pulse.
2.2.2.2 Cell density and properties
One essential factor for achieving good transfection and survival rate of the cells
is cell density in the suspension during the electroporation. Usually if the cell density is
lower than half million per milliliter ( 5105× /mL), the survival rate of cells is extremely
low. However, if the cell density is too high, the transfection efficiency largely decreases
as the result of the shield effect among the cells and electrofusion between cells in
contact (Tsong, 1991).
The capacity to undergo and recover from electroporation is highly cell-
dependent. Two significant factors are the size and growth phase of the recipient cells.
Larger the cell diameter, lower the recovery rate of cell membrane after electroporation.
(Potter, 1993) Electroporation of mammalian cells in logarithmic growth phase is much
more efficient than treatment in early or stationary growth phase. (Anderson et al, 1991)
Additionally, plasmids can access the nucleus more easily during the mitotic phase (M
phase) of the cell cycle (Figure 2.2) than entering post-mitotic state (G0 phase) outside
of the cell cycle. (Golzio et al., 2002a)
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Figure 2.2: Schematic of cell cycle. Outer ring: I = Interphase, M = Mitosis; inner ring:
M = Mitosis, G1 = Gap 1, G2 = Gap 2, S = Synthesis; not in ring: G0 = Gap 0/Resting.
(http://en.wikipedia.org/wiki/Cell_cycle)
2.2.2.3 Concentration and properties of genetic materials
Tsong and Xie (1997) have reported that the amount of DNA bound to the cell
membrane during electroporation is proportional to the amount of DNA transferred into
the cells post-electroporation. It is essential to facilitate binding of DNA to the cell
surface during electroporation. The increase of DNA concentration can arise the
possibility of DNA bound to cell membrane, especially around the effective area facing
the cathode, and thus enhance the transfection efficiency.
-17-
The size (molecular weight) affects uptake pathway: Smaller genes, such as
oligonucleofide, siRNA, and microRNA, are more easily transferred through the cell
membrane by diffusion of surface bound genes and electroosmosis of genes in the bulk
solution, while large plasmid delivery into cytosol is dominated by the electrophoresis of
surface bound DNA. The integration and expression of the loaded DNA is strongly
dependent on the DNA configuration (supercoiled, circular-relaxed, or linearized). Linear
DNA can be more effectively integrated into the host genome, but relative unstable in the
cytoplasm compared to supercoiled and circular DNA. (Tsong and Xie, 1997)
2.2.2.4 Buffer composition
Another key factor to influence the cell transfection is the recipe of
electroporation buffer. In many cases, the presence of high ionic strength in
electroporation buffer causes electric arching, especially during the application of longer
high-voltage pulses, and may kill most of cells. Therefore buffer solutions having a low
ionic strength and thus low conductivity were used in order to avoid cell damage as a
result of high currents when using high conductivity buffers. (Rols and Teissie, 1990)
Divalent cations, such as calcium (Ca2+) and magnesium (Mg2+) ions, are frequently
added to the electroporation buffer. Free Ca2+ and Mg2+ stimulates the pore resealing,
leading to an increase in cell survival rate. (Klenchin et al., 1991) Mg2+ (up to 10mM)
facilitates the binding of DNA to the cell membrane, resulting in an elevated transfection
rate. (Tsong, 1991) In addition, it has been reported that low concentration of glucose or
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sucrose has certain effect on transfection efficiency and cell viability in some type of
cells. (Myers and Tisa, 2003)
Change of the microenvironment of cells is believed to be the cause of cell death.
Although the role of loss of intracellular components is unclear, electroporation can
abolish the osmotic balance of cells with the external environment, especially the
equilibrium of individual ions, like the equilibrium of Na-K-ATPase at the plasma
membrane. More importantly, the colloidal-osmotic effect is lethal as it causes the
swelling of cells. The excess large molecules inside cells lead to the osmotic imbalance
inside. Ions are transported towards the outside to attain ionic equilibrium and water
flow inside, resulting in swelling. If the resealing of the cell membrane is ineffective,
continuous swelling may lead to cell bursting and death. Several surfactants, such as
dextran, PEG and poloxamer 188, were used in the external medium to balance the
colloid osmotic pressure changed by DNA or macromolecule delivery. (Kanduser et al.,
2003)
2.2.2.5 Temperature
Besides reversible breakdown of the lipid domain induced during the
electroporation, many voltage-sensitive protein channels in the cell membrane may open
when they experience an applied external electric field. Local temperature increase as a
result of Joule heating (or ohmic / resistive heating) generated with electric current, and
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high temperature around cells may denature these protein channels. To avoid the Joule
heating effect, it is commonly suggested to pre-incubate the cell suspension on ice before
electroporation.
On the other hand, a better cell survival rate was achieved when culturing the
cells in 37°C incubator right after electroporation rather than leaving them at room
temperature. It is possible that an increase of post-pulse temperature facilitates the
membrane recovery, and thus improved the cell viability.
2.2.3 In vivo vs in vitro electroporation
In vivo electroporation has been attempted in the past two decades to deliver
therapeutic materials into targeted tissues and organs. (Jaroszeski et al., 1999) Generally
it can be divided into two major groups: delivery of anticancer drugs (such as bleomycin
and cisplatin) for cancer therapy, and delivery of DNA, RNA or DNA vaccines for gene
therapy and DNA vaccination. For cancer therapy, successful examples of electroporation
trials have been done on animal or human patients with basal cell carcinoma, melanoma,
and head and neck cancers. In gene therapy or DNA vaccination, any type of cell or
tissue can be a target, and successful targeting locations have been reported in skin, liver,
muscle, brain and tumors; with muscle and skin as the two most popular targeted tissues.
For in vivo electroporation, the electric field distribution is a key factor. Different shapes
of electrode probes have been tested, such as plate electrodes and needle electrodes.
-20-
Needle electrodes are preferred due to good electric contact as well as stability. However,
the electric field distribution must be given particular consideration. Because of the lack
of a fundamental understanding of the biophysics of electroporation, current in vivo
process must be performed by trial-and-error.
To understand the fundamentals of the delivery mechanism, in vitro
electroporation has been widely used by researchers as a routine method for transient cell
transfection by plasmids or short strands of interfering RNA (eg. siRNA and microRNA).
(Coulberson et al., 2003) Because of its simplicity and reproducibility, in vitro
electroporation has also been used for introducing exogenous macromolecules, such as
antibodies and enzymes, into cells, and thus has been identified as a rapid and affordable
screening method to assess the behavior of fusion proteins within cells before using more
costly in vivo transgenic animal models.
Experimental results indicate that cell death is more likely in vitro than in vivo.
The most likely reason is that there is a large loss of intracellular molecules due to
abundant extracellular space for in vitro studies, while this space is much more limited
for in vivo studies inside tissues. (Chang et al., 1992)
2.2.4 Commercially available in vitro electroporation systems
In vitro electroporation systems are commercially available and parallel plate
electrode probes are widely used. The in vitro electroporation systems available in the
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U.S. market are listed in the Table 2.1. The leading systems, Amaxa Nucleofection
Biosystem and Bio-Rad Gene Pulser XCell system, are shown in Figure 2.3.
Table 2.1: Commercially available electroporation systems.
Company Generators Cost Advantage Disadvantage
Nucleofector Single cuvette:
$10K Amaxa
96-well Shuttle $20K
High
Efficiency
Unknown electric
condition;
Expensive cell-
specific buffer
Gene Pulser XCell Single cuvette:
$6K Bio-Rad
Gene Pulser
MXCell
96-well Shuttle:
$ 10K
BTX ECM 830 Single cuvette or
25-/96-well: $6K
Low cost; Limited efficiency
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Figure 2.3: (a) Amaxa’s Nucleofection Biosystem. (www.amaxa.com); (b) Bio-Rad
Gene Pulser XCell system. (www.biorad.com)
Although commercial electroporation systems have been reasonably successful,
the variation in transfection efficiency is large. Furthermore, the outcome of an
electroporation protocol is cell-type specific and varies among cells in a given
population. The quick development of proper protocols for different cells (in vitro) or
tissues (in vivo) is difficult due to the lack of detailed understanding of the
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electroporation mechanism and many variations presenting in the bulk systems. It is
necessary to develop new electroporation methods that can replace the current
approaches.
2.2.5 Microfluidic electroporation
Microfluidic devices have been used for electroporation since 1999 (Huang and
Rubinsky). Microfluidic electroporation offers several advantages compared to bulk
electroporation: (1) various configurations of electrodes can be patterned and placed in
close proximity so that very low potential differences are sufficient for electroporation of
the cell membrane (can be as low as 1 V/cm); (Khine et al., 2005) (2) cell handling and
manipulation are much easier and uniform electroporation is possible; (3) less reagents
are needed for transfection; and (4) the process can be monitored at the single cell level
for intracellular content transport.
Current microfluidic electroporation devices focus mostly on cell lysis. (Lee and
Tai, 1999; Lu et al., 2005) In general, solid electrodes are patterned in close proximity
with different shapes (e.g., saw-tooth structure) to concentrate electric field strength, and
the electric current is focused at a small constriction zone. In recent years, more and more
designs were being used for drug and gene delivery to mammalian cells. (Huang and
Rubinsky, 2003; Lin et al., 2001; Khine et al., 2005) Figure 2.4 shows two microfluidic
electroporation designs for mammalian cell transfection.
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Figure 2.4: Microfluid electroporation devices: (a) single cell electroporation (Khine et
al., 2005); (b) electroporation microchip (Lin et al., 2001).
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2.3 Genetically modified embryonic stem cells by electroporation
2.3.1 Embryonic stem cells and their properties
Embryonic stem (ES) cells are derived from the inner cell mass of a blastocyst.
ES cells were first isolated from mouse embryos in 1981 by two independent groups.
(Evans and Kaufman, 1981; Martin, 1981) More than 15 years of research on mouse ES
cells led to the breakthrough of human ES cells. In 1998, Thomson et. al. successfully
isolated hES cells from human blastocysts and grew the cells in the laboratory.
ES cells have two important characteristics that distinguish them from other types
of stem cells. The first is their developmental potential called pluripotency, which means
that they are able to differentiate into any of the three primary germ layers and thus into
different types of mammalian tissues. Second, ES cells can proliferate for one year or
more in cell culture without differentiating while maintaining the karyotype, which
allows the production of unlimited numbers of undifferentiated cells. Because of the
capability of self-renewal and vast differentiation, ES cells are identified as a promising
cell source that could be used therapeutically to treat tissue injury as well as genetic
disorders.
In general, ES cells are maintained in the undifferentiated state on feed layers of
embryonic fibroblasts or on feeder-free substrate (gelatin for mouse ES cells and Matrigel
for human ES cells) in the presence of basic factors (leukemia inhibitory factor for mouse
ES cells, and fibroblast growth factor for human ES cells). (Carpenter et al., 2003; Xu et
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al., 2001) When placed in suspension cultures, ES cells spontaneously form three-
dimensional cell spheroid-like aggregates called embryoid bodies (EBs), and differentiate
into various cell types, e.g. neural cells muscle cells, and blood cells. (Drukker and
Benvenisty, 2003) Cultured ES cells can also be induced to differentiate in vitro to
specific cell types by adding growth factors to the media. This provides the possibility for
cell-based therapies to repair damaged or destroyed cells or tissues.
2.3.2 Gene delivery to embryonic stem cells
Over the past decade, the use of genetically modified ES cells has become an
attractive tool for fundamental studies as well as clinical applications. (Ben-Nuna and
Benvenisty, 2006; O’Connor and Crystal1, 2006; Strulovici1 et al., 2007) Recent studies
in cell culture systems indicate that gene delivery to ES cells has the potential to treat
tissue injury and cure genetic disorders. (Fuchs and Segre, 2000) For example,
introducing the gene Nurr1 into ES cells has made it possible to regulate the formation of
dopamine-producing nerve cells for the treatment of Parkinson’s disease. (Kim et al.,
2002; Lindval and Kokaia, 2006)
Figure 2.5 summarizes three pathways to possibly reprogram multi-potent stem
cells for treatment of human disorders. (Fuchs and Segre, 2000) Two of the pathways
involve controlling in vitro differentiation of primary ES cells, and transplanting
differentiated cells. Control of ES cells differentiation to specific cell lineages in vitro is
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very challenging due to the complexity of signals involved in determining the fate of
stem cells. (Odorico et al., 2001) Genetically modifying ES cells with certain
transcription factors, growth factors and/or other signal molecules can guide their
differentiation process to specific cell types. (Zeng et al., 2003) Efficient gene delivery
techniques are the key to achieving the full potential of ES cells.
Figure 2.5: Three pathways to possibly reprogram multi-potent stem cells for treatment
of human disorders. (Fuchs and Segre, 2000)
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2.3.3 Electroporation of embryonic stem cells
Electroporation is one of the most popular non-viral gene transfer methods for ES
cell transfection. (Tompers and Labosky, 2004) However, high cell concentration (at
least five million per milliliter) and a large number of cells (at least half million for a
cuvette) are often required to obtain acceptable gene transfection and cell viability,
because the entire cell membrane is affected by the applied high electric field and the
electric field distribution is non-uniform due to randomly suspended cells and genes
during the electroporation process. Amaxa nucleofection (now Lonza), the best
commercial electroporation-based technique, demonstrated good transfection efficiency:
more than 85% transfection efficiency in mouse ES cells (Lorenz and Harnack, 2004),
and over 66% in human ES cells (Hohenstein et al., 2008; Siemen et al., 2005). However,
nucleofection relies on an expensive electroporation buffer that varies from cell to cell.
Neither the recipes of the cell-dependent electroporation buffer nor the electric
parameters are disclosed by the manufacturer. As designed for individual cell suspension
electroporation, it is hard to obtain good transfection of cell colonies or EBs with limited
programs suggested by the manufacturer. Also, there are concerns that the unknown
addictives in the electroporation buffer might affect the behavior of transfection cells,
limiting its use for research only.
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2.4 Generation of transgene-free induced pluripotent stem cells by electroporation
2.4.1 Induced pluripotent stem cells
Induced pluripotent stem (iPS) cells, just as the name implies, are artificial
pluripotent stem cells derived from non-pluripotent cells transfected by specific stem
cell-associated genes. Takahashi and Yamanaka of Kyoto University in Japan first
announced successful iPS cell generation in June 2006. Through direct reprogramming of
mouse skin cells, they obtained iPS cells by introducing four transcription factors,
Oct3/4, Sox2, c-Myc, and Klf4. (Takahashi and Yamanaka, 2006) In 2007, Yamanaka’s
group and Thomson’s group successfully generated iPS cells from human somatic cells.
Their work was published in Cell and Science respectively almost at the same time.
(Takahashi et al., 2007; Yu et al., 2007)
As natural ES cells, iPS cells possess high nucleus-to-cytoplasma ratio and typical
compact colony morphology. Chromosome analysis and flow cytometry expression
analysis shows that iPS cells have normal karyotypes, and expresse ES cell-specific cell
surface marker and genes. More importantly, iPS cells can be differentiatioted into
derivatives of all three germ layers. (Takahashi et al., 2006 and 2007; Yu et al., 2007 and
2009)
iPS cell lines provide an alternative source of autologous tissue for transplantation
in regenerative therapy, especially patient-specific cell-based therapy, as shown in Figure
2.6 (Passier et al., 2008; Maherali and Hochedlinger, 2008; Lensch, 2009) Not only can
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they be used for studing disease models and testing new drugs against those diseases, iPS
cells may also contribute to the fast screening of inducing pluripontent cells to
differentiate into desired cell types. (Figure 2.6) Besides its promise in scientific field,
iPS cells also solved ethical issues associated with the use of fertilized embryos to obtain
ES cells and oocytes for somatic cell nuclear transfer.
Figure 2.6: Schematic of iPS-cell-based treatment. (Passier et al., 2008)
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2.4.2 Strategies to generate induced pluripotent stem cells
There are three strategies to generate iPS cells as shown in Figure 2.7. (Lowry
and Plath, 2008) Prior to 2008, the generation of iPS cells was based on the integration
of the reprogramming genes into the host-cell genome by retroviral transduction, which
may also turn on cancer-causing genes. (Takahashi et al., 2006 and 2007; Yu et al.,
2007) To avoid the danger of insertional mutagenesis or potential oncogenesis using
retroviruses, scientists started to pursue the vector-free and transgene-free reprogramming
methods. Adenoviral transduction without genome integration (Stadtfeld et al., 2008)
and repeated plasmid transfection (Okita et al., 2008) were reported to achieve
reprogramming of mouse iPS cells. However, the reprogramming efficiencies of vector-
free and transgene-free approaches were less than 0.01%, much lower than that of
retroviral transduction. (Lowry and Plath, 2008) Using non-integrating episomal vectors,
Yu et al. (2009) has reported the generation of human iPS cells completely free of vector
and transgene sequences. Although the programming efficiency was improved to ~ 0.1%
using episomal vectors, it is still only one tenth of the highest efficiency achieved by
retroviral transduction.
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Figure 2.7: Three strategies to generate induced pluripotent stem cells: (a) retroviral or
lentiviral transduction, (b) adenoviral transduction, and (c) plasmid transfection. (Lowry
and Plath, 2008)
2.4.3 Electroporation for transgene-free induced pluripotent stem cells
To address the challenge of low efficiency of vector-free and transgene-free
reprogramming, scientists developed an alternative strategy involving delivery of a single
vector with all the required reprogramming genes by electroporation, and using piggyBac
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transposon technology to integrate the vector into the host-genome. (Kaji et al., 2009;
Nagy et al. 2009; Stadtfeld and Hochedlinger, 2009; Yusa et al., 2009) Kaji et al. (2009)
constructed all four genes, Oct4, Sox2, c-Myc, and Klf4, into a single plasmid vector
with a 2A-linkage. By using the 2A-peptide sequence, the vector is able to undergo self-
removal from a peptide undergoing translation, and thus realized complete removal of
vectors and transgenes after reprogramming (Pera, 2009; Stadtfeld and Hochedlinger,
2009) Using piggyBac technology, the single vector can easily integrate into the host-
genome. (Nagy et al. 2009; Stadtfeld and Hochedlinger, 2009) Through transient
expression of the transposase enzyme, the integrated genes can also be removed from the
host genome in a in a highly efficient and seamless fashion. (Nagy et al. 2009; Yusa et
al., 2009)
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CHAPTER 3: MEMBRANE SANDWICH ELECTROPORATION
3.1 Introduction
Among the physical and mechanical methods, electroporation-enhanced delivery
of plasmid vectors is gaining acceptance for both in vitro and in vivo applications. The
electroporation process induces transient openings in the plasma membrane by executing
electric pulses on cells and driving genes or drugs into the cytoplasm. It is applicable to a
wide variety of animal cells and tissues, simple to perform, and easy to use. However,
conventional bulk electroporation requires the use of a high electric voltage, leading to
low cell viability and limited transfection efficiency.
In this chapter, we present a much less invasive and more efficient gene delivery
method, called membrane sandwich electroporation (MSE). We trapped cells on a track-
etched polyethylene terephthalate (PET) membrane. Cell immobilization on a porous
surface leads to localized cell electroporation, allowing the use of a low applied voltage
to achieve temporarily dielectric breakdown of the cell membrane. When we placed
another track-etched PET membrane on the top of the immobilized cells and sandwiched
the cells between the two membranes, we observed a significant improvement of gene
transfection with minimal cell damage.
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3.2 Materials and methods
3.2.1 DNA preparation
Reporter Plasmids, gWizTM green flurescence protein vector (GFP, 5757 bp) and
secreted alkaline phosphatase vector (SEAP, 6569 bp), were purchased from Aldevron
(Fargo, ND), and purified with an EndoFree Plasmid Maxi Kit from Qiagen (Valencia,
CA, USA) according to the manufacturer’s instructions. Figure 3.1 shows the maps of
gWiz GFP and SEAP.
(Continued)
Figure 3.1: Plasmid maps of gWizTM green flurescence protein vector (GFP, 5757 bp)
and secreted alkaline phosphatase vector (SEAP, 6569 bp).
-36-
Figure 3.1 continued
3.2.2 NIH 3T3 fibroblast culture and preparation
NIH 3T3 fibroblasts (mouse embryonic fibroblast cell line, CRL-1658) were
purchased from American Type Culture Collection (ATCC, Manassas, VA), and were
grown in the culture medium. The culture medium consists of Dulbecco’s modified
Eagle’s medium: Nutrient Mix F-12 (D-MEM/F-12, Catalog No.10565), 2 mM L-
glutamine (Catalog No. 25030), 1mM MEM sodium pyruvate (Catalog No. 11360), and
10% (v/v) newborn calf serum (NCS, heat-inactivated, Catalog No. 26010). NIH 3T3
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cells were maintained in 25 cm2 T-flasks at 37oC with 5% CO2 and passaged every 2~3
days using 0.05% trypsin with 1mM EDTA·4Na (Catalog No. 25300).
Before experimentation, NIH 3T3 cells were plated on a Ф35mm plastic petri
dish and allowed to grow till 70 ~ 90% confluence. Cells were harvested by
trypsinization, and pelleted via centrifuge. The pellet was washed with Dulbecco's
Phosphate Buffered Saline (D-PBS, Catalog No. 14190) without calcium or magnesium
at least twice, and resuspended in GIBCO Opti-MEM I reduced-serum medium (Catalog
No. 51985). Cells were counted with a hemocytometer, and cell suspensions were
adjusted to desired cell concentration for electroporation.
All the media and solutions for cell culture and transfection were purchased from
invitrogen (Carlsbad, CA) unless otherwise specified.
3.2.3 Experimental set-up
The experimental set-up of membrane sandwich electroporation (MSE) is shown
in Figure 3.2(a). The MSE device and platform was fabricated using a high precision
computer numerically controlled (CNC) machine (AeroTech Inc, Pittsburgh, PA). The
MSE platform is connected with a square wave pulse generator designed and built in our
lab. The technical specifications of the square wave pulse generator are given in Table
3.1.
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Figure 3.2: Experimental set-up (a) and fluidic device (b) of membrane sandwich
electroporation (MSE). (Designed and fabricated by Mr. Shi-Chiung Yu, Dr. Weixiong
Wang, and Dr. Chuhe Zhang, 2006)
-39-
Table 3.1: Technical specifications of the square wave pulse generator.
Voltage Pulse length Pulse interval No. Pulses
(maximum)
Current
(maximum)
10 ~ 240 V 1 ~ 999 ms 1 ~ 999 ms 6 180 mA
3.2.4 Fabrication and assembly of microfluidic device
The fluidic device consists of a pair of cross channels connected by a center hole
as shown in Figure 3.2(b). One channel is present on the top of the device while the
other is on the bottom. Both channels are 500 µm in width and depth. The channel on the
top of the device intersects with a 1 cm diameter reservoir located at the center of the
device where membranes can be fixed to the device.
The MSE device was fabricated in 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 Scientific
Inc., USA) was 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
through reservoirs at the ends.
In order to remove any contaminants on the surface, the MSE device was cleaned
in acetone ultra-sonication bath for 10 min and successively rinsed with isopropyl alcohol
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(IPA) and deionized water (DI water) before thermal bonding. Each MSE device was
reused more than 20 times.
3.2.5 Electroporation procedure
3.2.5.1 Bulk electroporation
Bulk electroporation of NIH 3T3 fibroblasts was carried out using Bio-Rad Gene
Pulser XcellTM unit (Catalog No. 165) with CE module and ShockPod. 100 µL of
suspended cells (1 × 105 cells) and 5µg DNA sample was loaded into the 2-mm gap
electroporation cuvet. The electroporation conditions were chosen according to Tekle et
al. (1991)’s work, and are given in Table 3.2. After electroporation, every 10 µL of the
cell suspension was transfer to a well of the 24-well plate with 240 µL culture media. The
transfection efficiency and cell viability were measured at 24 or 48 hours after
electroporation.
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Table 3.2: Comparison of conventional bulk electroporation (BE), localized cell
electroporation (LCE), and membrane sandwich electroporation (MSE).
LCE and MSE
Method BE
(Tekle et al., 1991) Electroporation DNA Attraction
Field Strength (V/cm) 1,600 35 3.5
Pulse Frequency (Hz) 40 1 100
Pulse duration (ms) 0.4 500 5
No. of pulses 1 5 300
Electroporator Bio-Rad Gene Pulser Home-made
Note: The pulse type is a biopolar square wave. Electric field strength is defined as the
voltage amplitude divided by the distance between two electrodes. Voltage amplitude is
the absolute value of peak value minus zero.
3.2.5.2 Localized cell electroporation and MSE
For localized cell electroporation and MSE, a 3-mm diameter track-etched
polyethylene terephthalate (PET) membrane (BD Biosciences, San Jose, CA) was used as
the support membrane with an average pore size of 400 nm, and fixed at the center
reservoir of the fluidic device by sealing tape (shown in Figure 3.1(a)). First, a 10-µL
drop of suspended cells ( 4101× cells) was loaded onto the support membrane, and a
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vacuum of 334 ± kPa was used to trap the cells on the support membrane. Next, another
3-mm diameter track-etched PET membrane with an average pore size of 3 µm was
added on top of the immobilized cells with a spacer of ~ 10 µm between the two
membranes. Opti-MEM I reduced-serum medium was then loaded into the channels and
the center reservoir. Two thin silver wire electrodes were placed in the inlet and outlet
reservoirs, and 0.5 µg DNA sample was loaded into the reservoir with the cathode.
Finally, a DNA attraction step was performed, followed by electroporation (The
conditions are given in Table 3.2). The DNA molecules were migrated from the cathode
side to the anode side as shown in Figure 3.3(b). After 15 to 20 minutes, the support
membrane with the cells was transferred to a 24-well plate with 250 µL culture media in
each well. The transfection efficiency and cell viability were measured at 24 or 48 hours
after electroporation.
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Figure 3.3: Schematic drawing of (a) cell-binding substrate in MSE disk and (b) DNA
migration path during electroporation.
3.2.6 Detection of green fluorescence protein (GFP) expression
The transfection efficiency of plasmid GFP (pGFP) was qualified by the
percentage of the cells with green fluorescence among the cells observed by phase
contrast with the same visual area. An inverted digital microscope (Eclipse TS100,
Nikon, USA) equipped with X-Cite 120 fluorescence illumination system (EXFO Life
Sciences Division, Canada) was used to detect GFP expression and cell viability 24 hours
after electroporation. For each visual field, cells were first observed by phase contrast,
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then by fluorescence. B-2E/C fluorescence filter set (Excitation filter wavelengths: 465 –
495 nm, Dichouromatic mirror cut-on wavelength: 505 nm, Barrier filter wavelengths:
515 – 555 nm; Nikon, USA) was used for green fluorescence detection. Both phase
contrast and fluorescence images were taken with a digital camera (SPOT Insight 2MP
Firewire Color Mosaic, Diagnostic Instruments, Inc., Sterling Heights, MI) set gain at 4,
and controlled by the SPOT Advanced software.
3.2.7 Assay for secreted alkaline phosphatase (SEAP) Activity
The transfection efficiency of plasmid SEAP (pSEAP) was expressed as the total
SEAP activity per ten thousand initial input cells. Samples of culture media were
collected 48 hours after electroporation and determined by a colorimetric assay based on
the hydrolysis of p-Nitrophenyl phosphate (pNPP). pNPP substrate solution was fresh
prepared using SIGMAFAST™ pNPP tablets (Sigma-Aldrich, Catalog No. N1891, St.
Louis, MO). 100 µL of culture media and 25 µL of pNPP substrate solution were added
into each well of a 96-well plate. The plate was incubated in the dark for approximately
15 minutes at room temperature, and read at the wavelength of 405 nm on a multi-well
plate reader (GENios Pro, Tecan, Durham, NC, USA). A standard curve of absorbance
value at 405nm versus total SEAP activity (mU) was generated (Appendix A), and then
the experimental readings at 405nm were normalized to total SEAP activity per ten
thousand initial input cells.
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3.2.8 Cell viability
NIH 3T3 cells were collected in a single-cell suspension after electroporation, 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 then counted under a microscope. The trypan blue stainable cells were the
dead cells, and thus the viability of the cells was calculated in accordance with the
percentage of the cells excluded from staining.
3.2.9 DNA distribution study by spin-disk confocal microscopy
To facilitate visualization, large λ-DNA molecules stained with the dye YOYO-1
were used in the confocal microscopic experiments instead of plasmids GFP and SEAP.
Lambda DNA (λ-DNA, N6-methyladenine-free, 48502 bp) was purchased New England
Biolabs (Ipswich, MA) and used as received without further purification. According to
the staining procedure given by Perkins et al. (1997), the λ-DNA solution was diluted to
the concentration of 0.03 µg/ml in the imaging buffer, and incubated with YOYO -1
(EM491/EX509, Molecular Probes, Eugene, OR) at a dye-base pair ratio of 1:4 for 1 ~ 2
hours. The imaging buffer consisted of YOYO-1 iodide and 20% 2-mercaptoethanol
(Sigma-Aldrich, Catalog No. M7154) in sterile Tris-EDTA (TE) buffer (Fluka, Catalog
No. 93302). YOYO-1 iodide is an intercalating dye that stains the DNA backbone and
makes it possible to visualize the DNA. 2-Mercaptoethanol is a strong reducing agent that
-46-
retards photo bleaching of the YOYO-1 fluorescence dye by scavenging oxygen from the
solution. (Xiao et al., 2007)
A spin-disk confocal microscope (VisiTech International, Alexandria, VA) with
Z-stacking was used to trace the location of DNA molecules in the gap of the MSE setup.
The Yokogawa spin-disk scanning unit (CSU-22) synchouronized with Hamamatsu EM
CCD camera was connected to an inverted microscope (Olympus IX-81, Tokyo, Japan).
A solid-state laser line was employed with the supplying 50 mW at 491 nm wavelength.
A Jena piezo-controller was mounted underneath the 60X oil objective to carry out a Z-
direction scans with submicron accuracy (the minimum distance is 100 nm). The entire
system was controlled using VoxCell Scan software from VisiTech.
Table 3.3: Optical set-up of spin-disk confocal system. (Hemminger et al., 2007)
Parameter Value
Pinhole diameter 50 µm
Magnification (M) 60
Numerical Aperture (NA) 1.42
Refractive index (n) 1.52
Excitation wavelength 491 nm
Emission wavelength 515 nm
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3.3 Results and discussions
3.3.1 Comparison of MSE with bulk electroporation
Using plasmid GFP and NIH 3T3 fibroblasts as reporter gene and model cells,
bulk electroporation and MSE were tested. A significant improvement was observed of
green fluorescence protein (GFP) expression by using the MSE method over the
conventional bulk electroporation method (Figure 3.4(a, b)). In the conventional bulk
electroporation, a layer of foam was observed due to cell lysis in the high intensity
electric field (1,600 V/cm). In comparison, a much low electric field with the amplitude
of 35 V/cm was applied in our MSE method, and more than 90% of cell viability was
achieved. Cell viability was quantified right after electroporaion with the trypan blue
method.
3.3.2 Comparison of MSE with localized electroporation
Using plasmid GFP and NIH 3T3 fibroblasts as reporter gene and model cells,
two different cases were tested for localized cell electroporation. Cells and genes were
placed on either opposite sides (Figure 3.4(c)) or the same side (Figure 3.4(d)) of the
support membrane. In both cases, only a slight improvement was observed of green
fluorescence protein (GFP) expression over the conventional bulk electroporation method
(Figure 3.4(a)). When the MSE method was used, most cells survived after the
treatment, and GFP expression (Figure 3.4(b)) was much higher than in localized cell
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electroporation (Figure 3.4(c, d)). 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 localized cell electroporation (Figure 3.5).
(Continued)
Figure 3.4: Comparison of membrane sandwich electroporation (MSE) with
conventional bulk electroporation and localized cell electroporation (LCE) using plasmid
GFP. The green fluorescence indicated green fluorescence protein (GFP) expression 24
hours after bulk electroporation (a), MSE (b), and LCE with genes and cells on (c)
opposite sides and (d) the same side of the support membrane.
-49-
Figure 3.4 continued
0
5
10
15
20
25
30
35
SE
AP
Activity (
mU
)
Cell & gene on opposite sides
Cell & gene on the same side
MSE
Figure 3.5: Comparison of membrane sandwich electroporation (MSE) with localized
cell electroporation (LCE) using plasmids SEAP. The bars indicated the activity levels of
secreted alkaline phosphatase (SEAP) expressed by NIH 3T3 cells 48 hours after
electroporation. Data were plotted with the standard deviation from the mean (n=3).
-50-
3.3.3 Mechanism analysis by a spin-disk confocal microscope
To explain why the MSE method promotes transgene delivery, Z-stacking was
carried out using a spin-disk confocal microscope with a 60X oil objective to trace the
local concentration of DNA molecules during the electroporation process. This system is
similar to the Confocal Laser Scanning Microscopy (CLSM) micro-PIV system (Park et
al., 2004), but equipped with a Yokogawa CSU-22 spin-disk unit and Hamamatsu EM
CCD camera. It is capable of scanning more than 120 full-frame images (1024×1024
pixels) per second, sufficient to directly measure the DNA distribution inside the
sandwich gap in the MSE setup.
To facilitate visualization, YOYO-1 conjugated large λ-DNA molecules were
used instead of plasmids GFP and SEAP. The DNA solution was loaded to the cathode
side, while the anode side was loaded with the buffer solution only. Scanning was carried
out every 0.4 µm across the 10-µm gap between two membranes. The time for DNA to
diffuse across the two planes was calculated according to Fick’s second laws
D
lt
2
2
≈ (3.1)
where
t: time for DNA to diffuse across the two planes,
D : diffusion coefficient of λ-DNA molecules,
4.0=l µm: diffusion distance.
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The diffusion of λ-DNA molecules is predicted according to the Zimm model in
good solvent conditions. (Smith et al., 1996; Hur and Shaqfeh, 2001)
36.0~ ≈−v
DNALD µm2/s (3.2)
where
6.0=ν : scaling exponent,
17=DNAL µm contour length for λ-DNA.
Thus the time for DNA to diffuse across the two planes is 2.0~ second, while the
scanning time interval between two adjacent planes is 04.0~ second, about one fifth of
the time for DNA to diffuse across the two planes. Therefore, DNA molecules observed
in adjacent planes must be different individual molecules.
Three sets of consecutive images were analyzed at each z slice and experimental
results were then plotted in Figure 3.6. A large number of DNA molecules were found in
the gap after electroporation, with a decreasing number of DNA molecules near the
membrane surface. Without the top membrane, DNA molecules were hardly seen this
time within the same distance of 10 µm from the bottom membrane. During
electroporation, the extent of cell permeabilization is dependent on the amplitude of
electric pulses, while the transport of the polyanionic DNA molecules into the cells is
driven by the electrophoretic force (Mir et al., 2005) and is dependent on the duration and
number of electric pulses (Gabriel and Teissie, 1997). The nano-scale pores in the
support membranes allow a focused electric field on the cell membrane, and thus enhance
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cell permeabilization at a low electric voltage. However, negatively charged DNA
molecules quickly migrate away from the negatively charged cell surface after the pulse
duration because of electric repulsion. This can seriously limit gene transfer into the cells.
When a negatively charged track-etched PET membrane is placed on top of the cells,
DNA molecules are prevented from diffusing away, as demonstrated in Figure 3.6. Thus,
the sandwich membrane configuration is able to provide better gene confinement near the
cell surface to facilitate genes transport into the cells.
3.4 Conclusion
A new membrane sandwich electroporation (MSE) approach was demonstrated
using plasmids GFP and SEAP as model materials. NIH 3T3 fibroblasts were tested and a
significant improvement in transgene expression was observed compared to current
electroporation techniques. In the MSE method, the focused electric field enhances cell
permeabilization at a low electric voltage, leading to high cell viability; more important,
the sandwich membrane configuration is able to provide better gene confinement near the
cell surface, facilitating gene delivery into the cells.
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Figure 3.6: (a) DNA distribution in the gap between two membranes in the observed
domain. The zero position is set at the surface of the top membrane. 3 sets of consecutive
images were analyzed at each z slice. (b, c) Confocal images of the slices near the top
membrane (x = 0) and in the middle of the two membranes (x = 5.2µm).
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Successful examples of in vitro electroporation trials have been done on animal
and human patients. Since typically cells or tissues from the patients are very limited and
therapeutic materials such as plasmids and oligonucleotides are very expensive, our MSE
method with the ability to deal with small number of cells with high transfection
efficiency and cell viability, offers a great impossibility for ex vivo gene therapy. The
applicability of the MSE method to primary cells and hard-to-transfect cells (such as
mouse embryonic stem cells and leukemia cells) is currently under investigation in our
laboratory.
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CHAPTER 4: MICRO-NOZZLE ARRAY ENHANCED MEMBRANE
SANDWICH ELECTROPORATION
4.1 Introduction
In Chapter 3, we demonstrated a membrane sandwich electroporation (MSE)
technique. However, the design could not provide a uniform electric field distribution to
each cell because of randomly distributed pores on the track-etched membrane.
Consequently, the gene transfection efficiency was limited.
To address this limitation, we formed well-defined micro-pore array on nano-
porous polyethylene terephthalate (PET) track-etched membranes using femtosecond
pulsed laser ablation. By adjusting the laser output powers and laser beam focus points,
we were able to produce both converging micro-nozzle and straight micro-channel arrays
on the membrane. This new design was tested by plasmid gWiz SEAP transfection of
mouse embryonic stem (ES) cells. The micro-nozzle array enhanced MSE method was
optimized. Effect of membrane porosity and pore shape was explored. The observed
transfection results are further explained by numerical calculations of the transmembrane
potential distribution on the cell surface.
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4.2 Fabrication of micro-pore arrays on gelatin-treated polyethylene
terephthalate (PET) track-etched membrane by femtosecond laser ablation
4.2.1 Micro-patterning of pores by femtosecond pulsed laser ablation
In the past several decades, the pulsed laser beam systems have been used in
micro-manufacturing a wide range of materials. Due to its flexibility and non-cleanroom
operation, laser micro-machining offers rapid and cost-effective evaluation of design
concepts during prototyping phases. Recent studies demonstrated that there is nearly no
thermal damage on the surrounding material if the laser-material interaction time is
picoseconds or shorter. (Aguliar et al., 2005; Varel et al., 1997) The major benefits of a
femtosecond (or ultrashort) laser pulse include its ability to produce very high peak intensity
( 1610≥ W/cm2) and rapid deposition of energy into the material. The difference between a
long laser pulse and ultrafast laser pulses on material removal and heat transfer is shown
in Figure 4.1. Therefore, femtosecond laser pulsing appears to be a very promising tool
for biomedical application (Serafetinides, 1997), such as surface modification of poly(ε-
caprolactone) (PCL) membrane for tissue engineering application (Tiaw et al., 2005), and
patterning of collagen for directed cell attachment (Liu et al., 2005).
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Figure 4.1: Physical phenomena that are present when machining with a long laser pulse
(a) and ultrafast laser pulses (b). (http://www.cmxr.com/Industrial/Handbook.htm)
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4.2.2 Femtosecond laser system used in this study
Figure 4.2 shows the femtosecond laser system used this study. An ultrashort
CPA laser system (Model 2161, Clark-MXR, Dexter, MI) was used as a femtosecond
laser energy source. It has a central wavelength of 775 nm, pulse width of 150
femtoseconds, and pulse repetition frequency of 3kHz. The system has a single-mode
erbium (Er) fiber oscillator to provide a seed pulse for chirped pulse amplification in a
Ti:Al2O3 –based regenerative amplifier.
Figure 4.2: Femtosecond laser CPA system (Model 2161, Clark-MXR) with micro-
station. The arrow indicates the laser pathway.
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The seed pulse from the frequency doubled SErF laser (λ=775nm) is stretched to
protect optics during pulse amplification. The stretched seed pulse is then transferred to a
regenerative amplifier to amplify the laser intensity. The optimum injection or cavity
dumping time is controlled by electronic devices (DT-505, Clark-MXR) and is critical for
the power and pulse duration. As shown in Figure 4.3, the injected seed laser is amplified
through an Ti:Al2O3 cavity which is pumped by a frequency doubled Nd:YAG laser
(λ=532nm) through a dichouroic mirror. After multiple passes of amplification, typically
5 times, the polarization of a Pockels cell is changed to eject the amplified beam to the
compressor.
Figure 4.3: Schematic drawing of regenerative amplifier, including High Reflective (HR)
mirror, Faraday Rotator (FR), Pockels cell (PC), Dichouroic mirror (DM), and
Radiofrequency (RF) unit. (Clark-MXR, CPA 2110 User manual. 2nd Edition, 2004)
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Figure 4.4 presents the block diagram of the optical beam delivery system. The
maximum output power of the system is 2.5 W. Since the typical range of power for
micromachining is less than 10 mW, it was attenuated to the mW scale by thin-film
polarizing beam splitters (PBS) and a λ/2 wave plate. The PBS regulates the transmission
and reflection rate based on the polarization direction, which can be adjusted by a λ/2
wave plate and incidence angle of laser beam.
Figure 4.4: Block diagram of the beam delivery system set-up.
(Farson et al., 2006 and 2008)
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The diameter of raw beam was measured to be 5 mm and the beam quality, M2,
was measured by using auto correlator to be 1.3 where a perfect Gaussian beam is M2=1.
The laser beam can be turned on or off using an external high speed mechanical shutter
with 4 ms shutter control time (LS055, NMlaser Inc), and was finally delivered to the
target material on a high precision X-Y stage (Parker-Hannifin) with 0.5 µm resolution
and 50 mm travel distance. The focusing optics was mounted on the Z axis with 0.5 µm
resolution and 25 mm travel distance. A 50x infinity corrected microscope objective lens
with numerical aperture (NA) of 0.42 (50x M Plan Apo NIR, Mitutotyo) was used for
fine focusing. Attenuated laser power was measured by a power meter (PM100, Thorlab)
placed right under the laser focusing lens. (Farson et al., 2006 and 2008)
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 this selected optics.
4.2.3 Thermal effect of femtosecond laser fabrication on gelatin-coated
polyethylene terephthalate surface
Femtosecond laser is an excellent tool for direct micro-patterning of biomaterials.
Because of the ultrashort contact between the laser beam and materials, there is very low
heat transfer to surrounding materials. (Farson et al., 2006 and 2008) Gelatin has been
widely used as a feeder-free substrate for maintaining mouse embryonic stem (ES) cells
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in an undifferentiated status. Different output laser beam powers up to 6 mW were tested
to determine the desirable power range for a minimal thermal effect on the surrounding
gelatin-coated polyethylene terephthalate (PET) surface.
A PET surface was coated with 0.1 % (w/v) gelatin (from porcine skin, Catalog
No. G2500, Sigma-Aldrich, St. Louis, MO) at room temperature for 30 minutes, yielding
a thin layer with thickness of 2 to 3 µm. Multi micro-wells were produced by scanning
the focused femtosecond pulsed laser beams under various powers over the gelatin-
coated PET surface. The G-code was programmed to fabricate 100 µm diameter micro-
wells with a center-to-center distance of 200 µm in a 22 × array. The G-code was
imported into a computer controlled motion system (MX80L, Parker Hannifin, Rohnert
Park, CA) with a 0.5-µm resolution in the X, Y, and Z axes. After Ultraviolet (UV)
sterilization, mouse ES cells were seeded at desired density to determine the heat effect
on the bioactivity of the surrounding gelatin-coated surface. Figure 4.5 shows the
morphology and distribution of mouse ES cells after 2-day culture. Mouse ES cells grew
very well around the wells at both 2 and 4 mW, while nearly no mouse ES cells were
observed on the surrounding surface of the wells milled at 6 mW. This implies that an
average output laser beam power up to 4 mW can provide proper irradiance incident to
fabricate the gelatin-coated PET surface without denaturing the gelatin coating.
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Figure 4.5: Heat effect of various laser beam power on the surrounding gelatin-coated
polyethylene terephthalate (PET) surface.
4.2.4 Femtosecond laser drilling of gelatin-treated polyethylene terephthalate
track-etched membrane with micro-pore arrays
Polyethylene terephthalate (PET) track-etched membranes (thickness: 10 µm,
pore size: 400 nm) were coated with 0.1 % (w/v) gelatin at room temperature for 30
minutes, yielding a thin layer with thickness of 2 to 3 µm. After gelatin coating, most of
the original submicron pores were blocked (Figure 4.6). A range of laser beam power
from 1 to 4 mW was then used to drill a periodic array of micro-pores in a gelatin-coated
PET membrane. A scanning electron microscope (SEM, Hitachi S-4300 Field Emission)
was used to characterize the dimension and surface roughness of the pores on the gelatin-
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coated PET membrane after femtosecond drilling. To perform SEM, a thin gold layer (50
nm) was sputter-coated on the Samples by Emitech k550x sputter coater.
Figure 4.6: SEM image of PET track-etched membrane with average pore size of 400
nm after coating with gelatin. White arrows point out the pores blocked with gelatin.
By varying the laser beam power pattern, the shape and the size of micro-pores
could be controlled. The relationship between pore dimension and laser beam power is
plotted in Figure 4.7(a). Smooth and well-defined micro-pores with minimal gelatin
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denaturing was obtained by femtosecond laser drilling at relatively low pulse energies in
our work, but a pulse energy below 2 mW was not high enough to penetrate the gelatin-
coated PET membrane.
(Continued)
Figure 4.7: (a) shape and size of the micro-pores produced under various laser beam
power up to 4 mW; (b) SEM images of micro-pores on the gelatin coating side produced
at the average laser beam power of 2.5 (upper) and 3.5 mW (lower).
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Figure 4.7 continued
Two different micro-pore shapes were generated: a converging nozzle at the laser
output power between 2 and 3.5 mW, and a straight channel at the output laser power
higher than 3.5 mW. Figure 4.7(b) shows SEM images of micro-pores on the gelatin
coating side produced under an average output power of 2.5 and 3.5 mW respectively on
gelatin-coated PET membrane. For 2.5 mW, the average pore size was 3.5 µm on the
PET membrane side and 1.5 µm on the gelatin coating side, while the average pore size
on both sides was 3.5 µm under 3.5 mW.
4.3 Micro-nozzle enhanced sandwich electroporation
4.3.1 Experimental
4.3.1.1 Reporter plasmids
Reporter plasmids pmaxGFP (3486 bp), encoding the new green fluorescet
protein from Pontellina sp., were purchased from Amaxa (now Lonza, Switzerland), and
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gWiz SEAP (6569 kbp) from Aldevron (Fargo, ND). Figure 4.8 shows the map of
pmaxGFP, and the map of gWiz SEAP was shown in Figure 3.1(b).
Figure 4.8: Plasmid map of pmaxGFP, encoding the new green fluorescent protein from
Pontellina sp. (http://www.lonzabio.com/uploads/tx_mwaxmarketingmaterial/
amaxa_Newsletter_amaxa-news-03.pdf)
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4.3.1.2 Culture of mouse embryonic stem cells
Mouse embryonic stem (ES) cells (CCE strain) were purchased from StemCell
Technologies (Vancouver, BC, Canada). All mES CCE cell culture reagents were
purchased from Invitrogen (Carlsbad, CA) unless otherwise specified.
Mouse ES cells were maintained in an undifferentiated state on 0.1% (w/v)
gelatin coated dishes in 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 at 37°C with 5% CO2, and passaged every 3 days
by trypsinization. Mouse ES cells were typically used when reaching 50~70% confluent.
For single cell suspension, 0.25% trypsin with EDTA·4Na (Catalog No. 25200) was used
for harvesting mouse ES cells; for cell colony suspension, 0.05% trypsin with EDTA ·
4Na (Catalog No. 25300) was used.
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4.3.1.3 Experimental set-up
The second generation of MSE system, including a multi-functional pulse
generator and a platform, is shown in Figure 4.9. The multi-functional pulse generator is
able to generate either square wave pulse or exponentially decay pulse. The technical
specifications of the multi-functional pulse generator are given in Table 4.1. The new
platform is able to handle three fluidic devices (Figure 3.2) in parallel.
Figure 4.9: The second generation of MSE system, including a multi-functional pulse
generator, and a platform, which is able to handle three fluidic devices in parallel.
(Designed and fabricated by Mr. Mr. Shi-Chiung Yu and Dr. Shengnian Wang, 2008)
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Table 4.1: Technical specifications of the multi-functional pulse generator.
Pulse Type Voltage Capacitance
No. Pulses
(maximum)
Current
(maximum)
Exponentially
decay
10 ~ 300 V 100 ~ 1000 µF 10 20mA
Voltage Pulse length Pulse interval
No. Pulses
(maximum)
Current
(maximum)
Square wave 10 ~ 300 V 1 ~ 999 ms 1 ~ 999 ms 10 20mA
4.3.1.4 Electroporation procedure
Nucleofection by Amaxa Biosystem:
Mouse ES cells were pelleted via centrifugation and washed twice with
Dulbecco's phosphate-buffered saline (D-PBS, pH 7.4, Invitrogen, Catalog No. 14190).
One million ( 6101× ) cells were then resuspended in transfection solution from Mouse ES
Cell Nucleofector® Kit (Amaxa, Catalog No. VPH-1001) with 200 ng/µL plasmids and
transferred to the 2-mm gap nucleofection cuvette. Mouse ES cells were nucleoporated at
Program A-13, A-23, A-24, and A-30 according to manufactory’s suggestion, and A-23
was selected in this study (Appendix A).
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Bulk electroporation by Bio-Rad Gene Pulser X-Cell system:
Mouse ES cells were pelleted via centrifugation and washed twice with D-PBS.
The pellet was resuspended in high glucose DMEM, and then transferred to the 1-mm or
2-mm electroporation cuvette (Bio-Rad). Two different cell concentrations were tested as
shown in Table 4.2. The electroporation parameters were optimized (Appendix B) and
the best electroporation condition is shown in Table 4.2.
Table 4.2: Comparison of nucleofection, conventional bulk electroporation by Bio-Rad
Gene Pulser XCell system, and membrane sandwich electroporation (MSE).
Method Nucleofection Bulk electroporation MSE
Pulse type Unknown Exponentially decay
Pulse number Unknown 1 1
Field Strength (V/cm) Unknown 500 150
Capacitance (µF) Unknown 500 500
DNA concentration (µg/mL) 200 200 50 5
Initial cell number 6101× 6101× 5101× 4101×
Cuvette 2-mm 2-mm 1-mm N/A
Total buffer volume (µL) 100 100 100 200
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Micro-pore array enhanced MSE:
A gelatin-coated PET membrane with micro-pore arrays drilled by femtosecond
pulsed laser was used as the cell binding membrane. Briefly, the cell binding membrane
was mounted at the center reservoir of the fluidic device (Figure 3.2). A drop of
suspended cells was loaded onto the support membrane, and a vacuum of 334 ± kPa was
used to trap the cells on the support membrane. Another PET track-etched membrane
with an average pore size of 1 µm was placed over the immobilized cells. The cells were
sandwiched between two membranes sealed together (Figure 4.10(a)). The bottom
channel and inlet reservoirs were loaded with high glucose DMEM with 5 ng/µL DNA,
and the top channel and outlet reservoirs were then loaded with high glucose DMEM. A
low DC voltage of 10 V was applied to the system for 5 seconds; DNA molecules were
migrated from the cathode to the MSE disk (Figure 4.10(b)) and concentrated in the
micro-pores of the cell binding membrane. Finally, the cells were pulsed with a single
exponentially decay pulse at 150 V/cm and 500 µF (Table 4.2), and the MSE disk was
transferred to a 48-well plate with 250 µL culture media in each well.
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Figure 4.10: (a) Schematic of MSE disk set-up; (b) Schematic of DNA migration path in
the MSE device. DNA molecules migrate from cathode to anode.
4.3.1.5 Assay for transfection efficiency and cell proliferation
The transfection efficiency of pmaxGFP in mouse ES cells was qualified by the
percentage of the cells with green fluorescence, and the transfection efficiency of pSEAP
was quantified by the activity level of secreted alkaline phosphatase by the transfected
cells, determined by a colorimetric assay based on the hydrolysis of p-nitrophenyl
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phosphate (pNPP). The experimental procedures were the same as described in Sections
3.2.6 and 3.2.7.
CellTiter 96® non-radioactive cell proliferation assay (MTS) was used to measure
the cell viability at 24-hour post-electroporation. MTS reagent was diluted in culture
media at 1:10, and the work solution was then added to the culture wells. The absorbance
at 570 nm (OD570) was recorded after 2 hours of incubation at 37°C with 5% CO2.
Background absorbance can be corrected by including negative control wells on each
plate to measure the absorbance from culture medium in the absence of cells. A set of
positive control wells containing untreated cells performed in sister cultures was settled
as 100% cell viability. The cell viability of each sample was calculated as
%100570570
570570×
−
−=
backgroundcell
backgroundsmaple
ODOD
ODODViabilityCell (4.1)
4.3.1.6 Statistical analysis
Data analyses were performed using Student’s t-test and are expressed as
arithmetic mean + s.d.; t-test values of *P < 0.05, **P < 0.01, ***P < 0.005 were
considered statistically significant.
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4.3.2 System optimization
4.3.4.1 Converging micro-nozzle vs straight micro-channel
A gelatin-coated PET membrane with micro-pore arrays drilled by femtosecond
pulsed laser was used as the cell binding membrane. As mentioned in Section 4.2.4, two
different micro-pore shapes were generated: a converging micro-nozzle and a straight
channel. To determine the effect of micro-pore shape on the MSE performance, mouse
ES cells were transfected by the MSE method with the converging micro-nozzle arrays
and the straight micro-channel arrays. The converging micro-nozzles were obtained at the
output laser power of 2.5 mW (Average pore size: 3.5 µm on the PET membrane side and
1.5 µm on the gelatin coating side), and the straight micro-channels was fabricated under
3.5 mW (Average pore size: 3.5 µm for both sides). Plasmids gWiz SEAP were used as
reporter genes, and transfection efficiency and cell viability of pSEAP transfection were
evaluated 24 hours after electroporation. As shown in Figure 4.11, SEAP expression
using the micro-nozzles almost doubled over that with micro-channels, mainly because
the electric field is more concentrated at the small-end of the micro-nozzle, resulting in
better localized electroporation. Additionally, pSEAP are relative large plasmids, and
they may experienced strong stretching along the axis direction in the converging
direction of the micro-nozzle, (Hu et al., 2009; Wang et al., 2008), leading to easier
delivery through the cell membrane.
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0
15
30
45
60
75
To
tal S
EA
P A
ctivity m
U/
10,0
00 I
nitia
l se
ed
ing
ce
lls
Micro-channel Micro-nozzle
Figure 4.11: Effect of different pore shapes, micro-channel () and micro-nozzle (), on
mouse ES cell transfection by MSE. The bars indicate total activity of SEAP expression
24 hours after MSE under the optimized electrical field (Appendix B).
4.3.4.2 Effect of porosity and micro-pore shape of top membrane
With initial ten thousands ( 4101× ) mouse ES cells trapped on a 100100 × micro-
nozzle array, five different commercially available membranes were investigated as the
top membrane in MSE to determine the effect of porosity on transfection efficiency and
cell viability. Table 4.3 shows the properties of these membranes (Cases 1 to 5).
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Table 4.3: Top membranes with different pore size, pore density, and pore shape
Case No.
Pore size
(µm)
Pore density
(pores /cm2)
Porosity
(%)
Pores per cell Pore shape
1 a 0.4 4 x 106 0.5 4 ~ 5
2 a 3 2 x 106 14.1 2 ~ 3
3 a 0.4 1.6 x 106 0.2 2 ~ 3
4 b 1 1.6 x 106 1.3 2 ~ 3
5 b 3 8 x 105 5.7 1 ~ 2
Straight
micro-channel
6 Small end: 1;
Large end: 3
1 x 106 1.9 1 ~ 2 Converging
micro-nozzle
a: Polyester (Corning, Lowell, MA);
b: PET (BD Biosciences, San Jose, CA).
pSEAP transfection experiments were carried out and the transfection results are
shown in Figure 4.12.Since the micro-pore shape is a straight channel in the five
commercially available membranes (Cases 1 to 5), there are two main determinants of the
porosity: pore size and pore density. In Cases 2 to 4, the membranes have the similar pore
density but different average pore size. When average pore size increased from 400 nm in
Case 3 to 1 µm in Case 4, the SEAP expression largely increased while the cell viability
slightly decreased. Although the average pore size in Case 2 is 3 µm, three times larger
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than 1 µm in Case 4, there is no obviously difference observed in the transfection results
in both cases. For Cases with the same pore size but different pore density (e.g. Cases 1
and 3, Cases 2 and 5), either the SEAP expression or the cell viability is similar. Based on
the value of pore size and pore density, the porosity was calculated and the value is given
in Table 4.3. The transfection results can be divided into two groups: Group A includes
Cases 2, 4, 5 with porosity larger than 1% and Group B includes Cases 1 and 3 with less
than 0.5% porosity. The total SEAP activity in Group A is about one third higher than
that in Group B, while losing 10 ~ 15 % cell viability.
Transfection experiments of mouse ES cells using top membrane with either the
micro-nozzle array or the straight micro-channel array were also carried out to determine
the effect of pore shape of top membrane on the MSE design. Due to the limitation of
femtosecond laser micro-fabrication, the minimum distance between two converging
micro-nozzles (center to center) was 10 µm. It means the maximum pore density of the
micro-nozzle array is about 6101× pores/cm2, and thus there are 1 ~ 2 pores per cell. To
minimize the effect of porosity, Case 5 in Table 4.3, was used for the corresponding
straight micro-channel comparison. No significant difference of the transfection results
between two cases was observed (Figure 4.12). The results showed evidence that for the
top membrane the porosity effect is more important than micro-pore shape effect on cell
transfection.
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Figure 4.12: Comparison of top membrane with different micro-pore size and micro-pore
density in MSE: (a) transfection efficiency and (b) cell viability of mouse ES cells 24
hours after MSE.
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4.3.4.3 Effect of top membrane location
Mouse ES cell transfection by pSEAP with different top membrane positions
(Table 4.4) was also investigated. As shown in Figure 4.13, SEAP expression of Cases I
to III decreased with increasing the distance between two membranes. The cell viability
of the three cases with a top membrane is better than the case without the top membrane
(Case IV). It means the existence of the top membrane helps to protect the cells during
the electroporation process.
It is noticed that besides slightly deformation due to the gravity effect of cell
itself, there’s a compression effect of the top membrane on cell when the distance
between two membranes is less than 5 µm. As observed from an inverted microscope, the
average diameter of the cells is 15 µm in the X-Y cross-section area in Case I, and 12 µm
at the remaining three cases. It has been reported that cell viability decreased when cells
are compressed to ~ 50 % of their original diameter. (Takamatsu and Rubinsky, 1999)
This explained why the cell viability of Case II was better than that of Case I.
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Table 4.4: Top membranes with different distance to cell binding membrane.
Cell dimension
Case No. With top
membrane
With Spacer Spacer thickness
(µm) Major axis
(X-Y)
Minor axis
(X-Z)
I Yes No N/A 15 4.5
II Yes Yes 10~12 12 7
III Yes Yes 16~20 12 7
IV No No N/A 12 7
Note: A PET membrane (BD Biosciences, San Jose, CA) was used as the top membrane
with an average pore size of 3 µm. The spacers were spin-coated PCL membranes with a
3 mm diameter hole in the middle.
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Figure 4.13: Effect of top membrane location in MSE on cell transfection: (a)
Transfection efficiency and (b) cell viability of mouse ES cells 24 hours after
electroporation.
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4.3.3 Comparison of MSE with bulk electroporation and nucleofection
pSEAP transfection of mouse ES cells by micro-nozzle array enhanced MSE,
bulk electroporation using Bio-Rad Gene Pulser, and nucleofection was compared
(Figure 4.14). Although the transfection result of MSE was still not as good as that of
nucleofection using unknown cell-specific reagents, the amount of SEAP expression
mediated by the MSE method was higher than that in bulk electroporation by Bio-Rad
Gene Pulser XCell system using the same electroporation buffer. The main reason is that
the MSE method is able to pre-concentrate the genes near the cell surface and to focus the
electric field strength around the micro-pores for better gene transport during
electroporation. Also, the electric field strength applied in MSE was 150 V/cm, much less
than 500 V/cm used in bulk electroporation. Therefore, mouse ES cells experienced an
average survival rate of ~ 75 % in MSE with an initial cell number of 4101× , similar to
bulk electroporation with 6101× initial cells and much higher than bulk electroporation
with 5101× initial cells.
MSE provided the high transgene efficiency and cell viability, because the area of
the cell membrane with the highest transmembrane potential is exactly the same location
that negatively charged DNA molecules could permeate into the cells (Golzio et al.,
2002b). As comparison, a much larger area of cell surface experiences high electric field
strength in the bulk electroporation, and DNA molecules can’t transport across most of
the affected cell surface. Since only a small area of cell membrane was affected during
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MSE and each cell experienced a similar electric field with the micro-array, a very small
number of cells could be uniformly transfected.
(Continued)
Figure 4.14: Comparison of mouse ES cell transfection by micro-nozzle array enhanced
MSE, bulk electroporation by Bio-Rad Gene Pulser, and nucleofection. (a) Transfection
efficiency and (b) cell viability 24 hours after electroporation. From left to right, bulk
electroporation with initial input cell number of 6101× ( ) and 5101× ( ); micro-nozzle
enhanced MSE with initial input cell number of 4101× ( ); and nucleofection with initial
input cell number of 6101× ( ).
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Figure 4.14 continued
A hundred of mouse ES cells trapped on a 1010 × micro-nozzle array were
transfected by pmaxGFP. Mouse ES cells were analyzed 24 hours after electroporation
by phase contrast and fluorescence microscopy using a GFP filter. The best result is
given in Figure 4.15. Almost all the cells were transfected, and remained alive. This
indicated the potential of micro-nozzle array enhanced MSE for hard-to-harvest cells.
However, it is very difficult to get such a perfect result, because it is hard to handle such
a small amount of cells.
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Figure 4.15: GFP transfection of mouse ES cell by micro-nozzle enhanced MSE. A
hundred of cells were trapped on a 1010 × micro-nozzle array, and (a) phase contrast and
(b) fluorescent images were taken 24 hours after electroporation.
4.4 Simulation of transmembrane potential distribution
Although there are still arguments on the creation and evolution of nano-pores on
cell membrane during the electroporation, it is well established that the size of nano-pore
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is directly related to the value of transmembrane potential. (Weaver and Chizmadzhev,
1996) When the transmembrane potential is lower than the threshold value, there is no
formation of nano-pores in cell membrane. In a successful gene electrotransfection
process, nano-pores should be large enough for gene to diffuse through. A higher
transmembrane potential leads to larger nano-pores formed on the cell membrane, thus
the more DNA molecules can be delivered to achieve the higher transgene efficiency.
(Krassowska and Filev, 2007) Therefore, we can propose the possible reason for the
results in Sections 4.3.2 and 4.3.3 through the calculated distribution of transmembrane
potential for bulk electroporation, localized electroporation, and different set-ups of
MSE.
4.4.1 Three-layer model
Since the distribution of transmembrane potential is extremely important to the
cell transfection, the consideration of cell membrane should be indispensable in the
numerical simulation of cell electroporation. In this study, we simplify the cell with a
three-layer (or single-shell) model and solved the equations in the cytoplasm, the external
medium, and the cell membrane, respectively.
The Laplace equation of the three-layer model (Stewart et al., 2005; Zudans et al.,
2007) was used to calculate the distribution of electric field and transmembrane potential
(the potential difference across the cell membrane) of a single cell in bulk electroporation
and MSE:
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0)( =∇⋅∇ ii φσ (4.2)
0)( =∇⋅∇ ee φσ (4.3)
0)( =∇⋅∇ mm φσ (4.4)
where:
φ : electric potential,
σ : electric conductivity,
Subscript i: cytoplasm,
Subscript e: external medium,
Subscript m: cell membrane.
Correspondingly, there are two boundary conditions (B.C.s) for two interfaces,
i.e., the external interface between the external medium and the cell membrane, and the
inner interface between the cytoplasm and the cell membrane.
nn
mm
ee
∂
∂=
∂
∂ φσ
φσ
(4.5)
me φφ =
B.C.s on external interface
(4.6)
nn
mm
ii
∂
∂=
∂
∂ φσ
φσ
(4.7)
mi φφ =
B.C.s on internal interface
(4.8)
In our experiment, low-conductive PET membrane was used, and thus the electric
conductivity of the membrane can be neglected. Therefore, all the solid walls were
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treated as insulators. Since no electric field can penetrate into the walls, different
potentials are imposed at inlet and outlet in order to create a voltage drop. Once the
electric potentials were calculated, the electric field E could be known at different layers:
φ−∇=E (4.9)
The transmembrane potential was obtained as
)()( inlmexlmm SSV φφ −=∆ (4.10)
where
S: surface of cell membrane;
Subscript exi: external;
Subscript inl: internal.
Compared with the two-layer model, the three-layer model is more complicated
and very dense mesh needs to be generated near the cell membrane, but it can be easily
applied to calculate the transmembrane potential around electroporated cells with nano-
pores formed on the cell membrane.
4.4.2 Two-dimensional (2-D) simulation process
A two-dimensional (2-D) simulation with the three-layer model was carried out
using commercial FEM software, COMSOL (Mathworks, Natick, MA), and parameters
used in the simulation are given in Table 4.5.
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Table 4.5: Parameters of the three-layer model (Kotnik et al., 1997)
Symbol Value Definition
d nm 5 Cell membrane thickness
eσ
mS /2.0 Electric conductivity of external medium
iσ
mS /2.0 Electric conductivity of cytoplasm
mσ mS /105 7−× Electric conductivity of cell membrane
For a single cell in bulk electroporation, Zudans et al. (2007) have shown that 2-D
simulation results agree well with the experiments, and thus the accuracy of the 2-D
numerical simulation has been verified. The analytical solution of transmembrane
potential for a 2-D spherical cell was obtained (Appendix B):
)cos(θfERVm =∆ (4.11)
where
E: external electric field strength, V/cm;
R: radius of the cell, µm;
θ : angle between Eext and the point on the cell membrane;
f: shape factor, f = 2.
Mouse ES cells with an average diameter of 10 µm were used in the experiments.
In bulk electroporation, cells are suspended in buffer solution, and thus are simplified as a
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sphere shape with the diameter of 10 µm. In localized electroporation and MSE, the cell
is sit down on a flat surface, and slightly deformed as a result of the gravity. As observed
from inverted microscope, the average diameter of the cells is 12 µm in the X-Y cross-
section area, and thus we simplify its shape as an ellipse with the major axis 12 µm and
the minor axis 7 µm in localized electroporation. It is noticed that besides slightly
deformation due to the gravity effect of cell itself, there’s compression effect of the top
membrane on cell when the membrane distance is less than 5 µm. From the top view, the
average diameter of the cells is 15 µm, and thus we simplify its shape as an ellipse with
the major axis 15 µm and the minor axis 4.5 µm in MSE. 2-D simulation of
transmembrane potential for an oval cell was preformed using the finite element methods,
as used in Agawal et al. (2007). The mesh of finite elements was generated, and the
electric potential inside and outside the cell was then computed by solving Eq. (4.10).
4.4.3 Simulation results
4.4.3.1 Effect of cell shape
The transmembrane potential distribution of four cases with different top
membrane location (Table 4.4) was calculated and the results are presented in Figure
4.16. The cell shape is simplify as an ellipse with the major axis 15 µm and the minor
axis 4.5 µm in Case I, while in Cases II to IV the major axis is 12 µm and the minor axis
7 is µm (Figure 4.16(a)). The perk value of the transmembrane potential at Case I is
higher than the other three case as a result of the shape effect, which explained the
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transfection results in Figure 4.13 that better transfection efficiency in Case I than the
other three cases.
a
(Continued)
Figure 4.16: Simulation comparison of top membranes with different pore size, pore
density, and pore shape. (a) Schematic diagram of Cases I to IV, from left to right, with
electric field lines across/around a single cell; (b) calculated transmembrane potential
distribution. θ is the angle around cell surface.
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Figure 4.16 continued
b
4.4.3.2 Effect of porosity and pore shape of top membrane
Figure 4.17 shows the calculated transmembrane potential distributions of
different top membranes as described in Table 4.3. According to the calculation in
Figure 4.17, the membrane with lager pores at a certain pore density has higher
transmembrane potential; for a fixed pore size, the membrane with a higher pore density
has a higher transmembrane potential in the model. The transmembrane potential at θ = π
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increased with the increase of the porosity in the simulation results (Figure 4.17). The
transfection results in Figure 4.12 are divided into two groups, indicating that there is a
threshold transmembrane potential to permeate cell membrane. Since Cases 2, 4 and 5
have better transfection than Cases 1 and 3, the critical transmembrane potential is around
0.4 V according to the calculation in Figure 4.17.
Figure 4.17: Simulation comparison of transmembrane potential distribution of top
membranes with different pore size, pore density, and pore shape.
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4.4.3.3 Converging micro-nozzle vs straight micro-channel
For simplicity, we assume micro-channels on the top membrane are uniformly
distributed in the simulation. The thickness of top membrane is 10 µm, and the size of a
micro-channel is ~1 µm. According to the pore density of 6106.1 × pores/cm2 given in the
manufactory’s instruction, the center-to-center distance between two micro-channels is
7.7 µm. For the cell binding membrane, the thickness is 12 µm and the center-to-center
distance between two micro-nozzles or micro-channels is 20 µm. The simulation results
are given in Figure 4.18.
Figure 4.18(a) shows that the electric field is concentrated around the micro-hole,
and more electric field lines are forced to penetrate through the cell at the small-end of
the micro-nozzle. Correspondingly, the transmembrane potential at the small-end of the
micro-nozzle is much higher than that of the straight channel as shown in Figure 4.18(b).
This explains why our micro-nozzle MSE can provide better delivery efficiency than the
micro-channel MSE, as shown in Figure 4.11.
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Figure 4.18: Simulation comparison between support membranes with micro-nozzles
and micro-channels (a) electric potential distribution and electric field lines across/around
a single cell near a micro-nozzle (left) and micro-channel (right); (b) calculated
transmembrane potential distribution. θ is the angle around cell surface.
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4.4.3.4 Comparison of micro-nozzle enhanced sandwich electroporation with
bulk electroporation
Micro-nozzle enhanced MSE was also compared with bulk electroporation. The
transgene efficiency doesn’t depend on the total area of cell membrane with openings, but
on the effective area facing the cathode. (Golzio et al., 2002b) Although a much larger
area of cell surface experiences a high electric field strength in the bulk electroporation
(Figure 4.18(b)), the area of the cell membrane with the highest transmembrane potential
in MSE is at exactly the same location that negatively charged DNA molecules could
permeate into the cells. Furthermore, since only a small area of cell membrane was
affected in MSE and each cell experienced a similar electric field with the micro-array, a
very small number of cells could be uniformly transfected. In addition, the size of nano-
pores on the cell membrane surface is smaller, and thus DNA molecules can’t transport
across most of the affected cell surface.
4.5 Conclusion
In this study, we demonstrated the use of a femtosecond laser fabricated micro-
nozzle arrays on a gelatin-coated PET membrane for membrane sandwich electroporation
(MSE). Using micro-nozzle array enhanced MSE, we observed high and uniform gene
transfection, and good cell viability of mouse ES cells compare to the bulk
electroporation. The ability to treat a small number of cells (i.e. a hundred) offers great
potential to work with hard-to-harvest patient cells for pharmaceutical kinetic studies.
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Numerical calculation of transmembrane potential qualitatively explains the
observed differences among different cases of MSE and bulk electroporation. Since
there’s a reasonably well correlation between transfection results and transmembrane
potential calculations, the simulation process with the threshold experiments can be used
to predict the transfection results, and thus largely reduced the trial-and-error window
size.
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CHAPTER 5: NANOFIBER BASED MEMBRANE SANDWICH
ELECTROPORATION
5.1 Introduction
Over the last decade, the clinical applications of genetically modified embryonic
stem (ES) cells have increased considerably. For clinical trails, it is essential to have high
transfection efficiency as well as high cell viability. It has been showed that ES cells,
especially human ES cells, grow better when forming highly compact colonies. (Amit et
al., 2000) However, commercially available electroporation systems, especially the
leading Nucleofection, are based on single cell suspension. As a result, it is hard to treat
cell colonies using conventional bulk electroporation, especially if repeated transfection
is required. In this chapter, the membrane sandwich electroporation (MSE) method is
integrated into cell culture such that ES cells can form desirable colonies, be transfected,
and further cultured on a same substrate before and after MSE.
Electrospinning is a simple and versatile technique that can produce a porous
scaffold comprised randomly submicron-diameter polymeric fibers (or nanofibers) (Chew
et al., 2006). Electrospun nanofiber scaffolds characterized by a high surface area for cell
attachment and three-dimensional (3-D) microenvironment for cell-cell interaction
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provide stronger topographic cues by mimicking the filamentary extracellular matrics,
and enhance cell adhesion and proliferation compared to micro-porous polymer
membrane. (Wnek et al., 2003; Yashimoto et al., 2003) Poly(ε-caprolactone) (PCL), a
Food and Drug Administration (FDA) approved biocompatible polymer, has been widely
used as a tissue engineering scaffold to support cellular in-growth and proliferation for
the generation of biological tissues. However, it was reported that cell adhesion,
migration, proliferation, and differentiation were reduced due to its poor hydrophilicity.
(Kohl et al., 2005; Ingber, 2005) As a natural biopolymer derived from collagen by
controlled hydrolysis, gelatin is blended with PCL to obtain a scaffold with good
biocompatibility and improved mechanical, physical and chemical properties. (Chong et
al., 2007; Zhang et al., 2005) Additionally, electrospun PCL/gelatin nanofiber scaffolds
may serve as safe substitutes to Matrigel, a gelatinous protein mixture secreted by mouse
tumor cells. In a recent study, Gauthaman et al. (2008) evaluated the influence of
electrospun nanofibrous (PCL/collagen and PCL/gelatin) scaffolds for human ES cell
proliferation. Increased colony-formation, self-renewal properties, undifferentiation and
retention of stemness were observed in their study.
A combination of MSE and electrospun PCL/gelatin nanofiber scaffolds allows
gene transfection at different time points during cell colony formation without repeating
cell trypsinization, and thus may enhance the cell viability. Since the size of ES cell
colonies can influence pluripotent cell differentiation trajectories (Bauwens et al., 2008;
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Tsuruma et al., 2008), cell colony formation may be further controlled with micro-
patterned spacer bonded on nanofiber surface.
5.2 Materials and methods
5.2.1 Cell culture
NIH 3T3 fibroblasts and mouse embryonic stem (mES) CCE cells were cultured
and prepared according to the procedure described in Sections 3.2.2 and 4.3.1.2.
5.2.2 Fabrication and characterization of nanofiber scaffolds with micro-well
spacers
5.2.2.1 Preparation of electrospun poly (ε-caprolactone) (PCL) /gelatin nanofiber
scaffolds
A 6.7 % (w/v) solution of poly (ε-caprolactone) (PCL; Sigma-Aldrich, Mw =
65,000) in 1,1,1,3,3,3-Hexafluoro-2-propanol (HFIP, Sigma-Aldrich) and a 6.7 % (w/v)
solution of porcine gelatin (Sigma-Aldrich, Catalog. No. G2500) in HFIP was prepared
by stirring overnight at room temperature, separately. The two solutions were mixed
together at a 1:1 weight ratio, and stirred for 2 minutes. The mixture was then placed in a
60 mL syringe with a 20 gauge blunt tip needle, and electrospun using a high voltage DC
power supply (Glassman High Voltage, High Bridge, NJ) set to -26 kV voltage and 15
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mL/h flow rate. (Johnson et al., 2008, Lim et al., 2009) The support membrane was
placed on an aluminum foil with a 20 cm distance between the tip and the collector. The
electrospun sheet with nanofibers was deposited onto the support membrane, and the
thickness of the sheet was controlled by the electrospinning time. The electrospun sheet
was then placed in a vacuum oven overnight to ensure removal of residual HFIP. Figure
5.1 shows the electrospining process.
Figure 5.1: Schematic diagram of fiber formation by electrospining process where a
droplet of a polymer solution is elongated by a high electrical field.
(http://nano.mtu.edu/Electrospinning_start.html)
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5.2.2.2 Fabrication of PCL /gelatin nanofiber scaffolds with polystyrene (PS)
micro-well arrays
An 8 ~ 10 µm thick polystyrene (PS) (Sigma-Aldrich, melt flow index 4.0) sheet
with arrays of 100 or 300 µm diameter micro-wells was fabricated via soft lithography
(Figure 5.2). Standard photolithography using negative tone photoresists, NANOTM SU-8
(Microchem Corp., Newton, MA), was used to achieve an 8 ~ 10 µm thick master mold
with micro-well arrays. The fabrication parameters, including spin speed, exposure dose,
post-exposure bake times, and develop time, were set up according to the manufacturer’s
suggestion. The master mold was then used to create poly(dimethyl siloxane) (PDMS)
stamp with micro-pillar arrays. The PDMS stamp was spin-coated with 15 ~ 20 %
polystyrene (PS) solution in anisole under spin-speed of 3,000 rpm for 1 minute. The
PDMS molds were then placed on a hotplate at 100°C for 5 minutes to drive off the
residual solvent and anneal the PS sheet. After cooling, the PS sheet left in-between the
pillars was manually peeled off. The PS sheet with micro-well arrays was thermally
bonded to PCL/gelatin nanofibers with support membrane at 80 ~ 90°C.
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Figure 5.2: Schematic of fabricating electrospun nanofiber scaffold with polystyrene
(PS) micro-well arrays. (1) PDMS stamp with micro-pillar arrays; (2) Drop-cast PS
solution; (3) PS solution is spin-coated and it de-wets on the surface of the PDMS stamp;
(4) PS in-between the features is removed; (5) PS micro-well arrays were bonded to
electrospun nanofiber scaffold by thermal bonding. (Gallego et al., In preparation)
5.2.2.3 Structure characterization by scanning electron microscopy (SEM)
SEM was used to observe the structure of PCL/gelatin nanofiber scaffolds. The
samples were coated with 15 nm of osmium (model OPC-80T, SPI Supplies) prior to
viewing in a scanning electron microscope (Sirion FEI). The use of osmium plasma
deposition instead of gold or gold-palladium sputtering eliminated concerns regarding
PCL melting and allowed for higher resolution imaging of the fiber surface. SEM images
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of the surface of randomly distributed PCL/gelatin nanofibers and nanofiber scaffolds
with 300 µm PS micro-wells are shown in Figure 5.3.
Figure 5.3: SEM image of PCL/gelatin nanofiber scaffolds with 300 µm PS micro-wells.
5.2.3 Experimental set-up for nanofiber based MSE
The experimental set-up for electrospun nanofiber based MSE is shown in Figure
5.4. The cell binding substrate consists of a support membrane and a thin sheet of
electrospun nanofiber scaffold. Membranes with different properties were tested as the
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support membrane for nanofiber scaffolds. The electrospun fibers are randomly
distributed poly-ε-caprolactone (PCL) / gelatin blends (weight ratio: 50:50, mean fiber
diameter: 300-400 nm, thickness: 5-20 µm), directly bonded to 8 ~ 10 µm thick
polystyrene (PS) spacer with 100-500 µm micro-well arrays.
Figure 5.4: Schematic drawing of electrospun nanofiber based MSE.
5.2.4 Electric Resistance Measurements
The electric resistance of a sample was determined by the ratio of voltage to
current at room temperature, in accordance with Ohm's law:
I
VR = (5.1)
where:
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R: electric resistance, KΩ
V: voltage, V
I: current, mA.
The sample was placed in the central reservoir, and the inlet and outlet channels
and reservoirs were filled with high glucose DMEM for mouse ES cells. Two wire
electrodes were placed in the inlet and outlet reservoirs, and connected to PowerPac
Basic Power Supply (Bio-Rad, Catalog No. 164-5050). A DC voltage that increased from
a 30 to 60 V with increment of 10V was applied to the system, and the value of the
current at each specific voltage was recorded. The graph was plotted with voltage as Y-
axis and current as X-axis, and a linear curve was fitted to the data points. The calculated
slope of the line is numerically equal to the electric resistance of the sample.
5.2.5 Electroporation procedure
5.2.5.1 Single cell electroporation
For NIH 3T3 fibroblasts, electroporation followed the same procedure described
in Section 3.2.4.
For mES cells, single cell suspension was obtained using 0.25% trypsin with
EDTA·4Na. Bulk electroporation and nucleofection followed the same procedure given
in Section 4.3.1.4, and nanofiber based MSE followed the procedure described in Section
3.2.5.2.
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5.2.5.2 Cell colony electroporation
Mouse ES cell colony suspension was obtained using 0.05% trypsin with
EDTA·4Na (Catalog No. 25300).
Bulk electroporation and nucleofection followed the procedure given in Section
4.3.1.4.
For nanofiber based MSE, cells were seeded in the micro-wells on the cell
binding substrate and grow 1 to 2 days to form colonies. A PET track-etched membrane
with an average pore size of 3 µm was placed over the colonies, and the cell colonies
were sandwiched between nanofiber scaffold and PET membrane, which are sealed
together. The whole MSE disk was then sealed at the center reservoir of the MSE device
by sealing tape. The bottom channel and reservoirs (cathode side) were loaded with high
glucose DMEM, and the top channel and reservoirs were then loaded with high glucose
DMEM with 100 ng/µL DNA molecules. A low DC electric field of 3 V/cm was applied
to the system for 5 seconds to concentrate DNA molecules in the micro-pores, and the
cells were then electroporated by a single exponentially decay pulse at 150 V/cm and 500
µF.
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5.2.6 Assays for transfection efficiency and cell proliferation
As described in Sections 3.2.6 and 3.2.7, the transfection efficiency of pGFP in
NIH 3T3 fibroblasts and mouse ES cells was qualified by the percentage of cells with
green fluorescence, while the transfection efficiency of pSEAP was quantified by the
activity level of secreted alkaline phosphatase by the transfected cells, determined by a
colorimetric assay based on the hydrolysis of p-nitrophenyl phosphate (pNPP).
AlamarBlue® cell proliferation assay (Molecular Probes, Catalog No. DAL-1100)
was used to quantify the cell viability at 24 and 48 hours post-electroporation. The
alamarBlue® dye was diluted in the culture media at 1:10, and the work solution was
then added to the culture wells. The plate was shaken gently and incubated for 3 hours at
37°C with 5% CO2. The fluorescence detection was performed using a fluorescence
excitation wavelength of 570 nm, and the reading at emission wavelength of 595 nm was
recorded. (Nociari et al., 1998) Fluorescence background can be corrected by including
negative control wells on each plate to measure the fluorescence from culture medium in
the absence of cells. A set of positive control wells containing untreated cells performed
in sister cultures was settled as 100% cell viability. The cell viability of each sample was
calculated as
%100595595
595595×
−
−=
backgroundcell
backgroundsmaple
ODOD
ODODViabilityCell (5.2)
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5.2.7 Cell morphology characterization by confocal microscopy
Cell morphology on PCL/gelatin nanofibers was characterized by laser scanning
confocal microscopy (Zeiss LSM 510 Meta, Zeiss, Germany). Cells were fixed directly
with 70 % ethanol solution for 1 hour at room temperature, and labeled with the red-
fluorescent nucleic acid stain, propidium iodide (PI; Molecular Probe, Catalog No.
P3566). After incubated with 2 µg/ml PI for 15 minutes at 37°C and washed twice with
D-PBS, cells were then visualized by employing the excitation filter, 543 nm.
5.3 Optimization of nanofiber based membrane sandwich electroporation
5.3.1 Effect of support membrane
Preliminary experiments were carried out using electrospun nanofiber scaffolds
without any support membrane as a cell-binding substrate, and the results showed that it
is difficult to transfer the nanofiber scaffolds to membrane sandwich electroporation
(MSE) platform after overnight cell culture. To make the cell-binding substrate stronger,
a micro- / nano-porous membrane was used as a support membrane.
A track-etched polyethylene terephthalate (PET) membrane (average pore size: 3
µm) with a layer of PCL/ gelatin nanofibers was tested. Although the cell viability
increased, the transfection efficiency decreased compared to the transfection results using
PET membrane only (Figure 5.5(a,b)). Since the electric resistance of the cell-binding
substrate may increase by adding a layer of low-conductance nanofibers, the resistance
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measurement using PET membrane with and without nanofibers was carried out and the
data of voltage against current are plotted as shown in Figure 5.5(c). As the resistance is
determined by the gradient of the straight fitting line, a steeper curve corresponds to
increased resistance. By adding a layer of nanofiber scafflolds, the resistance value
almost doubled over that using the PET membrane only.
(Continued)
Figure 5.5: Comparison of different cell-binding substrates used for membrane
sandwich electroporation. The transfection efficiency (a) and cell viability (b) of mouse
embryonic stem cells, and the resistance (R) of MSE disk (c) were presented using PET
membrane only ( ), aluminum oxide membrane only ( ), PET membrane with
nanofibers ( ), and aluminum oxide membrane with nanofibers ( ).
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Figure 5.5 continued
To reduce the resistance of the MSE disk, AnodiscTM aluminum oxide membrane,
was tested as a support membrane. The properties of the membrane from the
manufactory’s website are given in Table 5.1. Without nanofiber scaffolds, the resistance
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of an aluminum oxide (Al2O3) membrane is similar to that of a PET membrane (Figure
5.5(c)); however, cell viability using the Al2O3 membrane significantly decreased
(Figure 5.5(b)). With a nanofiber scaffold, the resistance of the MSE disk using the
Al2O3 membrane as the support membrane is much lower than that of the PET
membrane; and good transfection results (Figure 5.5(a, b)) were achieved by using the
nanofiber scaffold supported with the Al2O3 membrane. Besides the material difference,
the Al2O3 membrane has more than 4 times higher porosity than the PET membrane.
Table 5.1: Properties of three different types of membranes used as support membrane.
Material
Pore size
(µm)
Porosity
(%)
Thickness
(µm)
Note
PET 3 6 10 BD Biosciences,
San Jose, CA, USA
Aluminum oxide 0.2 25 60 Anodisc™, Waltman/GE,
Piscataway, NJ, USA
Polycarbonate 0.2 20 20 Isopore™,
Millipore, USA
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To understand why Al2O3 membrane works better, another polymeric membrane,
polycarbonate (PC) membrane (Table 5.1) with a similar porosity as the Al2O3
membrane were tested. There was not any obvious difference observed in the electric
resistance measurements and transfection resultus between PC and PET membrane with
nanofiber scaffolds. Therefore, the difference in electric properties is more important than
the porosity difference. The most possible reason is that Al2O3 membrane became
electrically conductive under the external electric field, and consequencely reduced the
electric resistance of nanofiber scaffold covered support membrane.
5.3.2 Effect of nanofiber thickness
As shown in Figure 5.5, the resistance of the cell-binding substrate was increased
by adding PCL/gelatin nanofiber scaffold. As the PCL/gelatin nanofiber scaffold is a
non-conductive polymeric/protein matrix, the resistance of the cell-binding substrate
increased with increasing the thickness of the nanofiber scaffold layer, which was
controlled by the electrospinning time (Table 5.2). If the nanofiber layer was too thick
( 15≥ m), the transfection efficiency (Figure 5.6(a)) decreased greatly as the result of
significant resistance increase. However, if the nanofiber layer is too thin (≤ 5 µm), a
large number of cells would contact the Al2O3 membrane and thus died during the
electroporation process (Figure 5.6(b)). The optimal nanofiber scaffold layer thickness is
10 µm in the MSE set-up.
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Table 5.2: Thickness and corresponding resistance of nanofiber layer controlled by
electrospinning time.
Electrospining time (min) Thickness (µm) Resistance (KΩ)
0 0 5.46
1 4~5 6.19
2 8~10 6.46
4 15~20 8.11
(Continued)
Figure 5.6: Effect of electrospun nanofiber thickness on the transfection efficiency (a)
and cell viability (b) of mouse embryonic stem cells. The thickness of nanofiber layer
corresponds to electrospinning time as shown in Table 5.2.
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Figure 5.6 continued
5.4 Nanofiber based MSE of mouse embryonic stem (ES) cell colony
5.4.1 Mouse ES cell colony formation with controlled size
Figure 5.7 shows confocal images of mouse ES cells after 24 and 48 hours
culture on PCL/gelatin nanofiber scaffolds with and without 100 µm PS micro-wells.
With cell seeding density at 5,000 / mm2, mouse ES cells formed colonies with a uniform
size after 24 hours culture on PCL/gelatin nanofiber scaffolds with micro-wells. After 48
hours culture, mouse ES cells were expanded outside the 10 µm thick micro-wells.
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Figure 5.7: Confocal images of mouse ES cell colonies after cultured 24 (a, c) and 48 (b,
d) hours on randomly distributed PCL/gelatin nanofiber scaffolds without (a, b) and with
(c, d) 100 µm PS micro-wells. The cell seeding density was 5,000 / mm2. Cells were
fixed with 70% ethanol and stained with PI dye. The length of the standard bars is 100
µm.
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5.4.2 Nanofiber based MSE with vs without micro-well spacer
Mouse ES cells were seeded on the cell binding substrates, PCL/gelatin nanofiber
scaffolds with and without a micro-well spacer, respectively. The cell seeding density
was 5,000 / mm2. As shown in Figure 5.7, the PCL/gelatin nanofiber surface with 100
µm micro-wells were fully covered with mouse ES cells after 24-hour culture. After 48-
hour culture, mouse ES cells were grown beyond confluence on the flat nanofiber surface
and expanded outside the 10 µm thick micro-wells. To perform nanofiber based MSE at
confluent mouse ES cells with no more than double layers, SEAP transfection was
carried out after 24-hour pre-culture. Since the solid spacer blocked the passage of the
electric field, the amount of SEAP expression from mouse ES cell colonies cultured on
PCL/gelatin nanofiber scaffolds with the micro-well spacer is much higher than that
without micro-well spacer (Figure 5.8). More importantly, the size and thickness of cell
colonies was more uniform with the micro-well spacer (Figure 5.9).
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Figure 5.8: SEAP transfection of mouse embryonic stem cells by nanofiber based MSE
without ( ) and with ( ) micro-well spacer, bulk electroporation by Bio-Rad Gene Pulser
X-Cell system ( ), and nucleofection ( ): (a) transfection efficiency (b) cell viability 24
hours after electroporation.
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Figure 5.9: Confocal images of mouse ES cells 6 (a, c) and 30 (b, d) hours after
nanofiber based MSE without (a, b) and with (c, d) 100 µm micro-well spacer. Mouse ES
cells were fixed with 70% ethanol and stained with PI dye. The length of the standard
bars is 100 µm.
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5.4.3 Nanofiber based MSE vs Bulk electroporation of cell colony
SEAP transfection of mouse ES cell colonies by nanofiber based MSE was further
compared with bulk electroporation and nucleofection. Nanofiber based MSE with 100
µm PS micro-well spacers was performed as described in Section 5.4.2. For bulk
electroporation and nucleofection, half million ( 5105× ) mouse ES cells were seeded in
each well of the gelatin coated 6-well plate. After 24 hours pre-culture, mouse ES cell
colonies were harvested, and the total cell number from each well was around one million
( 6101× ). Colonies of mouse ES cells collected from each well were used for one bulk
electroporation or nucleofection sample. With only one percent of initial input cells, the
transfection efficiency of nanofiber based MSE with the micro-well spacer was
comparable to bulk electroporation (Figure 5.8(a)). Also, the electric field strength
applied in nanofiber based MSE was 150 V/cm, less than one third of 500 V/cm used in
bulk electroporation. Consequently, mouse ES cells experienced an average survival rate
of ~ 66 % in nanofiber based MSE, much higher than ~ 42 % in bulk electroporation
(Figure 5.8(b)). Although the transfection efficiency of nanofiber based MSE was not as
good as nucleofection, the survival rate of mouse ES cell colonies using nanofiber based
MSE was much better, ~ 35 % higher than that of nucleofection.
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5.5 Nanofiber based MSE of NIH 3T3 fibroblasts
Previous studies (Rodolfa and Eggan, K. 2006; Lowry and Plath, 2008; Pera,
2009) have shown that transient expression of reprogramming factors into embryonic
fibroblasts is sufficient to obtain iPS cells. However, the very low success rate (less than
10 colonies per one million input cells) and a substantial degree of clone-to-clone
variation limited its clinical applications of iPS cells. In addition, the reprogramming
factors are diluted during cell division in the case of plasmid transfection. (Okita et al.,
2008) To achieve better reprogramming efficiency, longer exposure to programming
factors and thus repeated transfection is needed.
5.5.1 NIH 3T3 fibroblasts with micro-well spacer
NIH 3T3 fibroblasts were seeded on the PET membrane surface with and without
micro-well spacer. Figure 5.10 shows the phase contrast images of NIH 3T3 fibroblasts
after 48 hours culture. With the micro-well spacer, NIH 3T3 fibroblasts did not spread
out to the fiber shape as observed on flat cell culture surfaces, instead formed high
compact cell clusters, similar to the morphology of mouse ES cell colonies. This suggests
the potential of using MSE for obtaining induced pluripotent stem (iPS) by plasmid
transfection.
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Figure 5.10: Morphology of NIH 3T3 fibroblasts with (a) and without (b) 300 µm micro-
well spacer after 48 hours.
5.5.2 Repeated SEAP transfection of NIH 3T3 fibroblasts
The repeated SEAP transfection results of NIH 3T3 fibroblasts by MSE (Figure
5.11) demonstrate that cell viability significantly decreased, resulting in a decrease of the
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transfection efficiency after the 2nd treatment. Since fibroblast proliferation was found to
be much better on nanofiber scaffolds (Park et al., 2007), the cell-binding substrate was
changed to nanofiber scaffolds. Nanofiber based MSE was conducted at day 0 and 1, and
the transfection results are shown in Figure 5.11. With repeated nanofiber based MSE,
NIH 3T3 fibroblasts expressed SEAP stably. Using electrospun nanofiber scaffolds as the
cell-binding substrate, cell viability was also higher than that using micro-porous
membrane, because the nanofibrous surface provides better cell adhesion and faster cell
recover rate after electroporation. The combination of MSE and electrospun nanofiber
scaffolds allows repeated plasmid transfection at different time points during cell culture.
Furthermore, the micro-well spacer provides well-defined microenvironment to form cell
colony with uniform size, and thus can reduce the transfection difference from colony to
colony.
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Figure 5.11: Repeated SEAP transfection of NIH 3T3 fibroblasts using PET micro-
porous membrane () and electrospun PCL/gelatin nanofiber scaffolds (): (a)
transfection efficiency (b) cell viability at day 1 and 2 post-electroporation. Both cell-
binding substrates had the micro-well spacer.
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5.6 Conclusions
The successful combination of MSE and electrospun nanofiber scaffold allowed
better plasmid transfection during colony formation of mouse ES cells. The micro-well
spacer provides well-defined microenvironment to form cell colony with uniform size,
and thus can reduce the transfection difference from colony to colony. Furthermore, the
electrospun nanofiber scaffold as cell-binding substrate improved the cell survival and
recover rate of embryonic fibroblasts during repeated plasmid SEAP transfection,
demonstrating the potential for better iPS cell generation.
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CHAPTER 6: CONCLUSIONS AND RECOMMENDATIONS
6.1 Conclusions
A new membrane sandwich electroporation (MSE) approach was developed and
demonstrated using plasmids GFP and SEAP as reporter genes. Several cell lines, such as
NIH 3T3 fibroblasts and mouse embryonic stem (ES) cells were tested and a significant
improvement in transgene expression and cell viability was observed comparing to
current electroporation techniques. In the MSE method, the focused electric field
enhances cell permeabilization at a lower electric voltage, leading to high cell viability.
Furthermore, the sandwich membrane configuration is able to provide better gene
confinement near the cell surface, facilitating gene delivery into the cells.
We also demonstrated the use of femtosecond laser fabricated micro-nozzle arrays
on a gelatin-coated PET membrane for MSE. Using micro-nozzle array enhanced MSE,
we observed higher and more uniform gene transfection with excellent cell viability of
mouse ES cells comparing to that achieved with the bulk electroporation methods.
The strategy of ex vivo human-stem-cell-based therapy for tissue regeneration
includes transfecting human stem cells in vitro to express a certain transgene or to
differentiate to certain cells, and then implanting them in vivo under pharmaceutical or
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blood bank standards of Good Manufacturing Practice (GMP). Successful pre-clinical
trials of genetically modificated human stem cells using in vitro electroporation has been
demonstrated for bone repair (Aslan et al., 2006) and dentin formation (Nakashima et al.,
2004). Since cells or tissues from the patients are often very limited and therapeutic
materials such as plasmids and oligonucleotides are very expensive, the ability to treat a
small number of cells (i.e. hundreds) instead of millions offers great potential for hard-to-
harvest patient cells in patient-specific ex vivo gene therapy and in vitro pharmaceutical
kinetic studies.
We also successfully integrated the electrospun nanofiber scaffold as a cell-
binding substrate in MSE, i.e. nanofiber based MSE. With a micro-well spacer, uniformly
sized colonies of mouse ES cells were obtained, and plasmid transfection by
electroporation was performed during colony formation. Also, repeated plasmid SEAP
transfection of NIH 3T3 fibroblasts was tested, and better cell survival and recover rates
were observed comparing to that using a micro-porous membrane. Due to its ability to
repeated transfection with reprogramming factors, the nanofiber based MSE method has
potential for efficient iPS cell generation by repeated plasmid transfection.
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6.2 Recommendations
6.2.1 Individual cell array trapping
In this study, we demonstrated the ability to treat a small number of cells using
membrane sandwich electroporation (MSE), and observed high gene transfection and cell
viability comparing to current bulk electroporation techniques. Using a 10 x 10 micro-
nozzle array, we once achieved very exciting transfection results that almost all 100 cells
were transfected with pmaxGFP and remained alive. However, it is very difficult to
repeat such a perfect experiment because of difficulty to align every individual cell on a
micro-nozzle using the vacuum-assisted seeding process. More robust and reliable
techniques should be developed allowing precisely control of the position of individual
cells on a large cell array without causing cell damage. For example, optical trapping
using an optical tweezer array as shown in Figure 6.1 can be a potential solution. In this
method, an array of individual cells is optically trapped by an optical tweezer array
created by focused laser beam through a micro-lens array. By designing a suitable micro-
lens array with the distance of two adjacent optical traps matching that of the adjacent
micro-nozzles, the trapped cells can be aligned right on top of micro-nozzles.
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Figure 6.1: Schematic illustration of individual cell array trapping by an optical tweezer
array created by focused laser beam through a micro-lens array.
6.2.2 Cell membrane permeability experiments
Our numerical simulation of transmembrane potential distribution qualitatively
explained the effects of cell shape, porosity and pore shape of top and bottom membranes
on MSE. However, there remained large discrepancy in some cases. This is because the
electroporation mediated gene transfection process (Figure 6.2) includes two parts: (1)
cell membrane break-down and reseal, and (2) genes bounding to the cell membrane
during the electroporation and entering cell plasma by endocytosis. The simulation in
Chapter 4 provides the distribution of initial transmembrane potential under an imposed
external electric field. If the transmembrane potential is larger than the critical
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transmembrane potential, cell membrane becomes permeable. Therefore, the membrane
permeability experiments should be carried out to obtain the critical transmembrane
potential for each cell type in order to link the simulation with transfection prediction in
both MSE and bulk electroporation. A method using the spin-disk confocal microscopy
with appropriate fluidic system and cell impermanent dye, propidium iodide (PI), is being
developed in our lab. Time series studies to track how the dye enters the cell should be
further explored such that the critical transmembrane potential and the cell permeable
location can be measured. Furthermore, the electroporation experiments with different
size of fluorescein-labeled dextrans under different electric fields should be used to
quantify the relationship between the transmembrane potential and size of nanopores on
cell membrane.
Figure 6.2: Electroporation of a cell. The electroporation mediated gene transfection
process includes two parts: (1) cell membrane break-down and reseal, (2) genes bounding
to the cell membrane during the electroporation, and entering cell plasma by endocytosis.
(http://www.inovo.com)
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In Chapter 5, we have demonstrated the possibility of nanofiber based MSE for
iPS cell generation by repeated plasmid transfection. During the repeated electroporation,
the MSE disks have to be transferred from culture wells to the current fluidic device
(Figure 3.2), and then placed back to culture wells every time. Since the cell-binding
substrates could not be perfectly fixed on the current fluidic devices after culture in the
media, the electric field might leak through the edge of the MSE disk. To address this
problem, a new design that allows cell culture with the cell-binding substrate was
developed, and Figure 6.3 shows this latest MSE system including a MSE stage with two
MSE disks and an electroporation box. Each MSE disk consists of a bottom and a top
piece. The cell-binding substrate and nanofiber scaffolds with micro-well spacer can be
fixed on the bottom piece, and two bottom pieces can be cultured together in one well on
a 6-well plate. Advantages of this new design should be further investigated, particularly
for transfection of delicate cells such as human stem cells and iPS cells.
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Figure 6.3: The third generation of MSE system, including (a) a MSE stage with two
MSE disks and (b) an electroporation box, which is able to connect with one AC pulse
generator and one DC power supply. Each MSE disk consists of a bottom and a top piece.
(Designed and fabricated by Mr. Shi-Chiung Yu and Dr. Weixiong Wang, 2009)
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One advantage of the MSE method is able to provide better gene confinement
near the cell surface, facilitating gene delivery into the cells. Although a low DC electric
field was applied to pre-concentrate genes in this study, the negatively charged DNA
molecules tended to migrate away from the negatively charged cell membrane when the
applied electric field was switched from the low DC electric field to a high voltage AC
electric pulse during. This time lapse was around 3-5 seconds. To avoid this problem, a
low DC electric field should be imposed on DNA molecules before, during and after
electroporation. A new electroporation box (Figure 6.3(b)) is constructed, which is able
to connect with one AC pulse generator with one DC power supply. This new device and
box is our third generation MSE system, and currently is under investigation.
Comparing the single cell electroporation and cell colony electroporation results,
we found that single cell nucleofection provided much better transfection efficiency,
while our nanofiber based MSE provided the highest cell survival and recover rates.
Therefore, an ideal procedure for iPS cell generation may include single cell transfection
by nucleofection, cell seeding on nanofiber scaffolds with a micro-well spacer, and
repeated cell colony transfection by nanofiber based MSE. Since one of the most
important factors of successful nucleofection is the cell-specific electroporation buffer, a
combination of nucleofection solution with our MSE method could provide a synergistic
effect. This idea should also be explored in the future study.
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6.3 Possible ways of in vivo applications
One possible way of in vivo applications is the combination of MSE with the
cannula for thymic cancer or breast cancer treatment. A good example is thymomas.
Although rare, thymomasis the most common type of thymic cancer and is hard to detect
at the early stage. Thymoma arises from thymic epithelial cells, which cover the thymus.
The thymus is a small organ that locats in the upper chest just below the neck. Some
preliminary results of transfecting embryoid bodies using the MSE method (Appendix E)
demonstrated that only few outer layers (less than 3 layers) were transfected. Since it is
nearly impossible to remove thymoma by surgery, MSE may be very suitable for
treatment in such case, because our MSE method is able to treat thymoma derived from
the outer thymic epithelial cells without disturbing the inside thymus.
Another possible in vivo application of MSE is to use a double-balloon
enteroscope to treat the solid tumors in the gastrointestinal system. The double-balloon
enteroscope (or push-and-pull enteroscope) was developed by Yamamoto et al. in 2001
and has been used for diagnosis and control of bleeding with electrocoagulation
(Nishimura et al., 2004). For the malignant tumors that are hard to be surgically resected,
MSE with the double-balloon enteroscope is a promising technique for performing
electrochemotherapy or electrogenetherapy in targeted tissue with minimal the damage to
surrounding tissues. The outer-balloon should be designed using micro-porous
membrane, and the inner-balloon should carry out a bundle of micro-electrodes and drugs
and genes. When the outer-balloon wraps up the targeted tumor, the drugs or genes are
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able to be injected into the tumor by electroporation. Before in vivo treatment, in vitro
test of a tumor sample from the patient by MSE should be first investigated, followed by
pre-clinical testing in small and large animal models.
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Zeng, X.; Chen, J.; Sanchez, J. F.; et al. (2003) Stable expression of hourGFP by mouse embryonic stem cells: promoter activity in the undifferentiated state and during dopaminergic neural differentiation. Stem Cells 21: 647-653.
Zimmermann, U.; Vienken, J. (1982) Electric field-induced cell-to-cell fusion. Journal of
Membrane Biology 67: 165-182.
Zudans, I.; Agarwal, A.; Orwar, O.; Weber, S. G. (2007) Numerical calculation of single-cell electroporation with an electrolyte-filled capillary. Biophysical Journal 92(10): 3696-3705.
-148-
APPENDIX A: STANDARD CURVE
y = 72.68x
R2 = 0.9908
0
10
20
30
40
50
60
70
0.0 0.2 0.4 0.6 0.8 1.0
OD@405nm
SE
AP
Acti
vit
y (
mU
)
Figure A.1: Standard curve of SEAP activity (mU) vs absorbance reading at 405 nm
(OD@405nm). 100 µL of standard alkaline phosphatase solution of 0, 5, 10, 20, 40, and
60 mU and 25 µL of pNPP substrate solution were added into each well of a 96-well
plate. The plate was incubated in the dark for approximately 15 minutes at room
temperature, and read at the wavelength of 405 nm on a multi-well plate reader.
-149-
APPENDIX B: OPTIMIZATION OF BULK ELECTROPORATION AND
NUCLEOFECTION OF MOUSE EMBRYONIC STEM (ES) CELLS
Figure B.1: Effect of initial cell number on bulk electroporation of mouse ES cells
using Bio-Rad Gene Pulser X-Cell system: (a) transfection efficiency and (b) cell
viability 24 hours after BE.
0
20
40
60
80
1.00E+06 5.00E+05 1.00E+05
Initial Cell Number
To
tal
SE
AP
Acti
vit
y m
U/
Init
ial
10,0
00 s
eed
ing
cell
s
0
20
40
60
80
100
1.00E+06 5.00E+05 1.00E+05
Initial Cell Number
Cell
Via
bil
ity (
%)
a b
-150-
Figure B.2: Optimization of bulk electroporation of mouse embryonic stem cells using
Bio-Rad Gene Pulser X-Cell system: (a) transfection efficiency and (b) cell viability 24
hours after BE.
0
25
50
75
375 500 625 750
Electrical field strength (V/cm)
Tota
l S
EA
P A
ctivity m
U/
10,0
00 I
nitia
l seedin
g c
ells
0
25
50
75
100
375 500 625 750
Electrical field strength (V/cm)
Cell
Via
bili
ty (
%)
a
b
-151-
Figure B.3: Optimization of nucleofection of mouse ES cells: (a) transfection
efficiency and (b) cell viability 24 hours after nucleofection.
0
25
50
75
100
125
A-13 A-23 A-24 A-30
Program
Tota
l S
EA
P A
ctivity m
U/
10,0
00 I
nitia
l seedin
g c
ells
0
25
50
75
100
A-13 A-23 A-24 A-30
Electrical field strength (V/mm)
Ce
ll V
iab
ility
(%
)
a
b
-152-
0
20
40
60
80
100 150 200
Electrical Field Strength (V/cm)
Rela
tive S
EA
P A
cti
vit
y
(mU
/10,0
00 c
ell
s)
0
20
40
60
80
100
100 150 200
Electrical Field Strength (V/cm)
Cell
Via
bil
ity (
%)
Figure B.4: SEAP transfection of mouse ES cell by micro-nozzle enhanced MSE at
different electrical field strength. Initially, ten thousands of cells were trapped on a 100 x
100 micro-nozzle array, and (a) transfection efficiency and (b) cell viability were
quantified 24 hours after electroporation.
-153-
Figure B.5: Optimization of bulk electroporation (BE) of mouse ES cell colonies using
Bio-Rad Gene Pulser X-Cell system: (a) transfection efficiency and (b) cell viability 24
hours after BE.
0
2
4
6
8
10
375 500 625 750
Electric field strength V/cm
Tota
l S
EA
P A
cti
vity
mU
/
Initia
l 35
,00
0 s
ee
din
g c
ells
0
20
40
60
80
375 500 625 750
Electric field strength V/cm
Cell
Via
bili
ty (
%)
a
b
-154-
Figure B.6: Optimization of nucleofection of mouse ES cell colonies: (a) transfection
efficiency and (b) cell viability 24 hours after nucleofection.
0
5
10
15
20
A13 A23 A24 A30
Program#
To
tal S
EA
P A
cti
vit
y m
U/
Init
ial 3
5,0
00
se
ed
ing
ce
lls
0
15
30
45
60
A13 A23 A24 A30
Program #
Ce
ll V
iab
ilit
y (
%)
a
b
-155-
APPENDIX C: ANALYTICAL SOLUTION OF TRANSMEMBRANE
POTENTIAL FOR A TWO-DIMENTIONAL (2-D) CELL IN BULK
For a 2-D cylindrical cell (three-layer model) in bulk electric field shown in
Figure C.1, the governing equation 0)( =∇⋅∇ φσ can be simplified as
01
)(1
2
2
2
2 =∂
∂+
∂
∂
∂
∂=∇
θ
φφφ
rrr
rr (C.1)
Figure C.1: Schematics of a 2D cylindrical cell in bulk electric field.
-156-
Equation (B-1) can be solved by the method of separation of variable )()( θφ grf=
and usually we take θθ cos)( =g , then 02 =−′+′′ ffrfr . And we have
rBArrf /)( += , or
θθφ cos)/(),( rBArr += (C.2)
So the electric field φ−∇=E is given by
θθ θ sin)/(cos)/( 22rBArBAr +++−= eeE (C.3)
where the relation between two coordinates systems is given as: θθ θ sincos eee −= rx
and θθ θ cossin eee += ry . Generally, we need to solve the electric potential layer by
layer.
(1) Layer of external medium
Since xe E eE 0−= for ∞→r , where 0E is the electric field far away from the cell,
we get 0EAe = , thus θθφ cos)/(),( 0 rBrEr ee += ,
θθ θ sin)/(cos)/( 2
0
2
0 rBErBE eere +++−= eeE .
(2) Layer of inner cytoplasm
iφ is finite at 0=r , then 0=iB , thus θθφ cos),( rAr ii = ,
θθ θ sincos)( iiri AA eeE +−= .
(3) Layer of cell membrane
θθφ cos)/(),( rBrAr mmm += ,
-157-
θθ θ sin)/(cos)/( 22rBArBA mmmmrm +++−= eeE .
Boundary conditions are given as:
nn
m
m
e
e∂
∂=
∂
∂ φσ
φσ (or rmmree eEeE ⋅=⋅ σσ ),
me φφ = on external interface Rr = (C.4)
nn
m
m
i
i∂
∂=
∂
∂ φσ
φσ (or rmmrii eEeE ⋅=⋅ σσ ),
mi φφ = on inner interface dRr −= (C.5)
Thus we have totally 4 unknowns and 4 equations as follows:
=
−−−
−
−−−−−
−
0
0
0)/(
//0
0)/(1)()(
/1/10
0
0
2
22E
RE
B
B
A
A
dR
RR
dRdRdR
RRR
e
e
m
m
i
mmi
emm σ
σσσ
σσσ (C.6)
The solution is:
))(())(()(
)(222
0
2
memimemi
emi
mRdR
ERA
σσσσσσσσ
σσσ
++−−−−
+−= (C.7)
))(())(()(
)()(222
0
22
memimemi
emi
mRdR
EdRRB
σσσσσσσσ
σσσ
++−−−−
−−= (C.8)
))(())(()(
422
0
2
memimemi
em
iRdR
ERA
σσσσσσσσ
σσ
++−−−−
−= (C.9)
))(())(()(
))(())(()(22
22
0
2
memimemi
memimemi
eRdR
RdRERB
σσσσσσσσ
σσσσσσσσ
++−−−−
−+−+−−= (C.10)
The transmembrane potential is given as )()( dRRV mmm −−= φφ and we have
-158-
))(())(()(
)2(cos2
22
22
0
memimemi
emei
mRdR
dRddREV
σσσσσσσσ
σσσσθ
++−−−−
−−= (C.11)
Because im σσ << , and em σσ << , you can treat 0=mσ , thus the transmembrane
potential for a 2D cylindrical cell in the bulk electric field is simplified as
θσσ
σσθ cos2
))((
)2(cos2 022
2
0 RERdR
RddREV
ei
ei
m =−−
−= (C.12)
Thus we complete the deduction of the distribution of transmembrane potential for a
2D cylindrical cell in the bulk electric field.
-159-
APPENDIX D: G-CODE GENERATION FOR FABRICATING MICRO-PORE
ARRAYS BY FEMTOSECOND LASER
Figure D.1: Interface of G-code generation software for femtosecond laser fabrication of
micro-pore arrays. (Programmed by Hae Woon Choi, December 2007)
-160-
APPENDIX E: ELECTROPORATION OF MOUSE EMBRYOID BODIES
Figure E.1: Formation of mouse embryoid bodies (EBs) by handing drop culture. The
initial cell number per drop is ten thousand in (a-c), and twenty thousand in (d-f). Phase
contrast images were taken at day 3 (a, d), day 5 (b, e) and day 8 (c, f).
-161-
Figure E.2: Electroporation of EBs by plasmid RFP at day 8. The initial size of EBs is ~
400 µm in (a, b), and ~ 500 µm in (c, d). RFP stands for red fluorescence protein. (a, c)
Shape of EBs; (b,d) RFP transfection results.
-162-
APPENDIX F: MEMBRANE PERMEABILITY EXPERIMENT
0.1
0.15
0.2
0.25
0.3
0.35
4 6 8 10 12
Diameter of cells (mm)
Tra
ns
me
mb
ran
e p
ote
nti
al
(V) 500V/cm
375V/cm
250V/cm
Figure F.1: Transmembrane potential vs cell size at different external electric field. The
pulse type is exponentially decay, and pulse capacitance is 500 µF. Cell impermanent
dye, propidium iodide (PI), was used in the experiment. Each data point presents that at
least 50% of transfected cells at such size were observed.
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