MICRO-/NANOFLUIDICS AND SINGLE DNA DYNAMICS IN NON-UNIFORM ELECTROKINETIC FLOWS
DISSERTATION
Presented in Partial Fulfillment of the Requirements for The Degree Doctor of Philosophy in the Graduate School of the Ohio State University
By
Shengnian Wang, M.S.
* * * * *
The Ohio State University
2006
Dissertation Committee: Approved by Professor L. James Lee, Adviser
Professor L.-S. Fan ______Adviser Professor Kurt W. Koelling Chemical Engineering Graduate Program
ABSTRACT
The purpose of this study is to fabricate polymer nanodevices and investigate micro-/nanofluidic and DNA dynamics in non-uniform electrokinetic flows.
Single DNA dynamic deformation was firstly studied in cross-slot microfluidic platforms. Three basic flow patterns (i.e., extensional, shear and rotational flows) were generated and polystyrene nanospheres were used to identify the flow characteristics. The conformational evolution of λ-DNA molecules was also investigated and it indicated that the initial conformation of molecules and their residence time in the flow play important roles in the dynamic of DNA stretching.
A new nanonozzle array was developed by Sacrificial Template Imprinting (STI),
which can provide more uniform and controllable DNA stretching compared to the cross-
slot design. A polymer sacrificial template was used to avoid structure damage or defects
during the de-molding process. It was produced by a two-step replication, starting with a
conically shaped nanotip array. Each nanonozzle is 3 µm high with the size at the small
end down to 80 nm. In conjuction with surface modification and silica synthesis on the
surface, the channel size was further reduced and the polymer structure was reinforced.
Further 2D dynamic complexation exhibited a two-stage complexation, extending
gradually from the small end towards the large end.
ii Nanonozzle can provide two important flow patterns: the converging flow and the diverging flow. 2D converging microchannels were used to investigate the migration behavior of rigid nanospheres and flexible DNA molecules. Vortices were observed both inside and at the small end of the converging channel. When nanoparticles have much smaller size than the channels (i.e., the hindered factor is small), for example, in dynamic assembly, non-uniform surface charge led to the formation of new vortex pair. The stagnation region between the double vortex pairs is believed to have the primary complexation. When hindered factor is large, hindered migration was shown in the diverging direction, while nanonozzles were easily clogged in the converging direction for rigid colloid nanospheres. But for flexible λ-DNA molecules, their molecular chain can be stretched to achieve easy pass in the converging direction even though their equilibrium size are much larger than the channel size.
iii
Dedicated to my family
iv
ACKNOWLEDGMENTS
First I would like to express my sincere appreciations to my adviser, Professor L.
James Lee, for his intellectual supports, encouragements and enthusiasm throughout my stay at The Ohio State University. I am in debt to him not only for his invaluable guidance and discussions that made this dissertation possible, but also in various other aspects, such as stimulation of my creativity and support to my career development.
I would also like to acknowledge Professors L.-S. Fan and Kurt Koelling for serving on my dissertation committee and for their invaluable comments and suggestions.
I also wish to thank Dr. Paula Stevenson and Mrs. Stacy Brannan Doepker for proofreading manuscripts I submitted for publication.
I would like to give my special thanks to Shizhong Yu at the Center for
Affordable Nanoengineering of Polymer Biomedical Devices, and Paul Green, Leigh
Edward, Carl Scott at the Chemical and Biomolecular Engineering Department for their endless technical assistance.
My appreciation also goes to all the fellow graduate students and post-doctors in our laboratory, especially to those who have heavily collaborated with me: Xin Hu (on simulation), Yi-Je Juang (on DNA dynamics), Siyi Lai (on CD-ELISA), Yubing Xie (on cell culture), and Changchun Zeng (on silica assembly). Experimental assistances from
v several undergraduate students, Edmond Yang, Mccauley C. Malcolm and Ericka
McKinney, are also greatly appreciated.
Last but not least, I want to thank my parents, sisters and brother for their love, encouragement, constant emotional support throughout this seemingly everlasting endeavor. Special appreciations to my wife, Shijin, for her love, accompany, encouragement, and support through all these years.
vi
VITA
May, 1974 ...... Born - Wuwei, Gansu, P.R. China
September 1993 - July 1997 ...... B.E. Chemical Engineering, Zhejiang University Hangzhou, Zhejiang, P.R. China
September 1997 - June 2000...... M.S. Chemical Engineering, Dalian Institute of Chemical Physics, CAS Dalian, Liaoning, P.R. China
September 2000 – August 2001...... Department Fellowship Chemical and Biomolecular Engineering The Ohio State University Columbus, Ohio, USA
September 2001 – Present...... Graduate Research Associate Chemical and Biomolecular Engineering The Ohio State University Columbus, Ohio, USA
vii
PUBLICATIONS
1. Wang, Shengnian, C. Zeng, S. Lai, Y.-J. Juang, and L. J. Lee, “Polymeric nanonozzle array fabricated by sacrificial template imprinting”, Advanced Materials (2005), 17(9), 1182-1186 2. C. Zeng, Wang, Shengnian, and L. J. Lee, “Dynamic silica assembly for fabrication of nanoscale polymer channels”, Materials Letters (2005) 59(24), 3095-3098 3. Y.-J. Juang, X. Hu, Wang, Shengnian, and L. J. Lee, “Electrokinetic interactions in microscale cross-slot flow”, Applied Physics Letters (2005), 87(24) 244105/1- 244105/3 4. Y.-J. Juang, Wang, Shengnian, X. Hu and L. J. Lee, “Dynamics of single flexible polymer under electrokinetics-induced stagnation flow”, Physical Review Letters (2004), 93(26), 218105/1-218105/4 5. S. Lai, Wang, Shengnian, J. Luo, L. J. Lee, S-T Yang, M.J. Madou. “Design of a CD-like microfluidics platform for enzyme-linked immunosorbent assays”, Analytical Chemistry (2004), 76(7) 1832-1837
FIELDS OF STUDY
Major Field: Chemical Engineering Studies in Micro-/nanofluidics, BioMEMS and Polymer Micro-/nanofabrication
viii
TABLE OF CONTENTS
Page
Abstract...... ii
Dedication...... iv
Acknowledgments...... v
Vita...... vii
List of Tables ...... xv
List of Figures...... xvi
CHAPTERS:
CHAPTER 1...... 1
1.1 BACKGROUND...... 1
1.2 OBJECTIVES OF STUDY...... 3
1.3 OUTLINE...... 4
CHAPTER 2...... 6
2.1 INTRODUCTION...... 6
2.2 FABRICATION OF NANOCHCANNELS ...... 7
2.2.1 Nanoimprint lithography and its derivatives...... 9
2.2.1.1 Nanoimprint lithography...... 9
ix 2.2.1.2 Bonding techniques...... 13
2.2.2 Soft lithography...... 14
2.2.3 Sacrificial layer removal technology ...... 16
2.2.3.1 Metal sacrificial layer...... 17
2.2.3.2 Si-based sacrificial materials...... 18
2.2.3.3 Polymer sacrificial materials...... 19
2.3 NANOFLUIDICS...... 22
2.3.1 Introduction...... 22
2.3.2 Electrokinetics...... 22
2.3.3 Molecular transport in nanochannels...... 27
2.3.3.1 Molecules transportation in nanotubule membrane ...... 28
2.3.3.2 Biomolecules transportation in 1D nanochannels on Si membrane...... 29
2.3.3.3 Molecule sensing and sorting in single nanopore or nanochannel...... 31
2.4 DNA MANIPULATION TECHNIQUES ...... 34
2.4.1 Introduction...... 34
2.4.2 Hydrodynamic DNA stretching under different flows ...... 35
2.4.3 DNA stretching study using nanopost and nanoscopic slit...... 38
CHAPTER 3...... 42
3.1 INTRODUCTION...... 42
3.2 EXPERIMENTAL...... 43
3.2.1 Materials ...... 43
3.2.2 Fabrication ...... 44
3.2.3 Sample preparation...... 44
x 3.2.4 Characterization ...... 45
3.2.5 Experimental setup...... 46
3.3 SINGLE CROSS-SLOT FLOWS VISUALIZATION ...... 47
3.3.1 Extensional flow...... 47
3.3.1.1 Velocity field imaging and analysis...... 47
3.3.1.2 Flow-induced molecular dynamics of DNA ...... 49
3.3.2 Shear and recirculation flows...... 55
3.3.2.1 Shear flow and recirculation flow generation ...... 55
3.3.2.2 Electrokinetic interaction analysis ...... 56
3.4 FIVE CROSS-SLOT ASSISTED MICROFLUIDIC PLATFORM ...... 59
3.4.1 Extensional flow and shear flow...... 60
3.4.2 Rotational flow...... 62
3.5 SUMMARY...... 63
CHAPTER 4...... 86
4.1 INTRODUCTION...... 86
4.2 SACRIFICIAL TEMPLATE IMPRINT...... 88
4.2.1 Materials ...... 88
4.2.2 Process description...... 88
4.2.3 Fabrication of optic-fiber nanotip ...... 89
4.2.4 Fabrication of sacrificial template ...... 92
4.2.5 Fabrication of nanonozzle array...... 94
4.3 PLANIZATION OF APERTURE PART ...... 96
4.3.1 Wet planization...... 97
xi 4.3.2 Plasma planization...... 98
4.4 DERIVATION OF SACRIFICIAL TEMPLATE IMPRINTING ...... 100
4.5 DYNAMIC SILICA ASSEMBLY IN NANONOZZLES ...... 101
4.5.1 Introduction...... 101
4.5.2 Experimental...... 102
4.5.2.1 Materials...... 102
4.5.2.2 Surface modification ...... 103
4.5.2.3 Dynamic assembly ...... 104
4.5.3 Results and discussions...... 104
4.5.3.1 Dynamic assembly procedure ...... 104
4.5.3.2 Dynamic assembly in STI nanonozzle array...... 105
4.5.3.3 Control of dynamic assembly procedure...... 107
4.5.3.4 Mechanical property testing...... 110
4.6 SUMMARY...... 111
CHAPTER 5...... 130
5.1 INTRODUCTION...... 130
5.2 2D MICROFLUIDIC STUDY ...... 131
5.2.1 Experimental...... 131
5.2.1.1 Materials...... 131
5.2.1.2 Fabrication 2D microfluidic channel...... 131
5.2.1.3 Charecterization ...... 132
5.2.1.4 Flow Imaging ...... 133
5.2.2 Results...... 133
xii 5.2.2.1 Flow profile inside the converging channels...... 133
5.2.2.2 Flow profile at the outlet ...... 136
5.2.2.3 Flow profile in the diverging and straight channels...... 137
5.2.3 Discussions and force analysis...... 138
5.2.3.1 Electrokinetic forces...... 138
5.2.3.2 Viscous drag and hydrodynamic interactions ...... 142
5.2.3.3 Dielectrophoresis(DEP) ...... 145
5.3 2D DYNAMIC COMPLEXATION ...... 149
5.3.1 Experimental...... 149
5.3.1.1 Materials...... 149
5.3.1.2 Fabrication 2D microfluidic channel...... 150
5.3.1.3 Experimental setup and imaging...... 150
5.3.1.4 Polyelectrolyte modification ...... 151
5.3.2 Results and discussions...... 151
5.3.2.1 Polyelectrolyte distribution...... 151
5.3.2.2 2D dynamic assembly process ...... 152
5.3.2.3 Dynamic complexation mechanism ...... 154
5.3.2.4 Effect of feed pattern on complexation...... 157
5.4 TRANSPORT IN 3D NANONOZZLE ARRAY ...... 160
5.4.1 Experimental...... 161
5.4.1.1 Materials...... 161
5.4.1.2 Fabrication of 2D microfluidic channel ...... 162
5.4.1.3 3D experimental setup...... 162
xiii 5.4.1.4 Characterization ...... 163
5.4.2 Transport of rigid nanospheres ...... 163
5.4.3 Transport of semi-flexible DNA...... 167
5.5 SUMMARY...... 170
CHAPTER 6...... 202
6.1 CONCLUSIONS ...... 202
6.2 RECOMMENDATIONS...... 204
6.2.1 Electrokinetic-driven flows...... 204
6.2.2 Dynamic complexation...... 206
6.2.3 Nanotip and nanonozzle drug/gene delivery...... 209
6.2.3.1 Nanonozzle array based drug/gene delivery ...... 209
6.2.3.2 Nanotip array based drug/gene delivery...... 211
6.2.4 Flow assisted organization of nanostructures ...... 212
6.2.4.1 Nanostructures with multiple functions ...... 213
6.2.4.2 Drug/DNA loading in naocapsules ...... 214
BIBLIOGRAPHY...... 221
APPENDIX A...... 229
APPENDIX B ...... 236
APPENDIX C...... 241
APPENDIX D...... 247
xiv
LIST OF TABLES
Table Page
Table 5.1 Zeta potential and mobility of Fluoresbrite Yellow Green (YG) polystyrene
micro-and nanospheres and λ-DNA. The solid content of micro/nanospheres is
0.00265%, 1/1000 from the stock suspensions unless is specified mentioned and the
concentration of λ-DNA is about 0.03 µg/ml...... 172
xv
LIST OF FIGURES
Figure Page
Figure 3.1 Optical image of single cross-slot device (a) and the schematic of the cross-
slot dimensions and the arrangement of electrodes (b) (drawn not in scale). The sale
bar represents 1 cm...... 65
Figure 3.2 Optical image of five cross-slots device (a) and the schematic of the five cross-
slots dimensions and the arrangement of electrodes (b) (drawn in scale). The sale bar
represents 500 µm...... 66
Figure 3.3 Experimental setup: the schematic drawing (a) and photography (b)...... 67
Figure 3.4 Schematic of surface arrangements for the generation of different flow
patterns by electrokinetics in single cross-slot: pure elongational flow (a), mixed
shear flow (b), and pure rotational flow (c)...... 68
Figure 3.5 An elongational flow in a single cross-slot: the streamlines by compounding
image of fluorescence microspheres trace driven by electrokinetics (a), by simulation
(b) and velocity vector field of experimental data (c) and simulation (d)...... 69
Figure 3.6 Comparison of the electric lines, the streamlines of a pressure-driven flow, and
the streamlines of a pure elongational flow...... 70
xvi Figure 3.7 Velocity distributions of DNA molecules at various locations in the spanwise
(x) and vertical (z) directions. z = 0 µm is at the bottom of the channel...... 71
Figure 3.8 The plot of x component of the velocity of DNA mass center versus x position
with the stagnation point as the origin. Data are taken their absolute value so that
they all fall in the first quarter of the cross-slot, with different symbols representing
data from different quarters. The extension rate in the pure elongational flow, ε& , is
given by the slope of the fitting curve, which is equal to 0.78 (1/s) in the
experimental conditions...... 72
Figure 3.9 Experiment and simulation of the movements of two DNAs in the intersection
region with similar initial conformation but different residence time (a), with
different initial conformation and residence time (b)...... 73
Figure 3.10 Different initial configurations of λ-DNA molecules in an elongational flow
driven by electrokinetics within single cross-slot: super-coiled (a), curled (b), half-
dumbbell (c and d), kinked (e), dumbbell (f and g) and stretched (h). The scale bar
represents 5 µm...... 75
Figure 3.11 Flow pattern evolution from strong electrokinetic interactions for different
particle charge density or λ (= µEP / µEO ) : λ is equal to (a) 0, (b) -0.2, (c) -1.0 from
simulation and (d) -1.0 from experiment (two asymptotes are outlined near the
intersect area for clarity). Scale bar represents 200 µm...... 76
Figure 3.12 Flow pattern evolution from electrokinetic interaction for different wall
surface zeta potential for positively charged arm pair and the negatively charged arm
EO EO pair or χ (= µ− / µ+ ) : χ is equal to (a) -2, (b) -5 with λ = −0.05 , and is fixed. (c)
xvii -1 from simulation and (d) -1 from experiment withλ = −1, and is fixed. The scale
bar represents 200 µm...... 78
Figure 3.13 Flow pattern generation of both elongational flow (in the center cross) and
mixed shear flow (in the side crosses) in a five cross-slot design: the design and
arrangement of electrodes (a), experimental result of the compounded pathlines of PS
microspheres, representing an elongational flow in the center cross (b), and mixed
shear flows in the side crosses (c). The scale bar represents 200 µm...... 80
Figure 3.14 Experimental and simulation results of the movements of DNA molecules in
the five cross-slot microfluidic platform in an elongational flow (a), in a mixed-shear
flow (b). Two DNA molecules experience different residence time...... 82
Figure 3.15 Flow pattern generation of rotational flow (in the center cross) in a five cross-
slot design: the design and arrangement of electrodes (a), experimental result of the
compounded pathlines of PS microspheres, representing an rotational flow in the
center cross (b), and microscopy image of λ-DNA (c). The scale bars in (b) and (c)
represents 200 µm, and 20 µm, respectively...... 83
Figure 3.16 Dimensionless vortex size versus the ratio of EP mobilty to EO mobility
λ (= µEP / µEO ) and two rotational flow patterns forλ = −0.2 and - 2 ...... 85
Figure 4.1 Schematic of the Sacrificial Template Imprinting (STI) process for fabricating
nanonozzle arrays with uniform, conically shaped nanochannels...... 112
Figure 4.2 SEM images of nanotip formation study in BOE (NH4F/HF=7:1) at room
temperature: (a) after 10 min; (b) after 20 min; (c) after 30 min and (d) the
xviii temperature effect on the nanotip geometry, resulting from the different etching rates
of core and cladding materials...... 113
Figure 4.3 SEM images of: (A) a nanotip array from an optic fiber bundle; (B) a nanotip
array cast by poly(vinyl alcohol) aqueous solution, 10 wt%; (C) a nanotip array cast
by chitosan 1M acetic acid solution, 1.0 wt%; (D) a nanotip array cast by
poly(methylacrylic acid, Sodium) aqueous solution, 10 wt%; (E) a nanotip array cast
by poly(methyl methacrylate) toluene solution, 10 wt%; and (F) a nanotip array cast
by poly(methyl methacrylate) acetic acid solution, 10 wt% ...... 114
Figure 4.4 The spin speed versus thickness curves for PMMA/toluene solutions with
different solid contents...... 116
Figure 4.5 SEM images of poly(methyl methacrylate, PMMA) nanonozzle arrays formed
with (A) 15 wt% and (B) 20 wt% PMMA solution in toluene...... 117
Figure 4.6 Comparison of size variation of nanonozzles and original optic fibers. The
solid circle represents for nanonozzles and hollow circle for optic fibers...... 118
Figure 4.7 The Schematic of (a) and SEM images of dynamic planization using
hydroxyethyl methacrylate, HEMA: the landscape-scale view (b) and an individual
channel (c)...... 119
Figure 4.8 The Schematic and SEM images after dry planization by oxygen plasma
etching: a short period removal with a thin PMMA layer (a-b) and a long period
removal with a thick PMMA layer (c-d)...... 120
Figure 4.9 The derivatives of sacrificial template imprinting I—obtain negative pattern
with the same material: schematic of the process (a) and SEM images of PDMS male
xix mold (b), PMMA female replica (c), PVA sacrificial mold (d) and PDMS female
mold (e)...... 121
Figure 4.10 The derivatives of sacrificial template imprinting II—obtain same pattern
starting with EBL patterns: schematic of the process (a) and SEM images of PMMA
photoresist nanochannels on Si substrate (b), PDMS male mold (c), SU-8
nanochannels (d), PVA sacrificial template and (e) PMMA nanochannels (f)...... 122
Figure 4.11 Strategies to introduce polyallyamine hydrochloride (PAH) on PMMA and
PET surface: through amidation reaction under basic conditions (a) and hydrolyzing
the ester bond under strong base solution (b)...... 124
Figure 4.12 Schematic of dynamic assembly of silica assisted by electrokinetics...... 125
Figure 4.13 Reaction mechanism of silica formation reaction: (a) the hydrolysis of
Tetraethylorthosilicate (TEOS) precursor to form Si(OH)4 at pH 2-7 and the
condensation of the deprotonated and neutral species to form silica...... 126
Figure 4.14 Scanning electron microscopy (SEM) images of polymer nanonozzle array
before (a) & (c) and after dynamic assembly (b) & (d): (a) & (b) on the large end, (c)
& (d) on the small end...... 127
Figure 4.15 Comparison of hardness of PET and PET/silica polymer layers...... 129
Figure 5.1 Images of 2D converging channels. The width of the small end is 20 µm in (a)
and 5 µm in (b)...... 173
Figure 5.2 The flow characteristics inside the converging channel by electrokinetics-
induced flow (a), pressure-driven flow (b), and electrokinetics-induced flow in a
straight channel (c)...... 174
xx Figure 5.3 The rheology data of PS suspensions of different dilute folds: (a) 700 nm,
diluted 1/100~1/1000 of the stock concentration and (b) 40nm, diluted
1/100~1/10000 of the stock concentration...... 175
Figure 5.4 The flow characterization outside the converging channels in the
electrokinetics-induced flow (a, b): 40 nm PS nanospheres (a) and λ-DNA molecules
(b), outside the straight channel with the same size of the small end of the converging
channel (c), and in the pressure-driven flow (d, e, f): Re=1.0 (d), Re= 10 (e) and
Re=250 (f). The arrows indicate the migration direction and the scale bars represent
20 µm...... 176
Figure 5.5 The flow characteristics of electrokinetic-induced flows outside the small end
of the diverging channels (a, b): E= 60 V/cm (a) and 120 V/cm (b), and pressure-
driven flows outside the small end of the diverging channels (c, d): Re= 100 (c) and
Re= 500 (d), and straight channels (e, f): E= 60 V/cm (e) and 120 V/cm (f). The scale
bars present 20 µm...... 177
Figure 5.6 Zeta potential of various polymer substrates...... 179
Figure 5.7 The electric field strength profile in a 2D converging channel and zone
division based on the dominating forces...... 180
Figure 5.8 The experimental data of lateral velocity profile measured along the centerline
of a 2D converging channel. The solid lines are the simulated curves for both
converging and straight channels with the channel size the same as the two ends of
the 2D converging channel. The small end of the converging channel is located at
x/L=0 and the large end is at x/L=-1.0...... 181
xxi Figure 5.9 Different velocity profiles including a bi-directional flow in a straight channel
(a) and the vortices in a converging channel (b) after the superposition of the
electrokinetic-induced flow (plug flow) and the induced pressure-driven flow
(parabolic velocity profile)...... 182
Figure 5.10 The contour of E2 in the converging channel: (a) the whole image and (b) the
local image near the small end and (c) the ratio of dielectrophoretic mobility to
electrokinetic mobility along the centerline of PDMS, untreated and PAH modified
PMMA 2D converging channels. Particles used here are 40 nm PS nanospheres. The
small end of the converging channel is located at x/L=0 and the large end is at x/L=-
1.0...... 183
Figure 5.11 The growth curves (a) and the concentration profile along the converging
channel surface (b) in dynamic complexation process with the solid content of 10-5 in
the suspension. The fluorescence intensity near the channel surface indicates the
amount of nanoparticles deposited. In (a), letters (a-d) label different locations on the
converging channel surface, starting from “a” on the small end, and in (b), the small
end of the converging channel is located at x/L=0 and the large end is at x/L=1.0. 185
Figure 5.12 Snapshots of dynamic complexation using 40 nm fluorescence-labeled PS
nanospheres (10-5 solid content) in 2D PAH-modified converging channel by
continuous feeding. The elapse time has a unit of second. The scale bar represents
100 µm...... 186
Figure 5.13 The extent of two-stage dynamic complexation. θ is equal to the instant
fluorescence intensity/equilibrium fluorescence intensity. The slopes, k1 and k2
xxii represent the complexation rates for primary and secondary complexation,
respectively. Here only curves “a” and “c” are shown for clear image purpose...... 187
Figure 5.14 The flow characterization of the startup of dynamic complexation with
converging channel surface initially positively charged (a). As comparison, flow
patterns at the outlet (b) and inside (c) a negatively charged converging channel are
also shown. The arrows indicate the migration direction and the scale bars represent
20 µm...... 188
Figure 5.15 The schematic of dynamic complexation mechanism: (a) startup with the
surface uniformly positively charged, (b) the small end becomes less positively
charged, (c) vortex pair appear with the small end becoming negatively charged, (d)
two zones are formed with the negatively charged zone expands and the positively
charged zone retreats, and (e) completely negatively charged zone. The formation of
vortex pair induced by the zeta potential gradient is explained by the schematic in (f)
and exhibited by the streak image of dynamic complexation experiment (g)...... 189
Figure 5.16 The growth curves (a) and the concentration profile along the converging
channel surface (b) in dynamic complexation process by single dosage feeding with
the solid content of 10-4 in the suspension. The fluorescence intensity indicates the
amount of nanoparticles deposited. In (a), letters (a-d) label different location on the
converging channel surface, starting from “a” on the small end and in (b), the small
end of the converging channel is located at x/L=0 and the large end is at x/L=1.0. 191
Figure 5.17 The growth curves (a) and the concentration profile along the converging
channel surface (b) in dynamic complexation process by multiple dosage feeding
xxiii with the solid content of 10-4 in the suspension. The fluorescence intensity indicates
the amount of nanoparticles deposited. In (a), letters (a-f) label different location on
the converging channel surface, starting from “a” on the small end and in (b), the
small end of the converging channel is located at x/L=0 and the large end is at
x/L=1.0...... 192
Figure 5.18 Snapshots of dynamic complexation using 40 nm fluorescence-labeled PS
nanospheres in 2D PAH-modified converging channel by multiple dosage feeding.
The elapse time has a unit of second. The scale bar represents 100 µm...... 193
Figure 5.19 Experimental setup for transport study in 3D nanonozzle arrays: schematic
(a) and experimental setup (b)...... 194
Figure 5.20 Plots of electrokinetically hindered transport of 40 nm PS nanospheres (a) in
a nanonozzle array with the small end diameter of 200 nm: circular symbols for
nanospheres transport from the small end to the large end; triangle symbols for
nanospheres transport in the opposite direction; (b) in a nanonozzle array with the
small end diameter of 200 nm (reverse triangle), 200 nm track-etched membrane
(square) and 1 mm track-etched membrane (circle)...... 195
Figure 5.21 Plot of electrokinetic hindered transport of various sizes of PS nanospheres in
nanonozzle arrays with the small end diameter of 200 nm (a) and the hindered factors
of different size PS nanospheres (b). The line is the prediction of Renkin’s equation.
...... 197
Figure 5.22 Microscopic images of 5-µm converging channel: before experiment (a),
after hindered migration study in the converging direction (b) and in the diverging
xxiv direction (c, d). The hindered factors in (b, c, and d) are equal to 0.20, 0.20, and 0.40,
respectively. The scale bar represents 20 µm...... 198
Figure 5.23 Experimental setup (a) and diagram of DNA migration through channel array
under electric field (b)...... 199
Figure 5.24 Snapshots of λ-DNA molecules passing through nanonozzle array in the
converging (a) and diverging (b) directions, respectively...... 200
Figure 5.25 Different stretching extents of λ-DNA in the 2D converging channel (a) and
diverging channel (b). The solid dots in the cartoons represent the positions of DNA
molecules in each image, but not drawn in scale for clear purpose. The scale bar
presents 5 µm...... 201
Figure 6.1 Various converging/diverging geometries and their combinations (a-c);
schematic of polymer conformation transition in converging/diverging geometry (d),
Electric field profile in a series of converging channels (e)...... 216
Figure 6.2 Schematic of dynamic DNA complexation or hybridization (a), DNA
conjugates with diverse functional groups or nanoparticles (b), replica of helix
structure (c)...... 217
Figure 6.3 Schematic of two strategies of polymer nanonozzle array cell patch for
gene/drug delivery (a) for drugs and small genes; (b) for large genes...... 218
Figure 6.4 Schematic of procedure for nanotip array drug/genes delivery (a) and the
compositions of nanotips for the delivery of drugs (b) and genes (c)...... 219
xxv Figure 6.5: Schematic of flow assisted organization of nanostructures: (a) multiple
functional nanostructure, (b) drug and (c) gene loading in nanocapsules. The lines
and blocks with different colors represent different materials...... 220
Figure A.1 Snapshots of two typical motions for T2 DNA in 1 wt% PEG solution inside
straight microchannel: (a) “worm-like” motion and (b) “hooked and sliding” motion.
...... 233
Figure A.2 Images of flowfront in a fountain flow with λ-DNA solution (0.3 µg/ml). . 235
Figure B.1 The series of snapshots of charged nanoparticles’ motion in the converging
channel with an air bubble trapped at the small end. (a-f) is for an individual
nanoparticle and (g-l) is for the second one. The second particle enters the image
domain in (d). The scale bar is 100 µm...... 238
Figure B.2 Cells trapping and electroporation study: (a) cells are trapped at the small end
of converging channel and (b) after the electroporation. The scale bar is 20 µm.... 240
Figure C.1 (a) Schematic of 5-step flow sequencing CD (reservoir 1 − antigen/sample,
2,4 − washing, 3 −2nd antibody, 5 − substrate, 6 − detection region, and 7 − waste),
(b) a CNC-machined CD having four microfluidic platforms with the same design as
shown in (a), and (c) the burst frequency of reagents from each reservoir...... 245
Figure C.2 (a) The signal variation checking along the whole microchannel and (b) the
calibration curves of rat IgG conducted on the microchip...... 246
xxvi
CHAPTER 1
INTRODUCTION
1.1 BACKGROUND
Miniaturization of silicon devices in microelectronics was one of the most significant enabling technologies of the twentieth century. In the past two decades, this technology has been extended to microelectromechanical systems (MEMS), microfluidics and applications in the biological area (i.e., BioMEMS). Miniaturization provides opportunities to massively parallel analysis with smaller quantities of materials as well as exploration of new effects and better performance.
With the emerging new imaging techniques, an exciting area, microscopic imaging of single polymer or DNA dynamics has attracted a great deal of attention from researchers in the past decade. Such study in submicron or even nanochannels could be more valuable as many new surface forces could be used to manipulate single molecules to better understand the molecular dynamics of polymeric fluids and to provide tools for
applications such as single molecular sensing. However, only limited work based on
1 hydrodynamic flows in quite large channels has been conducted so far. These techniques
ften resulted in highly non-uniform stretching of molecules in complicated flow cells.
Nanochannels with identical dimensions and large effective area may provide a completely new tool to investigate fundamental fluid transport and molecule dynamics
(i.e., nanofluidics) as well as the impetus for applications in analytical chemistry, biochemistry, biomedical diagnostics, pharmacy and health care. However, little work has been in these areas due to the lack of suitable and robust nanofluidic platforms.
Typically, nanoscale channels are fabricated by “top-down” methods. Most of
these methods involve the use of electron beam lithography (EBL) or focused ion beam milling (FIB) techniques to create open channels first. These open channels have to be sealed subsequently by an appropriate bonding procedure in order to handle fluids.
Compared to FIB, EBL followed by directional etching can provide a better surface
quality and dimensional control. However, both processes are slow and expensive,
especially when pattern density is high. With EBL prepared templates, the nanoimprint lithography (NIL) is often used to replicate nanoscale features onto silicon. Another alternative approach is using a sacrificial layer. This sacrificial-layer-removal approach can be applied to create silicon or metal nanochannels. Materials used as sacrificial layers include silicon nitride, silicon oxide, metals and some thermally degradable polymers.
The removal of the sacrificial layer in long nanochannels is very time-consuming if not impossible.
Polymers are more favorable than silicon in biological applications because many polymers can provide good biocompatibility and some are biodegradable. Moreover, polymer devices are inexpensive and disposable, which is highly desirable for many
2 biological applications that require cost-effective devices (e.g., biosensors for medical and food testing and point-of-care clinical diagnostics). The use of polymeric materials may also increase the opportunities for exploiting various functional channel surfaces for broader biological applications. However, nanoscale polymer processing is not as well developed as silicon processing, and suitable methods are not available to produce polymer nanochannels with high aspect ratios and high channel density. De-molding is a major challenge to fabricate these nanofeatures on polymer. Structure damage or defects is inevitable in de-molding when dealing with fragile nanoscale parts with a large contact area, a high density, and high aspect ratios. Mold release agents may be helpful to assist de-molding for nanofeatures with a low aspect ratio (typically less than one). But its effectiveness is quite dubious when applying to high-aspect-ratio features. Besides, residual mold release agents may introduce potential contamination for biological applications.
Therefore, there is an urgent need to develop new manufacturing methods that can achieve well-defined polymeric micro/nanoscale structures that present high-density patterns with low cost and high throughput, and to study the fluidic and transport phenomena in those nanochannels.
1.2 OBJECTIVES OF STUDY
The overall goal of this study is to develop polymer-based micro-/nanofluidic platforms, which can be used for many biological applications, e.g., immunoisolation of cells, DNA and protein sensing and manipulation, and drug/gene delivery, etc. This study focuses on the development of new nanofluidic platforms and the study of flow-based
3 manipulation of colloids and biomolecules under non-uniform electric field. Specially,
the objectives are:
1. To develop a low-cost and robust method to manufacture desirable nanofluidic
platforms. To make it affordable, the use of expensive equipments or clean room facility should be avoided. In addition, it must overcome current technique challenges mentioned above for nanofeature fabrication with high-aspect-ratios.
2. To manipulate and control DNA deformation in electrokinetic-induced flows.
Standard flow patterns, like elongational, shear and rotation flows will be generated by electrokinetic forces and applied in the study of DNA dynamic.
3. To study the transport phenomena of rigid colloids and flexible DNA molecules in the electric-field-enhanced nanofluidics and to design nanofluidic platforms for various biomedical applications.
1.3 OUTLINE
A general introduction is given in Chapter 1, addressing the background and motivations of this study, together with the objectives.
In Chapter 2, a detailed literature review is presented on the history and prospective of miniaturization of fluidic platforms, progresses in “top down” and “bottom up” approaches for nanofabrication, electrokinetic-induced flows, molecular transport and sensing in nanochannels, and DNA manipulation and deformation.
In Chapter 3, the conformational evolution of single flexible molecules (e.g. λ-
DNA) is introduced in elongational, shear and rotational flows. A single cross-slot flow is
first applied to generate the three basic flows by using different surface charge
4 arrangements on flow arm surfaces and electric bias configurations in liquid storage
reservoirs. The interactions of electrophoresis and electroosmosis are also explored.
Finally a five cross-slots microfluidic networks is designed, which could perform electrokinetics-induced flows largely independent of the charge density and polarity of analytes in the fluid.
Chapter 4 covers the fabrication of polymer nanonozzle arrays. The Sacrificial
Template Imprinting (STI) approach is first introduced to massively produce polymer
nanonozzle arrays with high convergence ratios. A novel dynamic assembly approach is
then introduced to further regulate the nanochannel size and reinforce the polymer
structure. Surface planization techniques and other derivatives to extend this sacrificial
template imprint are also addressed.
Chapter 5 presents the electric field enhanced transport in converging/diverging
channels. To investigate the mechanism behind the dynamic assembly process, 2D
converging/diverging flows and dynamic complexation are studied to simulate the
dynamic assembly process. In the second part of this chapter, particle transport through
the 3D nanonozzle array is investigated in both flow directions with electrolyte
suspensions.
In Chapter 6, the conducted research work and results are summarized and
recommendations are given on future research directions.
5 CHAPTER 2
LITERATURE REVIEW
2.1 INTRODUCTION
Miniaturization of silicon devices in microelectronics was one of the most significant enabling technologies of the twentieth century. In the past two decades, this technology has been extended to machining mechanical microdevices, which were later
named as microelectromechanical systems (MEMS). With the development of
microsensors, micropumps, and microvalves, the concept of microfluidics is emerged to
deal with transport phenomena and fluid-based devices at the micrometer scale.
Microfluidic chips has rapidly attracted the interest from analytical chemists, biochemists
and chemical engineers since Manz et al (1990) indicated that they can be applied to the
life sciences and chemistry. Miniaturization provides opportunities to massively parallel
analysis with smaller quantities of materials as well as to explore new effects and better
performance. Just like computers release people from tedious calculating labors,
microfluidic chips provide a high screening throughput of chemical reactions or drugs. In
addition, its large surface-to-volume ratio might enhance the efficiency of chemical
reactions by eliminating transport difficulties commonly met in the macroscopic world
6 (e.g., increasing the sensitivity of biosensors or assays). These advantages are extremely
important for chemistry or biochemistry production, biomedical diagnostics,
pharmaceutical and health care applications.
With the success of novel micro-/nanofabrication techniques, remarkable
progresses were made in experimental tools to downsize the well-defined channels to the
nanoscale. Even though only simple geometries such as slim slit or cylindrical pores are
available, it rapidly attracts great interests from researchers to open a new exciting area
called nanofluidics. This new fluidic study at the nanoscale might be beneficial for
understanding the fundamental fluid transport and molecule manners as well as for
applications such as single molecular sensing, targeted biorecognition, and cell and gene
therapy.
2.2 FABRICATION OF NANOCHCANNELS
Current “top-down” methods of fabricating nanoscale channels include two major
approaches: one is using well-developed techniques (e.g., electron beam lithography,
EBL or focused ion beam milling, FIB) to create high resolution open channels first,
followed by an appropriate bonding procedure to seal the channels for the handle of
fluids; while the other approach is to create channels with a sacrificial layer and remove
the sacrificial material by either wet etching or thermal degradation.
Several well-established methods for fabricating nanopatterns include lithography
with photons (X-ray or EUV), particles (electron or ion beams) and scanning probes (i.e.
AFM & STM). All these photolithographic methods share the same two-step operation:
7 introduce a latent image by exposure to the electromagnetic radiation (UV, DUV, EUV,
or X-ray) at first and then develop the image into relief structures through etching.
Different resolutions can be achieved by choosing different radiation methods. Among all these techniques, both e-beam lithography and x-ray lithography are slow, expensive and high-energy techniques, but offer high resolution (<10 nm). Compared to FIB, EBL followed by directional etching can provide a better surface quality and dimensional control. Scanning probes lithography (SPL), for instance, Atomic Force Microscope
(AFM) and Scanning Tunneling Microscope (STM) are relatively cheap but provide only low resolution (~10-50 nm), low density and low fabrication speed. UV photolithography has a resolution limitation (>100 nm) due to the diffraction limit. Many unsolved issues have limited applications of each method. All of them also suffer the same limitation of low throughput, especially when the pattern density is high.
Materials of the substrate used in all of these methods are mostly limited to Si and
SiO2. With new MEMS applications (e.g., using highly corrosive chemicals), other inertial or sturdy materials such as stainless steel or ceramics are desired. For BioMEMS applications (e.g., medical diagnostics, genetic sequencing, chemistry production, drug discovery, and proteomics), plastic based single-use disposable devices are highly desirable to clear a formidable price tag for most potential users. Many alternative and potential low-cost methods have been investigated, such as nanoimprint lithography
(NIL), soft lithography, and dip-pen lithography. These techniques are called “alternative techniques”, which are serious competitors for silicon micromachining due to their various material options and wide applications.
8 2.2.1 Nanoimprint lithography and its derivatives
2.2.1.1 Nanoimprint lithography
Nanoimprint lithography (NIL) is one of the promising technologies for high throughput nanoscale patterning. This technology was first invented by the Chou’s group to massively pattern features onto the Si substrate, and later extended to several polymer substrates. Nanoimprint lithography includes two steps: imprinting and pattern transfer.
The imprinting step, in principle, is the same as hot embossing but downsizing the feature size is reduced to nanometers. After imprinting, the residual material in the compression area is removed by reactive ion etching (RIE). The resolution of NIL depends not only on the rigid mould (its mechanical strength and pattern resolution), but also the physical properties of polymers. Silicon dioxide moulds were employed to fabricate dots and lines on the PMMA film (50-200 nm in thickness, Chou, S. Y., et al, 1995 and 1996). A resolution of 25 nm for holes and 70 nm for lines were achieved with the imprint temperature of 200°C and an optimum pressure about 13 MPa The highest resolution achieved by NIL is 6 nm in diameter and 60 nm in depth for holes, and 30 nm in diameter and 35 nm in height for pillars. The different resolutions for holes and pillars are due to the alignment accuracy. The pillar patterns could be easier destroyed than the hole patterns during demoulding. Other polymers such as cellulose acetate and standard optical resist S1805 were also employed in NIL at a lower imprint temperature.
Uniformity is as important as high resolution for NIL but it seems difficult to achieve desirable results for both at the same time. The flow behavior, the shrinkage and etching resistance can be different in different regions. Just like the parts produced by hot
9 embossing, the aspect ratio of NIL features is usually low. Slightly higher aspect ratio can
be obtained by multilayer (bilayer or trilayer) pattern. Using the same technique, NIL can
transfer patterns to a non-planar surface. In order to manufacture NIL nanostructures, the
continuous process can be applied, using either a bend mould or a smooth roller.
However, nanoimprinting lithography has plenty of challenges to be solved for
further progress. The limitations include: handling of complex patterns with varied
feature density; patterning over topographies and pattern alignment; new imprinting materials with desired properties. Many academic researchers and companies joined in research to improve NIL so it can become an important competitor among many next generation lithography technologies (NGL).
One of the most important progresses is the step-and-flash lithography (SFL), which was developed by Wilson’s group (Colburn, M., et al, 1999). Unlike the conventional NIL, a quartz template with nanoscale features and a photopolymerizable organosilicon solution were applied. A drop of low viscous solution was loaded between the gap of the template and the substrate. The fluid spread out and filled features under the capillary action. Then the gap was closed by pressing the template against the
substrate and UV light was irradiated for certain time to solidify the resist. Since the
process is operated at room temperature and at low pressure, it is easier than NIL to
pattern the whole wafer area by repeat same steps.
Replicate fidelity and defect control was a major problem for nanoimprint lithography. The deformation problem comes from the residual stresses generated during processing.
10 The first type of deformation results from the materials chosen for the mould and the substrate. Several materials have been tested, including silicon, silicon oxide, silicon
nitride and sapphire. The thermal expansion coefficient difference between the mould and
the substrate might result in pattern distortions or a stress build-up. Using Si as the
material for both the mould and the template is still the best pair so far, though not
perfect. Since nanoscale features penetrate the polymer layer, the adhesion of the polymer
material on the mould results in the generation of defects. Some means are used to
minimize the sticking problems, including plasma deposition of fluropolymers
(Jaszewski, R. W. et al, 1999) or by using a monolayer of perfluorosilane surfactant
molecules (e.g., 1H, 1H, 2H, 2H-perfluorodecyl-trichlorosilane, FDTS, Beck, M. et al,
2002). Through avoiding the liquid wetting problem to the nanoscale features, a better
imprint quality has been obtained. But the durability and efficiency of the surfactant
coating is still unclear. Some researchers attempted to use fluoropolymer thin film
directly as the template to alleviate this issue (Khang, D.-Y., et al, 2004a and 2004b).
Another type of deformation comes from the stability of the polymer after
patterning. The polymer has to suffer the mould separation and subsequent pattern
transfer steps. For the time being, there is no efficient ways to solve such a problem but
the problem can be alleviated by optimizing the processing conditions. Two major issues limit the possible solutions to this seemingly contradictory problem. Firstly, there is no commercially available polymers which could satisfy the requirements for excellent mould releasing properties without compromise the adhesion of the mould to the substrate. The surface treatment technique currently applied on the mould could hardly avoid the adhesion to the imprinted polymer and the situation becomes worse with the
11 increase of the density and aspect ratios of patterns, creating pattern defects not
acceptable for many applications at most time. Secondly, various applications might require different types of polymer or different polymer properties. A viable process should be as universal for materials as possible to expand its applications, instead of limiting its usage only to some specific materials.
Fabricating features with various micro/nano sizes at the same time may result in defects in nanoimprint lithography. Either incomplete polymer filling or a uniform thin layer of residual polymer will prevent the production of accurate replications. The alternative approach, called combined nanoimprint and photolithography (CNP), may solve this problem. This method combines photolithography with nanoimprint lithography just as shown in its name. A hybrid mask is applied as both NIL mould and photolithography mask (Chen, X., et al, 2004). In this hybrid mask, nanofeatures are placed at the UV transparent region and other regions are designed as a photomask for photolithography. Nanoscale features are formed by penetrating into the photoresist while large size features are formed through the standard photolithography process. Since only
thin residual resist is left on the substrate, the light diffraction effect is avoid at the region with nanoscale features.
Other derivatives of nanoimprint lithography have also been explored to solve the challenges mentioned above. One is to use solvents or monomers to soften polymers for imprinting at low temperatures (e.g., room temperature) and pressure (~1 N/cm2) (Khang,
D.-Y., et al, 2000). The disadvantages of this approach include involving non-
environmental friendly solvents and the relatively long evaporation time (~10min).
12 2.2.1.2 Bonding techniques
Most micro/nanofabrication techniques only create open nanochannels. The
bonding technique is required to seal the channels to complete functional nanofluidic devices. For Si or SiO2 based nanochannels, anodic bonding can be applied to seal
nanochannels to a rigid plate (e.g., cover slides, Stjernstrom, M., et al, 1998). But it can
not be extended to other materials, like polymers. The high voltage and high temperature
may also prevent this method from low-cost manufacture processes. Another alternative
sealing technique is to use PDMS (Mcdonald, J. C., et al, 2000; Chou, H.-P., et al, 1999).
Because PDMS can easily form conformal contact with most substrates, it can provide
good sealing against nanofluidic channels. But sometimes, the sagging of PDMS might
partially or completely block the nanochannels due to its rubber characteristics. Recently,
there are successful attempts to seal the nanochannels by using multiple vacuum
evaporations or shadow sputtering (Cao, H., et al 2002). But the whole process is very
complicated and non-trivial. Additionally, the channel geometry and dimensions can not
be precisely controlled during the sealing step. It might only practically useful to seal
very small nanochannels. Guo et al (Guo, L. J., et al, 2004) from University of Michigan
put forward another way to seal the nanochannels by imprinting against a glass cover
slides with a thin polymer film. The final nanochannel dimensions can be adjusted by
controlling the initial thickness of the polymer layer and the mould pattern configuration
(e.g., the ratio of the ridge width to the trench width on the template). Because of the
meniscus of polymer melts during sealing, nanochannels have a curved shape on the top.
13 2.2.2 Soft lithography
Soft lithography using PDMS is another option to create micro-/nanofeatrues.
Using the patterned elastomer (usually PDMS) as the mould, Whitesides’ group at
Harvard University developed a series of pattern transfer approaches, including
microcontact lithography (µCP), micromolding in capillaries (MIMIC), solvent-assisted
micromolding (SAMIM), microtransfer molding (µTM), and near-field conformal
photolithography (Xia, Y. et al 1998a, 1998b). Soft lithography is their collective name
for all these lithographic techniques. Compared to nanoimprint lithography, soft
lithography employs solvents to soften the polymer material, instead of heating the
material to several hundred degree centigrade that prevents the application of these methods in the biomedical field for peptides and protein delivery. Surface properties important for the pattern formation in soft lithography. It offers a simple process with high-resolution (~101-102 nm) and some of them can be used for nonplanar substrates,
unusual materials, and large area patterning.
Microcontact printing (µCP) enhances the development of self-assembly
membranes (SAMs). Its simple process may offer a low-cost, fast and mass transfer of these structures. In µCP, the PDMS stamp is wetted with an ink and then is brought into contact with the substrate surface. The ink is some chemical that can form SAMs on the substrate. Since the chemical is only transferred in the patterned area by the stamp, the
SAMs are generated with the same pattern on the substrate. These SAMs patterns can either serve as a resist protecting the underlying substrate in wet etching or as templates to selectively deposit other materials. Two key factors determine the success of µCP: the rapid formation of the SAMs and the autophobicity that prevent the ink from spreading
14 on the substrate. In µTM, a thin layer of liquid precursor is applied to the patterned
surface of a PDMS mold and the excess liquid is removed by scraping. Then this mold is
placed in contact with the surface of a substrate and the precursor is cured subsequently.
When the mold is peeled away carefully, a patterned microstructure is left on the surface
of the substrate. Usually, a thin (~100 nm) film will be left and can be removed by O2
RIE. Microtransfer molding can be applied in a wide variety of polymers or glassy carbon, sol-gels, and ceramics on relatively large areas (~3 cm2) within a short period of
time (~10 min). If letting a PDMS mold conformal contact the substrate at first to form
empty channels, it converts to another lithography method, MIMIC. In MIMIC, the
capillary force will push the low-viscosity prepolymer to fill the channels spontaneously.
So the pattern dimensions made by this method are limited due to the viscous flow and
gravity. SAMIM shares a similar principle as the hot embossing. Instead of using
temperature to soften materials and using a rigid mold to transfer patterns into the
substrate, SAMIM uses an appropriate solvent and an elastomeric PDMS mold to obtain
patterned structures. It can replicate some complex relief structures over large areas in a
single step. The limitation of SAMIM is the solvent. It must dissolve (or soften) the
material without affecting the PDMS mold. Good solvents for SAMIM should have a
high vapor pressure and a moderate surface tension (e.g. methanol, ethanol, and acetone),
which ensure a rapid evaporation of the excess solvent and minimal swelling of the
PDMS mold. Other solvents cannot be used in SAMIM, including those with low vapor
pressure (e.g. ethylene glycol and dimethyl sulfoxide) and those that swell the PDMS
mold (e.g. hexane, toluene and chloroform). However, solvents with a high surface
tension (e.g. water) can also be applied in SAMIM if choosing hydrophilic elastomers or
15 PDMS after surface modification (for example, by plasma treatment). Other materials can
also be added into the solvent if necessary for the resulting microstructures.
2.2.3 Sacrificial layer removal technology
Sacrificial layer techniques are not new to the microelectronic field. The
photoresists used in the pattern development can be called as the first generation of
sacrificial materials. In the rapid development of MEMS and BioMEMS techniques,
sacrificial layer techniques might play an important role in the fabrication of parts having
problems on de-moulding, e.g., creating moving parts or free standing parts, microcombs
with high pattern density, diverging shape parts or high aspect ratio parts.
The sacrificial layer removal approach can also be beneficial for creating nanofeatures. Since nanofeatures are more fragile, fabrication should try to avoid processes like demoulding, which are prone to damage the nanofeatures or generate defects. In the examples involving sacrificial materials to fabricate nanochanels or nanotubes, the sacrificial layer is temporarily embedded in other materials to define a desired geometry and then removed in subsequent processing. Materials used as sacrificial layers include silicon nitride, silicon oxide, metal and some thermally degradable polymers. Empty nanochannels are created after its removal by solvent dissolution, wet etching, or thermal degradation. The most promising advantage to create nanochannels this way is that it avoids the subsequent sealing of the open channels through bonding techniques.
16 2.2.3.1 Metal sacrificial layer
The first metal sacrificial layer was employed by Tonucci’s group (Pearson, D.
H., et al, 1995). Sacrificial aluminum layer was used to help release the thin metal layer from the substrate. An aluminum layer is sandwiched between the template and the substrate and finally dissolved in NaOH. This avoids the challenge of handling fragile
metal membrane (100nm in thickness).
Another examples using metal as the sacrificial material is to fabricate hybrid
nanotubes. Nickel nanorods (with the average diameter of 65±15 nm, length of 1.5±0.25
µm) were employed as the sacrificial materials (Subramanya Mayya, K. et al, 2001).
Through the layer-by-layer approach, polymer and organic-inorganic composite layers
were deposited on the outer surface of the nickel nanorods. After dissolving the nickel
core part, the hybrid nanotubes are formed.
Very recently, polyimide nanoslit was created using patterned aluminum stripes
as the sacrificial layer (Eijkel, J. C. T. et al, 2004). The aluminum thin layer (100~500 nm
in thickness) was sputtered onto the polyimide surface and patterned into long stripes by
photolithography and lift-off. With a second polyimide layer covering, empty
nanochannels were created after etching away the sacrificial aluminum layer. The
nanochannels demonstrated have the width varying from 2 to 30 µm and the length of 4
mm. To fabricate such nanochannels, 48 hours are required for the removal of the
sacrificial layer and the corrosive etchant (Merk) from the nanochannels.
17 2.2.3.2 Si-based sacrificial materials
SiO2 was firstly used as the sacrificial material to create nanochannels by silicon
surface machining (Chu, W., et al, 1995; Desai, T. A., et al 1998; Desai, T. A., 2000).
The size of Si nanochannels is tunable by control the thickness of the SiO2 sacrificial
layer. A series of silicon machining steps are involved and only a brief description of the
procedure is given here. Microchannels (2 µm wide separated by 2 µm and 5 µm deep) are created firstly on the Si substrate using dry etching. A thin SiO2 layer is then grown
over the entire exposed surface (including the surface of the microchannels). The microchannels are later filled by depositing polysilicon, followed by planarization for the
removal of excess polysilicon to expose the pores. A silicon nitride protection mask is
then patterned on both sides of the wafer to help opening membrane windows (2 × 3.5
mm) by etching Si wafer through the backside with KOH. After that, both the protective
nitride mask and the sacrificial oxide are removed using an HF etchant. This leaves open
nanochannels and provides the desired membrane structure. The SiO2 thickness can be
precisely controlled with the tolerance of less than one nanometer with the appropriate
thermal oxidation condition. Thus nanochannels created in this way have a uniform size
and well defined geometry. The size of channels can be precisely tuned through using
different oxidation periods. However, too many steps are involved to make the
fabrication process much intricate. Using SiO2 as the sacrificial material, the removal
involves the corrosive chemical etching. Since the long travel distance and short contact
area of reaction interface, the removal of the sacrificial layer in the long nanochannels is
very time-consuming. Sometimes, the process is impossible to be finished because of gas
18 bubble formation inside the narrow channels, blocking the transport path of reactants for
further etching.
2.2.3.3 Polymer sacrificial materials
Photoresists play an important role in photolithography. Depending on the
response to the exposure of light, photoresists can be divided into two types: positive
photoresists (the exposed regions dissolve and unexposed regions remain during the
development process) and negative photoresists (response in the opponent manner). Most
photoresists are mainly composed of a polymer resin, a photoactive compound and a
solvent that controls the mechanical properties of the resists. Since the photoresists would
be removed in the later process, they can be called the first generation of polymer
sacrificial materials. Polymer sacrificial materials reviewed here exclude the photoresists
used in photolithography.
A thermally decomposable polynorbornene, Unity400 (Goodrich), was probably
the first sacrificial polymer layer used (Kohl, P. A., et al 1998). It was applied to create
air-cavity. A polynorbornene was first spin-coated on the substrate and patterned into a
desired shape by standard photolithography and reactive ion etching (RIE). After
encapsulated inside another material, the sacrificial polymer decomposes into volatile
products upon heating to approximately 400oC. The decomposed products diffuse through the encapsulating material and leave behind a hollow cavity. Later,
o polynorbornen (PNB) was employed to fabricate nanochannels in SiO2 at 440 C. The decomposition temperature of PNB can vary in a wide range. These high temperature
19 decomposed polymers can bear moderately high temperature Si processes (e.g., silicon
oxide deposition).
Since the decomposition occurs at rather high temperature, only those materials
which can stand during decomposition are qualified to serve as the encapsulating material for final patterns. These materials include silicon dioxide, silicon nitride and other Si-
based materials. To extend such a process to other less thermal stable materials (e.g.,
epoxy-based materials and some polyimides), some polycarbonates (e.g., polyethylene
carbonate, PEC and polypropylene carbonate, PPC) were employed, which are known to
degrade in the range of 200-300oC (Metz, S., et al, 2004). Reed et al (2001) successfully
used such polycarbonate materials in the fabrication of microchannels (25 µm wide and 5
µm high) in inorganic glass, thermoplastic polymers (e.g., an Avatrel dielectric polymer) and thermoset polymers (e.g., Cyclotene 3022-57).
It requires specific properties for both sacrificial and encapsulating materials
when using thermally decomposable polymers as the sacrificial layer. The sacrificial
polymers must have good adhesion to the substrate and should not swell or degrade
during further processes involving solvents and other chemicals. The decomposition
products should easily permeate through the encapsulating layer with little or no solid
residue. The encapsulating material requires an adequate mechanical strength to avoid
sagging or distortion during decomposition and gases diffusion. These limitations plus
the relatively high temperature preclude the use of most functional polymers in the
sacrificial layer approaches.
20 Charles Martin and his associated synthesized gold nanotube membrane (GNM) using commercially available filters as the template (Cepak, V. M., et al, 1999; Menon,
V. P., et al 1995; Jirage, K. B., et al 1997). Track-etched polycarbonate membrane and anodic aluminum are two types of templates typically employed since their nearly monodisperse cylindrical pores. Such filters with uniform pores (10-100 nm in diameter
and 108-109 pores per cm2 of membrane surface) have been reported to synthesize
nanotube membranes made of gold, carbon, silica and even polymers, with gold nanotube
membrane as the most successful and useful one. Gold nanotubes were formed within
nanopores on the templates by an electroless plating method. In brief, the template
membrane was first deposited with Sn (II) and then Ag (0) onto all of the membrane’s
surfaces (pore walls and membrane faces) as the catalyst. The membrane was then
immersed into the electroless plating solution with Au (I) species and a chemical
reducing agent. There, a surface redox reaction occurred to yield Au (0) metallic thin film
on the exposed surface of the template (both the pore walls and membrane’s faces). Since
the reduction only occurred in the presence of the catalyst, this surface film did not block
the mouths of the nanotubes, leaving open nanochannels through the whole thickness of
the membrane. Finally, the template was dissolved to liberate the nanotube membrane.
By controlling the electroless plating time, the inside diameter of these nanotubes can be
controlled at will, down to molecular dimensions. Besides, a low deposition rate creates
nanotubes with a uniform thickness while a higher rate provides nanotubes with smaller
size on two ends than in the middle.
21 2.3 NANOFLUIDICS
2.3.1 Introduction
The advent of microfabricated systems has led to a growing interest in microfabricated fluidic systems with the characteristic scales in the range of several hundreds of micron to nanometer scale. Most microfluidic systems rely on two type of fluid transport: pressure driven and electrokinetic driven flow. At the early stages of microfluidics, the pressure driven flow was the favorite of most researchers as it is similar to the handling of fluids in the macroscopic pipes. With the dimensions of fluid channels shrinking to 50 micron or even smaller, it becomes difficult and unwise to use the pressure driven flow for further development. The pressure drop becomes tremendously high to make sealing and liquids handling impossible. In contrast, electrokinetic flows exhibit their advantages in micro-/nanoscale channels. Electrokinetic flows are relatively easy to operate as it avoids moving parts in devices for pumping and valving liquid. When scaling down the dimensions of fluid channels, no significant challenges occur in electrokinetic flows while an extremely high pressure drop is required in pressure driven flows.
2.3.2 Electrokinetics
Electrokinetic phenomena can be broadly divided into four types: electroosmosis
(the induction of flow at a charged surface by an electric field), electrophoresis (motion of charged particles in an electric field, the opposite of electroosmsis), streaming potential (the creation of a potential by fluid flow, the reverse of electroosmosis) and
22 polarization potential (the creation of a potential by particle motion, the reverse of
electrophoresis). Electroosmosis and electrophoresis are two important processes for
applications.
As mentioned above, electroosmosis is the bulk liquid motion induced by an
electric field in charged channels. Many materials, like glass, fused silica and some
polymers (i.e., acrylic or polyester), acquire surface charges when contacting with the
electrolyte solution. The extent of redistribution of surface charges relies on the local pH
and ion concentration of the solution (Hunter, R. J., 1981). With the surface charges, an
electric double layer (EDL) is formed and the counter-ions (with an opposite sign of
charge to the solid surface) is redistributed near the solid surface. They reside in two
regions: a Stern layer (where counter ions stay stagnant) and a Gouy-Chapman Diffusion
Layer (where mobile ions obey Boltzmann distribution). The two layers are separated by
the shear plane. In most cases, it satisfies the Debye-Huckel limit of the EDL and the
-ζ Debye length (λD) is the length from the shear plane where the EDL potential falls to e s
value (ζ s is the zeta potential of solid surface). The value of the Debye length can be calculated by the following equation:
εkT λD = 2 2 (2-1) 2z e n∞
where z and n∞ are the valence and the bulk ionic number concentration of type-i ions. ε is the medium permittivity, e is electron charge (=1.602×10-19 C), k is Boltzman constant
(=1.38×10-23 J/K) and T is the absolute temperature.
23 In many electrokinetic systems, the Debye length scale is much smaller than the characteristic dimensions of the channels (i.e., the hydraulic diameter). For these thin
EDL cases, the potential through the cross section area is zero and the velocity of the electrosmosis flow under zero pressure gradients is described by
εζ u = − s E (2-2) η
Through the definition of mobility (the ratio of velocity to electric field), µ = u / E , the mobility of electroosmosis flow (EOF) under zero pressure gradient becomes
εζ µ = − s (2-3) EOF η
In homogeneous microfluidic systems, the EO mobility is a constant for a given wall material and solution chemistry in the absence of Joule heating effects and EOF has a nearly plug velocity profile (Molho, J. I., 2002). This comes from the moving boundary at the electrical double layer region associated with the motion of ions and electrolytes.
The variation is limited inside a slim region adjacent to the channel wall. As the result, less solute dispersion occurs in the cross section. EOF has advantages in the flow control and species transport in microfluidic systems without a high external pressure. Thus, EOF has been applied in micropumping to delivery liquid, such as buffer solutions in microfluidic systems.
EOF can also generate non-plug like flow profile when heterogeneous zeta potentials are created on the wall surfaces of microfluidic systems (Herr, A. E., et al,
2000). The Joule heating effect can change the conductivity and viscosity of the fluid in
24 EOF, leading to the non-uniformity of the velocity profile (Xuan, X., et al, 2004). More interesting flow patterns, like multidirectional flow or recirculating flow, can be created by purposely varying charge – sign and density – on the local surface in a certain direction: perpendicular to the applied electric field for bidirectional flow and parallel for recirculating flow (Strook, A. D., et al, 2000 and 2003; Barker, S. L. R., et al, 2000).
Such patterned flows can be applied to mixing in microfluidics and the control of dispersion (band broadening).
However, as liquid motion is involved in EOF, pressure gradient will be generated sooner or later if the liquid is not move out. There might not be mismatch of fluid level at the reservoirs at the starting of the experiments, but the pressure gradient created by EOF will eventually be inevitable. If such a phenomenon is not favorable for applications, electrophoresis is a better choice (e.g., biomaterials, like DNA and proteins, transport study in nanofluidics).
Electrophoresis is the induced motion of charged objectives (i.e., colloidal particles or molecules) suspended in ionic solutions resulting from the external electric field. Most applications of electrophoresis are well approximated by one of two limits described below. The first limiting situation happens when the size of particles or molecules (mostly, ions and solutes) is much smaller than the Debye length. The presences of charged objectives and its counter ions around will hardly affect the electric field lines. The motion of these molecules can be described by Stokes-Einstein equation
(a balance between the electrostatic force and the viscous drag associated motion on the molecule). The electrophoretic mobility of a charged molecule is
25 q µ = ,(d << λ ) (2-4) EP 3πηd D
It is a function of the molecule's effective size (d, the diameter of a Stokes sphere) and directly proportional to its carried charges (q, the total charge on the molecule). The second one take places when microspheres with a thin Debye layer (e.g., polystyrene spheres with the diameter of 100–10,000 nm, cells or organisms) migrate under an electric field. In this case, the surface/charge interaction is similar to the flat wall electroosmosis flow. The electrophoretic velocity is affected by the electrostatic forces
(both on the surface charge and on their charge double layers) and the viscous drag associated with both the body motion and the ionic cloud motion. Thus the mobility of an electrophoretic particle reduces simply to
εζ µ = p ,(d >> λ ) (2-5) EP η D
Electrophoresis has been extensively applied in separating, sizing, and sequencing biomolecules, such as DNA, RNA and protein, using either gel or linear polymer solutions as the sieving media. (e.g., slab gel electrophoresis, capillary electrophoresis).
Recently, electrophoresis has been extended in microchannels to alleviate the high energy dissipation drawback (due to Joule heating) of conventional techniques. Microchannels have a larger surface-to-volume ratio, which is much beneficial for overcoming such a challenge. Electrophoretic microchips can greatly reduce the reagent consumption, shorten the analysis time and cut the cost. In addition, it can also increase diagnostic efficiency, obtain high throughput and integrate to form portable units. A number of such chips are now available in market, such as LabChip® for integrated analysis of DNA,
26 RNA, and protein, providing fragment size and concentration information in half an hour
(Caliper Technologies Corporation) and LabCard™ for rapid DNA fragment sizing, using plastic chips with pre-filled gel (Aclara Biosciences Inc).
2.3.3 Molecular transport in nanochannels
Geometric constrictions to molecular transport have been studied for more than half a century. Starting from the investigation of the hindered diffusion in porous materials (e.g., membranes, chromatography, and catalysis), many unusual phenomena are uncovered, which help understand the physics of molecular manners at the nano or even molecular scale and to develop new techniques for separations, reaction engineering, and sensing. Steric and/or hydrodynamic hindrances have been well studied, but most stayed as theories or statistic average results. The availability of track-etched membranes provides good model experimental tools to directly check the validity of a theory without any tortuosity correction. Since then, great progresses have been made in both theory and experiments in quantifying the effects of solutes (e.g., concentration, charges, size, shape and rigid degree), pores (size, shape, charges) and interactions between solutes and pores (electric interaction, hydrodynamic viscous drag, flow-induced deformation, etc) (Deen, W.M., 1987). Basically, the transport coefficients are dramatically reduced with the increase of the size ratio of the solute to the pore. Charged solutes have a sharper declined rate on the diffusion coefficients. Different from the hindered transport of rigid solutes, flow induced deformation of flexible polymers might happen, depending on the product of strain rate and the longest relaxation time. The
27 interactions of solute to solute begin to be significant when the molar concentration exceeds a threshold value C* (refer to Appendix IV).
The experimental studies lag behind to the development of theory for the hindered diffusions. Few experimental data are available to check the validity of those theories and most of them are restricted only to the cylindrical shape pores as well as a few rigid colloids and flexible polymers. Recently, remarkable progresses were made in experimental studies with the success of many novel techniques to obtain well-defined channels with desirable geometries.
2.3.3.1 Molecules transportation in nanotubule membrane
As mentioned in the previous section, nanotubules in Gold or Carbon Nanotubule
Membranes (GNM, or CNM) can have the same magnitude dimensions as molecules.
Hindered diffusion occurs in these Au nanotubules, showing a higher molecule selectivity on the basis of molecule size. For a pair of different size molecules, the tris-bipyridal
2+ 2+ complex of Ru(II), Ru(bpy)3 (large) and methyl viologen, MV (small), the selectivity was substantially raised from 1.5 (in the free aqueous solution) to 50 with 5.5 nm nanotubules and to almost completely block of the large molecule with 0.6 nm nanotubules (Jirage, K. B., et al 1997). The later result also exhibited that the Au nanotubule membrane could cleanly separate large molecules from small molecules with nanotubule dimensions larger than the molecules. Through spontaneously chemisorbing of thiols (i.e., linear C3, C10, and C16 alkanethiols and hydrophilic thiol HS-C2H4-OH) to the inner tubule walls, they demonstrated that Au nanotubules membranes could be applied to separate molecules based on their different chemical properties, i.e., the
28 separation of hydrophobic from hydrophilic molecules (Jirage, K. B., et al 1999). With
C16 modified nanotubules, pyridine and toluene could be successfully separated with 2 nm nanotubule membrane. They also found that nanotubule membranes could carry switchable charges on the Au wall surface, depending on the orientation of biased potential (Nishizawa, M., et al 1995). Surface charges had been successfully attained either directly submerging into electrolyte solutions carrying non-adsorbing anions (e.g.,
- - - F ), or combining with thiol-modification (e.g., C3) when adsorbing anions (e.g., Cl , Br ) present.
Track etched membranes have some structural imperfections: (1) the nanopores are not absolutely monodisperse and serious defects exist due to the overlapping of two or more pores. The problem is exacerbated when the pore size becomes smaller; (2) the pore length is not consistent because of variation in track tilt angle during manufacture.
These drawbacks limit their applications and the development of hindered transport studies. Very recently, great progresses were made in experimental studies with the success of many novel techniques to obtain well-defined channels with desirable geometries.
2.3.3.2 Biomolecules transportation in 1D nanochannels on Si membrane
Ferrari and his colleagues created silicon membranes consisting of rectangular nanochannel array by top-down silicon micromaching techniques (reference to the fabrication review part and Martin, F. J., 2003). These nanochannels provide one dimensional nanoscale channels with tunable uniform channel width ranging from 7 nm to 50 nm. They made capsules for islet cells using these Si nanochannel membranes as
29 the only connection to the external environment, working as “nanogates”. As the channel size is tunable, different transport characteristics occurred to the passage of nutrients and wastes (i.e., insulin, glucose, oxygen and carbon dioxide) and to the blocking efficiency of immuno molecules (i.e., cytotoxic cells, macrophages, antibodies and cytokines). With the decreasing the width of the nanochannels from 27 nm to 7 nm, the expected exponential increase of the cumulative flux of glucose (obeying the Fick’s law, first-order kinetics) was slowed down and finally decayed to a linear relationship with time (zero- order kinetics) when the nanochannel width shrank to 7 nm. This was further confirmed by the factor that the initial flux of glucose was independent of the initial glucose concentration (165-1000 mg/ml). In vivo release of bovine serum albumin (BSA, 66,000
Dalton) was studied with such Si nanochannel membranes and similar zero-order release rates were obtained for both 13 nm and 26 nm nanochannels. To test the blocking efficiency for immuno molecules, IgG was applied with these Si nanochannel membranes
(Desai, T. A., et al, 1999). The data showed that the complete isolation of IgG requires an absolute channel width below 18 nm. After 24 hours, less than 0.4% IgG diffused through the 18 nm membranes and only 2% after over 150 hours. Compared to other colleagues’ results using commercial polymeric membranes, this immunoprotection was highly efficient.
Electric forces can be useful to study the transport through well-defined nanochannels. As mentioned previously, a motion driven by the electric field avoids a large pressure drop between two sides of the thin layer with nanopores or nanochannels.
Electrokinetics can dramatically enhance molecule transport through nanochannels or
30 nanopores. Besides, electrophoresis is the most popular one used in the sensing of single nucleic acid and other biomolecules in nanopores or well-defined nanochannels.
2.3.3.3 Molecule sensing and sorting in single nanopore or nanochannel
Nanopores have been explored in the past decade to analyze single nucleic acid molecules. Single nucleic acid molecule carries a net negative charge and can be electrophoretically driven through by applying an electric potential across the nanopores.
Since the nanopores are the only path for macromolecules, each translocating molecule blocks the open pore ionic current, forming a transient current drop. Through studying the transient ionic conduction signal, the translocation information of molecules in the nanopores can be obtained and used to estimate the local macromolecule concentration.
These electrical signals depend on the characteristics of biological molecules. To resolve the molecular structure, the dimensions of the nanopores must fall in a range: large enough for molecules to pass while small enough to avoid averaging molecule configurations due to Brownian motion.
α-haemolysin (alpha-HL) ion channel is a heptameric protein with a ~2 nm diameter inner pore which allows translocation of single-stranded DNA (Kasianowicz, J.
J., et al, 1996; Deamer, D. W., et al, 2002; Sauer_Budge, A. F., et al, 2003). Analysis of externally induced ion current pulses across the pore during its interaction with DNA can provide information about the DNA molecule, including length, base composition and dehybridzization kinetics of dsDNA. In conjunction with the chemical modification of the surface of the pore or the analytes, the sensitivity of alpha-HL pores can be improved to the translocation of particular molecules or to the detection of other biomolecules and
31 analytes of interest. Some attempts include grafting ssDNA fragments and attaching polyethylene glycol (PEG, 3.4 kDa) or β-cyclodextrin. However, these biopores are out of practical applications like other natural materials because of their stability and noise are restricted by their chemical, electrical, mechanical and thermal constraints.
Desired solid-state nanopores might overcome some of these limitations. Focus ion beam (FIB) was applied to create single 3-nm nanopore or 10-nm pore in silicon nitride (Si3N4) thin films with the thickness of 5-10 nm (Li, J., et al, 2003). The dimensions of these nanopores are comparable to the alpha-HL biopores. 10nm Si3N4 nanopores showed a shorter current blockage duration time but a bigger (100-200 pA) ionic current drop for 10 kbp dsDNA. The translocation time inside the nanopores is approximately proportional to the dsDNA molecular weight and inversely proportional to the transmembrane potential. Three level blockades were observed in a histogram of the ionic current drops over all events, which were explained as the molecular folding when passing the nanopores.
Saleh, O. A., et al (2003a and 2003b) have created a single 200-nm diameter straight nanochannel in poly(dimethylsiloxane), (PDMS) using multistage lithography techniques. In these nanopores, the result of λ-phage DNA molecules (48.5 kbp) showed an ionic current drop of less than 12 pA with the total open-channel current of approximately 15 nA. The signal is not very resolvable over noise but provides an idea of the limitation of nanochannel dimensions for sensing the conformation of molecules of such a size. Additionally, their approach provides a possible alternative way for the development of low-cost robust nanochannels.
32 A single conical nanopore was fabricated by Mara and coworkers in the track- etched polyimide membrane (Mara, A., et al, 2004). The pore has a ~2-7 nm on the sharp end and 2 µm on the other. Its ultrahigh aspect ratio contributes to the long duration time of DNA translocation in the pore. As a demonstration, three different lengths of fragments of dsDNA (286 bp, 974 bp and 4126 bp) were used in their study. The blockage duration time was roughly proportional to the fragment length, similar to the conclusion drawn in α-hemolysin pore and silicon nitride pore studies. Short fragments showed unclear current peaks unless the fragment length was closer to 1000 base pairs.
Through tuning the geometry of nanopores, it might provide a higher resolution than the solid-state nanopores and alpha-HL biopores so that it might be able to detect fragments with only several hundred base pairs.
A single carbon nanotube membrane also has been used for sensing molecules
(Ito, T., et al, 2003). The channels reported have a diameter from 50 to 160 nm and a length of 0.94-1.26 µm. since the inner surface of the channel is uncharged, the motion of molecules driven by the electric field reduces to purely electrophoretic transport. Ito and his coworkers took advantages of this characteristic to measure the size, charge and mobility of polymeric nanoparticles. This method provides a way to directly obtain the size distribution of individual particles dispersed in solution. It might also conveniently determine the pKa values of charged molecules or nanoparticles.
33 2.4 DNA MANIPULATION TECHNIQUES
2.4.1 Introduction
Direct manipulation of single macromolecules can provide a wealth of new biochemical information and greatly improve our understanding of the nature of molecule interactions and polymer physics at the molecular level. Compared with average properties over an ensemble of macromolecules through traditional “bulk” measurements, direct testing and visualization of single molecules in the flow field can better reveal fundamental insights in complex fluid systems. For the time being, long DNA segments are the first choice of scientists because of their monodispersed molecular weight and easily obtained visible long chains. Through fluorescence labeling, the behaviors of individual molecules can be observed and recorded under fluorescence microscopy.
DNA superhelix structure consists of two polynucleotide strands connected loosely by hydrogen bonds through the base pairs: adenine (A) to thymine (T) and guanine (G) to cytosine (C). This structure carries the life genetic code and can be transferred strictly through the constraint binding rule for the four bases. In a short length scale, the electrostatic repulsion between two negatively charged strands makes the structure rather stiff. Such length scale is known as the persistence length, which is about
50nm or 150 base pairs. DNA segments much longer than the persistence length twist and tangle into a superhelix coil-like configuration. A supercoiled λ-DNA, 48.5 kbp) has a random coil structure with a diameter of about 1 µm and a contour length of around 17
34 µm if fully stretched to be straight. The supercoiled DNA behaves like ideal, flexible polymers in solutions.
Mechanically distorting or stretching of DNA under controlled conditions has been comprehensively explored by both experiments and physical theories. The stretching of DNA includes mainly several types: (1) stretching DNA molecule parallel to the helix axis; 2) unzipping DNA helix by pulling perpendicular to the helix axis; (3) twisting DNA molecule.
Many micromanipulation techniques have been employed to study the extension and relaxation of DNA double helix. These techniques can be divided into two main groups: one employs mechanical forces to stretch DNA and study its force response. This includes using AFM, optical and magnetic tweezers and so on. Many characterization data of DNA were determined by these techniques. The other uses fluid flows to induce
DNA stretching. This includes using different fluidic techniques to study the configuration histogram of DNA. Here, only the second group of techniques is summarized.
2.4.2 Hydrodynamic DNA stretching under different flows
The coil-stretch transition of single DNA molecules in well-defined extensional flow has been carried out using hydrodynamic forces. For example, the deformation of λ- phage DNA was studied in Newtonian flow by a Taylor four-roller mill device (Sasaki et al, 1996). The birefringence profiles indicated the reduced criticality of the coil-stretch transition in such an extensional flow field. The dynamics study of a pressure-driven planar extensional flow was explored by Chu and his coworkers at Stanford University
35 with a microfabricated cross-slot flow cell (Perkins, T. T., et al, 1997; Smith, D. E., et al,
1998; Babcock, H. P., et al, 2003). They focused on the relationship of the strain rate and the probability distribution of the molecular extension by tracking individual molecules.
The onset of polymer stretching takes place when the Deborah number (defined as the product of strain rate and the longest relaxation time, De = ε&τ ) exceeds the rigorous theoretical prediction value ( De ≅ 0.5 ). The conformation of DNA were classified into seven different configurations: ball (coiled), folded, kinked, half dumbbell, dumbbell, extended and uniform. Different conformational shapes showed diverse dynamics when the strain rate exceeds a critical value (i.e., 0.5 s-1). They also tracked the entire time evolution of DNA deformation in the startup of an extensional flow. Such studies are direct evidences to discover the nature of the conformation development during stretching. The stretching rate of individual molecules varied highly, depending on the initial equilibrium conformation of polymer coil and the stretching development it experienced.
Recently, a hydrodynamic focusing technique was applied to study the dynamics of single DNA molecules under the extensional flow (Wong, P. K., et al, 2003).
Sandwiched by two other flow streams of a buffer solution, a stream of DNA solution in the middle is forced to converge by hydrodynamics focus. In the middle flow, slip boundaries are achieved, leading to a plug-like velocity profile in the spanwise direction and a constant spatial acceleration in the streamwise direction. With such a hydrodynamics focusing technique, the coil-stretch transition was observed for T2 DNA with the Deborah number large than 0.8. By suddenly stopping the flow, they tracked the relaxation and recoil development for stretched DNA molecules. The longest relaxation
36 time determined in this way for T2 DNA is around 0.63 s with the scaling viscosity of 0.9 cP.
Shear flow is another type of flow commonly met in practice. Similar to the
Deborah number mentioned in the elongational flow, Weissenberg number (defined as the product of shear rate and the relaxation time,Wi = γ&τ ) is applied to relate the probability distribution of DNA extension. According to classical polymer theories, a flexible polymer starts stretching and aligning when the Weissenberg number is equal or larger than one. Chu’s group also studied the DNA dynamics in the shear flow (Smith, D.
E., et al 1999: Hur, J. S., et al, 2001). As the DNA molecules stretch, various aperiodic temporal fluctuations occur. The molecular chain can continue stretching or tumbles end- over-end, depending on the dominant items of several forces (hydrodynamic drag,
Brownian motion, etc). LeDuc et al from Johns Hopkins University found that various conformations of flexible DNA (i.e., T2 DNA) existed in a shear flow, with the orientation varying from parallel to perpendicular with respect to the shear plane (LeDuc,
P., et al, 1999). Different from the theoretical prediction, they found a shear induced deformation of T2 DNA with shear rates much smaller than the inverse of the relaxation time (Wi < 1.0).
Muller and her coworkers at University of California-Berkeley studied the stretching of DNA in a microfluidic device with a converging inlet (Shrewsbury, P. J., et al, 2002 and 2001). They focused on the concentration effects and fluid effects to the deformation of DNA. They found that flexible DNA molecules showed less stretching in a concentrated solution than those in a diluted solution. Profound stretching of DNA was
37 found in both the contracted region (from the reservoir to the channel) and high shear regions (close to the channel walls).
Other studies about flow stretching or aligning DNA include moving a solid surface which is adhesive to DNA solutions, like spinning DNA solution on a smooth surface and pulling glass from DNA solutions (Michalet, X., et al, 1997; Namasivayam,
V., et al, 2002; Jing, J., et al, 1998). Spin stretching DNA is rapid and efficient and the stretching is the combination of centrifugal force, hydrodynamic force and surface tension. The transient flow involved can provide stretching and fixation simultaneously as the liquid becomes more viscous due to evaporation and fixation. A similar method includes using fluid flows developed within tiny evaporating droplets to elongate and fix
DNA molecules onto derivatized surfaces. A dynamic molecular combing technique was reported to stretching the whole human genome by pulling silanized solid surface. The meniscus at the interface during the pulling of cove slides exerts a constant dragging force for DNA stretching. The quick dry characteristic of hydrophobic surface helped fix the stretched state almost instantaneously. This dynamics molecular combing can provide stretched and aligned DNA molecules in a single direction over the whole solid surface and also precisely uniform stretching at a resolution as high as 2 kbp/µm.
2.4.3 DNA stretching study using nanopost and nanoscopic slit
Although the mechanism of electrophoretic DNA separation in the gel or linear polymer solution is still not very clear, the entanglements of DNA molecules with polymer molecules or polymer clusters is widely believed to play a very important role.
Directly observation of DNA conformation in various polymer solutions (e.g., hydroxyl
38 ethyl cellulose, HEC and linear polyacrylamide, PA) under a DC or AC electric field provides evidences of two basic migrating ways: U-shape motion or I-shape motion
(Namasivayam, V. et al 2002; Ueda, M., 1999; Blanch, H. W., et al, 1998). In a U-shape collision, the DNA collides with a polymer obstacle and starting to extend into a U-shape or V-shape and then slides around the polymer obstacle like a pulley in the direction of the longer partition. Occasionally, multiple entanglement points exist, resulting in more complex deformation, like W-shape. The frequency of these interactions increases as the
DNA size increase, so do when the polymer size or concentration increases. For example,
λ-DNA (48.5 kbp) keeps its coiled conformation while T4 DNA (166 kbp) is deformed frequently, in a 0.09% solution of HEC 438K.
Some artificial obstacles were fabricated by micro-/nanotechnologies to serve for the DNA stretching or separation studies using only buffer solutions. For example, a
Japanese group (Kaji Noritada et al, 2004) built high aspect ratio nanopillar arrays inside the DNA separation microchips. The space between two pillars (500 nm) is comparable to the sieving matrix of an agarose gel at the concentration of 1%. The U-shape conformation was found for T4 DNA in such a periodic nanopillar channel but not for λ-
DNA. Optimal nanopillar designs have been demonstrated to effectively separate large
DNA fragments within minutes. Another example of artificial obstacles is using nanospheres. Such nanospheres have a cross-linked PLA core and a PEG shell. The migration of DNA between the gaps among packed nanospheres is similar to those in nanopillar array channels. With packed nanospheres channels, Tabuchi Mari reported that
39 DNA fragments up to 15 kbp had been successfully analyzed within 100 seconds
(Tabuchi, M., et al, 2004).
With such well-defined artificial obstacles, the impact dynamics of single DNA– obstacle collision was also studied systematically. Similar to the entanglement of DNA with polymer molecules, DNA obstacle collisions can be classified as “hook” or “roll- off” (Randall, G. C., et al, 2004). The probability of these two types of collision depends on a geometry ratio (the ratio of the obstacle radius to the DNA equilibrium radius of
gyration, Robs /Rg ) and a dynamic parameter (the ratio of DNA maximum deformation
rate to the relaxation, De = 2µEτ / Robs which is similar to the Deborah number defined
before). With Robs / Rg ≥ 1. 0 and 0.5 < De < 40 , the “hook” collision is more likely to form. The deformation of DNA was concluded to be carried by the purely extensional velocity field induced by those obstacles in the external electric field. Simulation results showed that smaller DNA molecules might experience more “hook” collision than larger
DNA because of its fast relaxation. The long collision time of larger DNA molecules makes them lag behind the smaller DNA to obtain the efficient electrophertic separation.
Another novel obstacle design for the study of DNA stretching and separation is the so-called entropic trapping (Han, J., et al 1999 and 2000). With a sequence of deep and shallow channels, the mobilities of various size DNA molecules are different in an external electric field. Because of the gyration radii of DNA used is much larger than the microfabricated thin nanoslits, DNA were trapped in the thick regions for a certain time, depending on the DNA molecule size. The recoil of trapped DNA in the confined nanoslits was also studied by the pulse field electrophoresis (Turner, S. W. P., et al,
40 2002). Entropy is regained when DNA molecules escape and relax into the random coiled configuration. Larger DNA molecules escape the trapping regions faster than the smaller
DNA, probably because larger molecules do not have a high probability to stay entirely within the thin nanoslits and are able to coil back outside rapidly.
41
CHAPTER 3
SINGLE DNA DYNAMICS IN CROSS-SLOT ELECTROKINETIC FLOWS
3.1 INTRODUCTION
In this chapter, single or multiple cross-slot(s) microfluidic platforms were designed and applied in the study of single DNA dynamics in electrokinetic flows. An electrokinetics-induced stagnation flow was at first created inside a single microscale cross-slot channel. Compared to hydrodynamic-driven microfluidics, this flow system can be readily assembled and easily operated. No complicated fabrication and assembly of flow cells are required, compared to the flows driven by hydrodynamic forces.
A fairly homogeneous, two-dimensional elongational flow was observed and the dynamics of flow-induced DNA deformation was studied. The initial conformation of
DNA molecules and residence time inside the flow field play important roles in determining the extent of DNA stretching. Other widely used flow patterns, for instance, shear and rotational flows were also generated by electrokinetics. Considering the complicated interactions between charged species in the fluid and surfaces of channel walls, more stable flows were performed in a fractionally extended five cross-slots
42 microfluidic network. Unlike other microfluidic devices, the flow patterns generated in this five-cross design are largely independent of the charge density and polarity of species in the fluid. Different flow types can be quickly switched by changing the voltage inputs. Charged polystyrene microspheres were used to identify the flow characterizations and the conformational evolution of single λ-DNA molecules under these three basic flows were investigated. A coarse-grain molecular simulation also was applied in several cases, providing reasonably well agreement with experimental observations.
3.2 EXPERIMENTAL
3.2.1 Materials
The testing fluids include Fluoresbrite Yellow Green (YG) microsphere suspensions (Polysciences, Inc, Warrington, PA) and bacteriophage λ-DNA (48.5 kbp,
New England Biolabs, Ipswich, MA). β-mercaptoethanol, glucose and sucrose were purchase from Sigma-Aldrich (St. Louis, MO) and used as received. 5x TBE (tris-Borate-
EDTA, pH=8.3), TE10 (Tris-HCl, 1mM EDTA, 10mM NaCl, pH=8) were purchased from Fisher Scientific (Pittsburgh, PA) and used as received for preparation of DNA solution. λ-DNA was labeled with a fluorescent dye, YOYO-1 (Molecular Probes,
Eugene, Oregon), at a dye-base pair ratio of 1:5.
The polymethyl methacrylate (PMMA) sheets and films with different thickness were purchased from McMaster-Carr (Cleveland, OH). After fabrication, PMMA was
43 cleaned ultrasonically in the mixture of isopropanol and distilled water for a duration of
2~5 minutes before use.
3.2.2 Fabrication
For prototyping, computer numerically controlled (CNC) machining is used to fabricate single or multiple cross-slot(s) microfluidic platforms. The designed microfluidic patterns were drawn using commercial AutoCAD software (AutoCAD 2000,
AutoDesk, Inc). Channels (with D/depth = 250/150 µm or D/depth = 1/1 mm) and reservoirs were machined on a polymethyl methacrylate (PMMA) plate by a CNC machine (Sherline Products, Inc, Vista, CA). The end mills (single-end two-flute sub- miniature end mill) were purchased from McMaster-CARR (Cleveland, OH) and the diameter of the mills ranged from 0.005” to 0.039”. Figure 3.1a shows a micromachined poly(methyl methacrylate, PMMA) plate with cross-channels of 250 µm in width and
125 µm in depth. The diameter of the wells is 1.5 mm and the length of each arm is 7.5 mm. Figure 3.2a shows a five cross-slot poly(methyl methacrylate, PMMA) platform with the ratio of width to depth equal to 1 or width/depth = 250/250 µm). A 45 µm thick
PMMA film was then thermally laminated onto the surface of the substrate to form closed channels.
3.2.3 Sample preparation
For clear imaging purpose, polystyrene microsphere (700nm, 2 µm and 3 µm) suspensions of 0.00265% were made by diluting stock suspensions (2.65% solid content)
44 by a factor of 1000 in demineralized distilled water (pH = 5.5). 5 min ultrasonic treatment is done right before loading into microchannels to avoid possible aggregation.
Fluorescently labeled bacteriophage λ-DNA was prepared using the following protocol. Stock λ-DNA solution (~0.5 mg/ml) was diluted by a factor of 50 in TE10 and heated to 65 oC for 10 min to free the complementary sticky ends of DNA molecules.
After cooled to room temperature, the DNA solution was then diluted by another factor of 40 into a solution that was made by mixing 100 parts TE10 and 1 part 10-5 M YOYO-1 solution. The solution of 10-5 M YOYO-1 was made by diluting YOYO-1 stock solution by a factor of 100. The DNA/dye solution was then allowed to sit for at least 1 hour at room temperature in the dark before being used. To increase the stress added on DNA molecules, the bulk viscosity of DNA solution was finally adjusted by adding glucose and sucrose. In most cases, 18% (w/w) glucose and 40% (w/w) sucrose were added and the final bulk viscosity of DNA solution is 30 cp.
3.2.4 Characterization
The cross section of fabricated cross slots was examined by a scanning electron microscopy (Hitachi 4300, Hitachi, Japan). The dimensions and surface roughness of the microchannels were characterized by an optical profilometer (WYKO NT3300 Profiling
System, Veeco Metrology Group, Tucson, AZ). The electrophoretic (EP) mobility of particles was measured using a ZetaPALS zeta potential analyzer (Brookhaven
Instruments Company, Inc). The electroosmotic (EO) mobility of different solid surfaces was characterized using a BI-EKA streaming potential analyzer (Anton Paar, Inc,
Austria).
45 3.2.5 Experimental setup
For single cross slot, the microscale cross-channel was first filled with 1x TBE buffered solution. 1X TBE buffered solution (pH= 8.3) was made by diluting the stock
TBE buffer (5X) by a factor of 5. An electric bias of 147 volts was applied to wells 2 and
4 with wells 1 and 3 were grounded. The arrangement of the voltages is shown in Figure
3.1b. The whole chip was mounted onto the stage of an inverted epi-fluorescence microscope (TE 2000S, Nikon, Japan), as shown in Figure 3.3. Negatively charged polystyrene microspheres (700nm, C=0.00265%) were used as the tracer for flow visualization with 10x and 40x objective lens. The streamlines of the flow pattern were generated by compounding the video graphs. The dynamics of DNA conformation was observed with a 100x/1.3 NA oil immersion objective lens and recorded by a Coolsnap
HQ CCD camera (Roper Scientific, Inc, Tempa, AZ).
For five cross-slots, the device (with D/depth = 250/250 µm or D/depth = 1/1 mm) was micro-machined on a poly(methyl methacrylate) (PMMA) plate sealed by a 45
µm thick PMMA film. The microchannels first were filled with DI water, followed by adding fluorescence-dyed polystyrene microspheres with a diameter of 3 µm as the tracer at the concentration of 0.00265% for flow visualization. The arrangement of the voltages is shown in Figure 3.13a for extensional and shear flows, where the actual voltage (in
Volts) needs to be multiplied by a factor of 25. To generate rotational flow, the arrangement of voltage inputs is switched to case shown in Figure 3.15a, with the actual voltage (in Volts) needs to be multiplied by a factor of 50.
46 3.3 SINGLE CROSS-SLOT FLOWS VISUALIZATION
With the same applied voltage arrangement, different flow patterns, i.e., extensional flow, shear flow and rotational flow, in principle, can be generated in the same single cross-slot, as shown in Figure 3.4. For example, if the four arms carry the same charge (positive or negative), extensional flow will be achieved. If the pair of arms in the diagonal carries the same charge but contiguous arms have different charges, shear flow and rotational flow can be achieved. If one pair is neutral and the other pair is charged, shear flow will be generated. If two pairs carry opposite charge, rotational flow will be generated.
3.3.1 Extensional flow
3.3.1.1 Velocity field imaging and analysis
In the single cross slot, if a two-dimensional pure elongational flow is generated, the velocity profile should satisfy(u, v) = (ε&x, − ε&y ) , and the velocity magnitude V is given as:
2 2 V = ε& × x + y (3-1) where x and y are the distance away from the stagnation point and ε& is the elongational rate, which is a constant for a given fluid. The streamlines inside this single cross-slot is shown in Figure 3.5a. As the comparison, the compounded image of electrokinetically driven elongational flow is shown in Figure 3.5b. It nicely followed the streamlines governed by equation 3-1. The velocity vector map also is shown in Figure 3.5c and experimental data is plotted in Figure 3.5d. Because of symmetry, the result is only
47 shown in the first quarter. The maximum velocity gradient exists along the centerline of the cross. Other regions have less steep gradient and much non-uniform velocity profile.
Since cross-slot(s) are made of PMMA, their surfaces carry weak negative electrostatic charges. However, those charges are derived from weak acids, e.g., -COO- and in solutions, not all such groups are dissociated and the charge density is strongly dependent of the pH value of the solution. Besides, the dissociation reaction happened randomly along the polymer chain, which could readily change due to the change of local condition. Therefore, electroosmosis also plays some roles beside electrophoresis for those charged species in this cross-slot microfluidic platform. Under the applied electric field, the overall velocity of particles is the combination of electrophoretic movement of the particles and electroosmotic flow of the fluid, which can be expressed as follows:
Voverall = VEP + VEOF (3-2) where V is the velocity and the positive direction is defined from cathode to anode,
VEOF = µ EO E , VEP = µ EP E , where µ EO and µ EP are given by equations 2-3 and 2-4 or
2-5, respectively. Note that both PS particles and channel surface carry negative charge, the component of velocity contributed by electrophoresis and electroosmosis have opposite motion directions. At the experimental conditions, the actual direction of particle movement was observed from wells 1 and 3 to wells 2 and 4, which indicated that the electrophoretic movement of polystyrene microspheres overcame the electroosmotic flow. The elongational motion inside the cross region is electrophoresis- dominated.
48 Figure 3.6 compares the electric lines of the electric field (the thin solid lines) and the streamlines of the pressure driven flow (the dashed lines) with streamlines of the pure elongational flow (the thick solid lines) for the first quarter of the intersection area in the cross-channels. Since no electric source is inside the microchannel, the Laplace equation is satisfied for the electric potential φ and the electric field vector E = −∇φ can then be calculated. The streamlines of the pressure driven flow are calculated using the Stokes aequation under steady state (Re = 0). Because of the existence of sharp channel corners, both the electric lines and the streamlines of the pressure driven flow are different from the streamlines of the pure elongational flow near the channel wall. However, we can still see that the electric lines are closer to the streamlines of the pure elongational flow than the streamlines generated by the hydrodynamic pressure. The dissimilitude of streamline and electric line become more significant at regions far away from the stagnation point. At a certain y position inside the cross region, the dissimilitude exhibits at a certain x position. For example, at y = 67 µm, the electric lines agree closely with the streamlines of the pure elongational flow up to around x = 90 µm, which supports our experimental results for the measurements of DNA velocity (shown in Figure 3.7).
3.3.1.2 Flow-induced molecular dynamics of DNA
A dilute solution (~ 0.03 µg/ml, about 10-3 of the overlapping concentration) of λ-
DNA (48.5 kbp) in Tris-EDTA buffer was used and labeled with fluorescent YOYO-1 dye at a dye-base pair ratio of 1:5. In these conditions, λ-DNA has a gyration radius of about 0.73 µm in equilibrium; after being fully stretched, it has a contour length of 16.3
49 µm (unstained, Bustamante et al, 1994) and about 21 µm (dye: base pair = 1:5, Perkins et al, 1997).
Figure 3.7 shows the measured velocity distribution of DNA molecules in the spanwise (x) and vertical (z) directions. To minimize the influence of stretched polymer chain to the velocity of DNA center of mass, no sucrose was added in this specific case so that DNA molecules were kept their super-coiled structure. At y = 67 µm (inside the intersection), the velocity of DNA molecules was nearly the same in the spanwise direction up to x = 40 µm and then gradually increased afterwards. The velocity distribution can then be calculated using equation (3-1), which is represented by the solid line in Figure 3.7. It shows a very well agreement with experimental measurements up to around x = 90 µm, indicating that a nearly pure elongational flow field can be generated by electrokinetic forces. At y = 225 µm (outside the intersection), a nearly constant velocity of DNA molecules was observed in the spanwise direction up to x = 100 µm but then gradually decreased afterwards. Since DNA molecules have left the elongational flow field in the intersection area and are inside the straight channel, a more plug-like velocity profile is expected. However, the observed velocity of the DNA molecules near the channel wall decreased, which mainly result from the increase of viscous drag due to the hydrodynamic interactions between the DNA molecules and the solid wall. For the same reason, obvious depletion of DNA molecules also was observed in the near wall region (Fang, L., et al, 2005). More details about the hydrodynamic interactions will be discussed in Chapter 5.
50 In the vertical direction, the velocities of DNA molecules at selected x and y positions with z locations ranging from 20 to 80 µm from the bottom of the channel are fairly uniform. These results verify that a fairly homogeneous, two-dimensional (planar) flow field can be generated by the electrokinetics-induced flows. The velocity gradient is largely minimized in both the spanwise direction and the vertical direction. This advantage overcomes the limitation of other DNA coil-stretched transition study in the microscale cross-channels by hydrodynamics-driven flows, where the observation has to be done in the region very close to the centerline. Within this electrokientic-induced flow, such studies can be focused on almost any planes in the vertical direction, as long as not too close to the wall.
According to the DNA stretching theories in elongational flows, “electrokinetics- induced” Deborah number (De) is one of the major criteria to control the coil-stretch conditions, which is defined by the following equation:
De = λε& (3-3) where λ is the maximum relaxation time for DNA molecules in the certain solution and
ε& is the extension rate. In principle, stretching only happens when the value of Deborah number is larger than half of the unit ( De > 0.5 ).
To identify the extension rate, x component of the velocity of DNA mass center were measured and plotted versus x position, as shown in Figure 3.8. The x component was chosen because of less stretching conformation when DNA entering the cross-slot (in x direction) than those moments they were leaving (in y direction). Data from all four quarters were taken and were plotted in the first quarter by taking their absolute value. A
51 straight line was obtained when fitting all of those data and the extension rate, ε& , is given by the slope of the line, which is equal to 0.78 (1/s) in the experimental conditions.
Theoretically, the relaxation times of DNA molecules are linearly proportional to the viscosity of solution (Chapman et al, 1969). Using the experimental data of λ-phage
DNA molecules published by other researchers (e.g., Perkins et al, 1997, Smith et al,
1999 and Babcock et al, 2000), a linear relationship is established and given as (Fang, L. et al, 2005)
λ(s) = 0.094η s (cP) (3-4)
Considering that the viscosity of DNA solution used in our dynamic stretching experiments is ~30 cP, the relaxation time is 2.82 s. Then, the “electrokinetics-induced”
Deborah number (De) could be calculated and it was about 2.2, which is way larger than the critical value (De=0.5) for molecule stretching.
Two major pre-deformation exists in pressure-driven elongational flow: one is the shear-induced pre-deformation because of its non-uniform velocity profile and the other is the entry deformation upon the departure of the reservoirs. The first pre-deformation was eliminated by studying at the startup of the flows and the second could be much reduced by using long entry channel on the inlet. In our cases, such pre-deformation, in principle, could be largely avoided. Long entry channel was also used and more uniform velocity profile is obtained by applying electrokinetic-induced flows. DNA molecules experience the same and long residence time prior to entering the pure elongational flow region in the cross-slot geometry. However, several initial conformations were still
52 observed when entering the imaging domain, which is believed to be induced by
Brownian motion.
Beside the various initial conformations resulting from pre-deformation, residence time is another important factor controlling the degree of DNA molecules stretching.
Even though uniform extension rate exists inside the cross region, DNA molecules following in different streamlines have different residence time. With a similar initial conformation (a semi-dumbbell shape in this case), the DNA molecule having a longer residence time can be stretched longer than that having a shorter residence time as shown in Figure 3.9a. On the other hand, two DNA molecules with different initial conformations (a curled and a partially stretched shape in this case) may end with a similar extent of stretching as shown in Figure 3.9b, even though the residence times are substantially different. Therefore, it is also true that both the initial conformation and the residence time of DNA molecules play important roles in DNA stretching inside the electrokinetics-induced stagnation flow, which has been verified inside the pressure- driven elongational flow field (Perkins, T. T., et al, 1997). All of those various conformations, including initial and final conformations, were classified into several big groups, like super-coiled, kinked, half-dumbbell, dumbbell, curled, stretched and so on
(shown in Figure 3.10).
Brownian dynamics (BD) simulation was also applied to make a comparison with the experimental results. Worm-like chain model (Larson, R. G., et al, 1999; Hur, J. S., et al, 2000; Panwar, A. S., et al, 2003) was applied to simulate the coil-stretch transition of
DNA molecules. The governing equation for the movement of the i-th bead is given as follows (Panwar, A. S., et al, 2003):
53 Brownian S S ξ r&i = ξ vi + Fi + (Fi − Fi−1) + qEi (3-5)
where ri is the i-th bead’s coordinates, r&i is its derivate with respect to time, ξ is the
Brownian drag coefficient, v i is the flow velocity generated by electroosmosis, Fi is the
S S Brownian force, Fi − Fi−1 is the total spring force for the i-th bead, and qEi is the electric force. Following the examples of ((Larson, R. G., et al, 1999; Hur, J. S., et al,
2000), we used a 20-bead chain (20 beads and 19 springs) with persistence length (half of one Kuhn step length) of 0.066 µm to simulate the movement of λ-DNA.
Because the counterions move in a direction opposite to the DNA molecules and screen the hydrodynamic interactions over a distance larger than the Debye length (only several nanometers, Viovy, J.-L., 2000), we can neglect the hydrodynamic interactions in equation (3-5), when more attentions is paid to regions away from the wall (i.e., >10 µm).
Since the flow cell is made of PMMA, which has a surface with a very low charge density in the buffer solution, the electroosmotic effect is much weaker than the electrophoretic effect. Since the flow generated by electroosmosis is similar to the electric field (Cumming, E. B., et al, 2000; Dutta, P., et al, 2002), we can remove the electroosmosis in equation (3-5) by simply combining this effect into the “effective”
charge density q of the DNA molecule. In the simulation, r&C ≈ qEC /ξ , where r&C is the
velocity for the center of mass of DNA and EC is the electric field at rC (the location for the center of mass of DNA). By fitting the center of mass velocity of DNA with the electric field, we got q /ξ = −1.16×10−8 m2 /(V ⋅ s ) and the value was used in the
Brownian dynamics simulation.
54 Several cases of DNA stretching were made comparison between experiments and simulation results. The coarse-grain molecular simulation by using corresponding initial
DNA shapes and the calculation results were placed side-by-side with the experimental observations (shown in Figure 3.9a and 9b). They agree qualitatively with each other.
3.3.2 Shear and recirculation flows
3.3.2.1 Shear flow and recirculation flow generation
With the same single cross-slot geometry, flow patterns other than pure elongational flow can be generated. By applying similar criteria described by Babcock
(Babcock, H. P., et al, 2003), the flow type can be characterized by a factor Λ defined as
|| L || − || Ω || Λ = || L || + || Ω || (3-6)
T where L = ∇u p + (∇u p ) ) / 2 is the symmetric or extensional part of the particle velocity
T gradient ∇u p , Ω = ∇u p − (∇u p ) ) / 2 is the anti-symmetric or rotational part, and is the Frobenius norm of a tensor. The value of Λ falls between –1 and 1. When Λ = −1,
0, or 1, the flow is pure rotational, simple shear or pure extensional, respectively. When
−1 < Λ < 0 , it is a rotational-dominated mixed flow and when 0 < Λ = 1, an extensional- dominated mixed flow.
For pressure-driven flows in cross-slot devices, such as “four-roll” mill, those flow patterns are controlled by adjusting the rotation speed set of the four rolls (so does the flow rate for each arm). In principles, electrokinetic-induced flows can also form those flows. By using different surface charge arrangements on flow arm surfaces and
55 various electric bias configurations in liquid storage reservoirs, elongational, shear and rotational flows also can be carried out. The schemes for different setup are shown in
Figure 3.4. Here, two arms of a single cross-slot in the diagonal position are considered as a pair. If zeta potentials of these two pairs are identical (both the magnitude and sign), pure elongational flow is presented in the cross region. Flow patterns other than pure elongational flow will be presented as long as the zeta potentials of these two pairs are different. The other special case happens when the magnitudes of zeta potential of two pairs of arms are identical but carry different sign of charge. In this case, pure rotation flow is generated. Other cases will be generated upon adjusting the zeta potential of the two pairs of four arms.
3.3.2.2 Electrokinetic interaction analysis
All of the above discussions are base on the assumption that no influence results from species existing in the fluid. In practice, the species used in the flows carry more or less charge. This will add additional electrophoretic component to the specie velocity when driven by electroosmotic flow. The velocity of a massless charged particle in the
2D electrokinetic field can be expressed as u p = E0 (µ EOu + µ EP E ) , where E0 is the characteristic electric field magnitude, u = (u,v ) is the dimensionless electroosmotic
εζ velocity, E is the dimensionless electric field vector, µ = s is the EO mobility, and EO η
q µ = is the EP mobility. Here, ε is the permittivity, ζ is the zeta potential of the EP ξ s
56 channel surface, η is the viscosity of the solution, q is the charge density of the electrolyte, and ξ is the drag coefficient.
Figure 3.11a shows the schematic of the cross-slot device. Here, one pair arms in
EO the diagonal are negatively charged with the surface charge density µ− , and the other
EO pair positively charged with the surface charge density µ+ . To help explain the interaction of electrophoresis and electroosmosis, two new parameters are defined here:
χ and λ . χ is defined as the ratio of the surface charge density of these two pairs of four
EO EO arms in the cross slot ( χ = µ− /).µ + λ is the ratio of EP mobility to the characteristic
EO surface EO mobility ( λ = µ EP / µ EO ), where µEO = − µ− . Different values of these two parameters present different flow patterns generated in this single cross slots.
Figure 3.11a assumes that the value of χ is equal to -1, that is, EO mobility of the
EO EO two surfaces satisfy µ− =0− µ + < . Pure rotational flow in the center of the cross slot
will present when the particle EP mobility, µ EP , is negligible compared with the
characteristic surface EO mobility, µEO , which means λ =0. When µ EP becomes non- negligible, the EO-dominated rotational flow is skewed (Figure. 3.11b). With EP mobility of particles are larger than the value of surface EO mobility, that is, λ ≤ −1.0 when negatively charged particles are used), the flow becomes mixed-shear flow with extensional component becomes dominating when λ keeping increasing (Shown in
Figure 3.11c). Experimentally, that will happen in most of the cases since the tracer particles usually carry more or less negative charges. In the case shown in Figure 3.11d, the EP mobility of liposome nanoparticles is -2.75 × 10-8 (m2/sV) while the four arms are
57 constructed by two pieces of No. 1 glass slide and two pieces of APTES-modified No. 1 glass slide with dimensions 25 x 25 x 1 mm, whose EO mobilities are -3 × 10-8 and +4 ×
-8 2 EO EO 10 (m /sV), respectively. This gives χ = µ− / µ+ = −1. 33 and λ =-0.92, which is
EO EO pretty close to the simulation case (χ = µ− / µ+ = − 1 and λ = −1) shown in Figure
3.11c. The actual flow patterns match pretty well with the simulation results. Liposome particles are used here to minimize the complex dynamic adsorption when two different sign of charge surface are involved in the flow channels, due to their grafted PEG functional groups.
As mentioned above, the rotation flow pattern can be stretched because of the different ratio of the EP mobility and the surface EO mobility (Shown in Figure 3.11a and 11b). Similar situations can also be generated by controlling χ , the ratio of the EO mobility of the two pairs of arms in the cross slot (shown in Figure 3_12a and 12b). With the increase of the absolute value of χ , the stretched rotation flow patterns could be split into two symmetric small circulation flows at the two corners in the diagonal position, as shown in Figure 3.12b. Actually, this is the common case observed in the real experiments. Figure 3.12c is the simulated flow pattern with the consideration of geometric “offset”, from misalignment of four pieces of glass. It agrees fairly well with the experimental result shown in Figure 3.12d. The “offset” changes the vortex size but not the flow pattern.
58 3.4 FIVE CROSS-SLOT ASSISTED MICROFLUIDIC PLATFORM
As discussed in the precious section, it is not easy to obtain favorable surface charge density in practice by current available various surface modification methods, such as multiphase laminar flow patterning and surface coating, in electrokinetic-induced shear and rotational flows. Moreover, the movement of electrolytes in aqueous fluid is strongly affected by the interactions between electroosmosis (EO) and electrophoresis
(EP), in particular when the electrophoretic mobility of a particle is greater than or equal to the electroosmotic mobility. The flow pattern is also highly dependent on the surface charge pattern of channel walls. Since the surface charge varies with solution properties
(i.e., pH values and ion concentration), it is difficult to control the surface charge to an exact certain level. In addition, the electrolyte adsorption on the oppositely charged surface is another major concern when performing those different flows. Although special treatments such as coating polyethylene glycol (PEG) on the particle surface can minimize this problem, it is impractical in many chemical and biological applications.
In this section, a new five cross-slots design is developed, which can generate electrokinetically modulated various flow patterns mentioned above by simply changing the voltage inputs without the need of surface patterning and coating. This five cross-slots design can be considered as the extension of single cross-slot: if four single cross slots are connected with one corresponding arm from each single cross-slot merging with the other three, that will give you the initial five cross-slot design. As shown in Figure 3.2, the fluidic network consists of five cross-slots with 12 electrodes (A1 to A12) placed in 12 reservoirs. A DC power supply is used to provide different voltage inputs. For
59 demonstration, the channel width D is chosen as 250 µm or 1 mm. The length of the 12 shorter arms is 1.5D, and the length of the 4 longer arms is 2.5D. The relative lengths of the arms could be adjusted. Since most solids used in microfluidics, such as glass and plastics, possess negative static charge, it is assumed that all channel walls are negatively charged. The dielectrophoresis is insignificant due to the low voltage drop and small electric gradient, and is confirmed by the numerical simulation.
3.4.1 Extensional flow and shear flow
In this five cross-slot design, there are several ways to generate the pure extensional and mixed-shear flow. The simplest method is to use only 4 electrodes and ground the other 8 electrodes. For example, by using only electrodes A1, A6, A7, and A12, the network is essentially a symmetric single cross-slot flow device that can generate
either the EO dominated (if µ EO > µ EP ) or the EP dominated (if µ EO < µ EP ) extensional flow. These two flows have similar streamlines, but opposite flow directions. By using only electrodes A1, A2, A3, and A12, we can obtain an asymmetric single cross-slot flow device. The flow is either EO or EP dominated mixed-shear depending on the relative
magnitude of µ EO and µ EP . They obviously have opposite flow directions.
A more sophisticated design is shown in Figure 3.13a, where the extensional flow is generated in the middle cross-slot and the mixed-shear flow in the four corner cross- slots simultaneously. The streamlines of the flow pattern are shown in Figure 3.13b and
13c by compounding the video graphs of at least 200 frames. An elongational flow was observed in the center cross and mixed-shear flows were observed at all four other side crosses, which agree well with the simulation prediction.
60 Similar to the single cross slot section, the conformation dynamics of single flexible polymer molecules also was studied in this five cross-slot device in extensional and mixed-shear flows. The same experimental conditions are used here. In the center cross, the electrokinetic flow is an EO dominated extensional flow. The
“electrokinetically-induced” Deborah number (De) was calculated to be ~1.4, which is larger than the critical value of 0.5 for molecule stretching. DNA chains are gradually stretched when they approach the center of the cross, the stagnation point. As shown in
Figure 3.14a, DNA molecule A, which is closer to the centerline, was stretched to a longer length (close to 9 µm) compared to DNA molecule B, which is away from the centerline, although both experienced almost the same length of residence time in the observed domain. This is because DNA A has an unfolded initial conformation, while
DNA B is initially in the folded shape, making it more difficult to be stretched.
In the mixed-shear flow regions, sheared DNA molecules experienced different configurational changes in both size and orientation, depending on their initial conformations and flow paths. As shown in Figure 3.14b, DNA molecules C and D can only be stretched to 3-5 µm with the corresponding Weissenberg number equal to ~ 1.7 to
2.1, respectively. Considering the critical value of the Weissenberg number to stretch the
DNA molecules is 1.0, it is quite reasonable to get only 15%-25% extension of the contour length of the DNA molecules in this mixed shear flow.
Again, the same method of Brownian dynamics (BD) simulation was carried out to describe the coil-stretch transition of DNA molecules in this elongational and shear flows. For both extensional flow and mixed-shear flows, the calculated configurational
61 changes from our coarse-grain molecular simulation could match the experimental results well with appropriate initial DNA conformations as shown in Figures 3.14a and 3.14b.
3.4.2 Rotational flow
The electrophoretic flow is purely extensional because ∇E is symmetric, considering that ∇ ⋅ E = 0 and ∇ × E = 0 . The rotation of particles can only come from the electroosmotic flow. To generate a rotational flow independent of the particle charge density, the electric field E has to be suppressed and the rotational part in the
electroosmotic flow u EO has to be enhanced. This can be easily accomplished in this five cross-slot device. By setting the voltage inputs as shown in Figure 3.15a, a rotational flow forms in the center of the five cross-slot due to the electroosmotically-driven secondary flow. Because of the symmetric geometry and imposed electric potentials with respect to the center of the five-cross network, the electroosmotic flows from four outer cross-slots impinge on one another at the center of the middle cross-slot. Thus, the electroosmotic flow in the middle cross-slot has to be a rotation type as it is the only way
to satisfy the continuity condition∇ ⋅u EO = 0 . Furthermore, the electric lines generated from the symmetry of geometry and imposed electric potentials intersect with one another in the middle cross-slot, thus the electric field there is greatly suppressed. In fact, the electric field is zero at the center of the five-cross network.
This predicted rotational flow is also successfully demonstrated experimentally using the same platform described above for the mixed-shear and extensional flow. The only difference is in the arrangement of voltage inputs, as shown in Figure 3.15a. Here, the actual voltage (in Volts) needs to be multiplied by a factor of 50. The rotational flow
62 was observed at the center of the device for both polystyrene microspheres and λ-DNA molecules, as shown in Figure 3.15b and 15c. The rotational direction is in accordance with the direction of the simulated streamlines. As opposed to both shear and extensional flows, the rotational flow does not generate any strong gradient difference. Therefore, most DNA molecules stay in their coiled configuration and the coil-stretch transition is rarely observed in this case.
The rotational direction is independent of the charge type and charge density of the particles. However, the size of the vortex varies. To investigate the effect of EP mobility of microspheres on the vortex size, two types of polystyrene microspheres were used: the 3 µm plain polystyrene microspheres used in all other experiments and 2 µm polystyrene microspheres with multiple carboxyl groups on the surface. The measured
-8 -8 2 µ EP for these two microspheres in DI water are –0.78×10 and –7.02×10 (m /sV), respectively. Different sizes of vortex were observed when using different microspheres.
The computed vortex size (in diameter and non-dimensionalized based on the channel width D) is plotted as a function of the relative magnitude of EP vs. EO mobility,
λ = µ EP / µ EO as shown in Figure 3.16. The actual experimental points were also added for comparison. The observed vortex sizes agree quite well with the simulation prediction.
3.5 SUMMARY
Single or multiple cross-slot microfluidic platforms were designed to generate elongational, shear and rotational flows by using different surface charge arrangements
63 on flow arm surfaces and various electric bias configurations in liquid storage reservoirs.
Fairly homogeneous, two-dimensional flows can be generated readily by electrokinetics in low-viscous fluids (e.g., aqueous solutions). Charged polystyrene microspheres were used to identify the flow characteristics, and the conformational evolution of single flexible molecules (e.g. λ-DNA) was investigated under those three basic flows. Different cases of coupled electrophoresis and electroosmosis also were explored, considering the complicated interactions between biomolecules and charged surface of channel walls.
Except elongational flow, other electrokinetic-induced flows rely so heavily on a spatially non-uniform surface charge distribution on channel walls that it becomes too hard to control and maintain the flow patterns while actual adsorption happens between oppositely charged species and channel surfaces. By extending to a five cross-slots design, the situation is significantly improved because of the independence of flow patterns to the surface properties, especially for shear and rotational flows.
Even though various degree of deformation is largely eliminated in our electrokinetic-induced stagnation flows, a variety of DNA conformations still exhibited within cross-slot geometry. Two natural drawbacks lead to such diversity: one is the different residence time for each streamline and the other is that most of the extended stretching happens in the near-stagnation point where Brownian motion is also significant. New designs, which can provide uniform and controllable DNA stretching, will be developed and discussed in the following chapters.
64
(a)
(b)
Figure 3.1 Optical image of single cross-slot device (a) and the schematic of the cross- slot dimensions and the arrangement of electrodes (b) (drawn not in scale). The sale bar represents 1 cm.
65
(a)
(b)
Figure 3.2 Optical image of five cross-slots device (a) and the schematic of the five cross- slots dimensions and the arrangement of electrodes (b) (drawn in scale). The sale bar represents 500 µm.
66
(a) 4 5 3 2 6 1
495 nm 510 nm 1. Power Control 2. Power Supplies/Relay 7 3. Moving Stage 4. Chip holder 8 5. Microchip 6. Object lens 9 7. Epifluorescent filter cube 8. Mercury light source 9. Detector/Cooled CCD Camera 10. Computer/MetaMorph software 10
(b)
Figure 3.3 Experimental setup: the schematic drawing (a) and photography (b).
67
(a) (b) + (c) + _ _+ _ _ _ _ _ + + _ + _ + _+ _ + + _ + _ + _+ _ _ _ _ _ + + _ + + _ _ _ + _ _ ++ _ + _ _+ ______+ + ______+ ______+ _ + ______+ + + +_ +_ +_ +_ _+ _ + _ + + + + + + + + + + + _ _ + + + + + + _ _ + + + + + + _ + + + + + + + + + +_ _+ _+ _+ _+ + + + + + ______+ + _ _ _ _ _ + ______+ _ _ + + _ _ + _+ _ + + + + + + _ + + _ _ + _ + _ + _ + + _ _ + _ + _ + _ _ + + _ + _ + _ + ______+ + _ _ + _+ _ + ++ + + +
68 + + +
Pure extensional flow Mixed shear flow Pure rotational flow
Figure 3.4 Schematic of surface arrangements for the generation of different flow patterns by electrokinetics in single cross-slot: pure elongational flow (a), mixed shear flow (b), and pure rotational flow (c).
68
(a) (b)
(c) (d)
Y
X, µm 0 X
Figure 3.5 An elongational flow in a single cross-slot: the streamlines by compounding image of fluorescence microspheres trace driven by electrokinetics (a), by simulation (b) and velocity vector field of experimental data (c) and simulation (d).
69
Figure 3.6 Comparison of the electric lines, the streamlines of a pressure-driven flow, and the streamlines of a pure elongational flow.
70
350
300
250 m/s µ
200 y = 225 µm
150
100
of DNA, Velocity y = 67 µm 50
0 0 20406080100120 x, µm y = 225 µm, z = 40 µm y = 225 µm, z = 60 µm y = 225 µm, z = 80 µm y = 67 µm, z = 40 µm y = 67 µm, z = 60 µm y = 67 µm, z = 80 µm y = 67 µm, z = 50 µm y = 67 µm, z = 70 µm
Figure 3.7 Velocity distributions of DNA molecules at various locations in the spanwise (x) and vertical (z) directions. z = 0 µm is at the bottom of the channel.
71
50
40
30
m/s µ Vx = 0.78x lVxl, 20
10
0 0 10203040506070
lxl, µm
Figure 3.8 The plot of x component of the velocity of DNA mass center versus x position with the stagnation point as the origin. Data are taken their absolute value so that they all fall in the first quarter of the cross-slot, with different symbols representing data from different quarters. The extension rate in the pure elongational flow, ε& , is given by the slope of the fitting curve, which is equal to 0.78 (1/s) in the experimental conditions.
72
(a)
Continued
Figure 3.9 Experiment and simulation of the movements of two DNAs in the intersection region with similar initial conformation but different residence time (a), with different initial conformation and residence time (b).
73
Figure 3.9 continued
(b)
74
(a) (b) (c) (d)
(e) (f) (g) (h)
Figure 3.10 Different initial configurations of λ-DNA molecules in an elongational flow driven by electrokinetics within single cross-slot: super-coiled (a), curled (b), half- dumbbell (c and d), kinked (e), dumbbell (f and g) and stretched (h). The scale bar represents 5 µm.
75
(a) + R - D + 4 - C + ------+ + + + +
R R1 2
- - - - - + + + + + - + A - + B R - 3 +
(b)
Continued
Figure 3.11 Flow pattern evolution from strong electrokinetic interactions for different particle charge density or λ (= µEP / µEO ) : λ is equal to (a) 0, (b) -0.2, (c) -1.0 from simulation and (d) -1.0 from experiment (two asymptotes are outlined near the intersect area for clarity). Scale bar represents 200 µm.
76
Figure 3.11 continued
(c)
(d)
77
(a)
(b)
Continued
Figure 3.12 Flow pattern evolution from electrokinetic interaction for different wall surface zeta potential for positively charged arm pair and the negatively charged arm pair
EO EO or χ (= µ− / µ+ ) : χ is equal to (a) -2, (b) -5 with λ = −0.05 , and is fixed. (c) -1 from simulation and (d) -1 from experiment withλ = −1, and is fixed. The scale bar represents 200 µm. 78
Figure 3.12 continued
(c)
(d)
79
0
(a)
+2 +1
-1 -2
0 0
-2 -1
+1 +2
0
Continued
Figure 3.13 Flow pattern generation of both elongational flow (in the center cross) and mixed shear flow (in the side crosses) in a five cross-slot design: the design and arrangement of electrodes (a), experimental result of the compounded pathlines of PS microspheres, representing an elongational flow in the center cross (b), and mixed shear flows in the side crosses (c). The scale bar represents 200 µm.
80
Figure 3.13 continued
(b)
(c)
81
82
(a) (b)
Figure 3.14 Experimental and simulation results of the movements of DNA molecules in the five cross-slot microfluidic platform in an elongational flow (a), in a mixed-shear flow (b). Two DNA molecules experience different residence time.
82
0 (a)
-1 +1
+1 -1
0 0
-1 +1
+1 -1
0
Continued
Figure 3.15 Flow pattern generation of rotational flow (in the center cross) in a five cross- slot design: the design and arrangement of electrodes (a), experimental result of the compounded pathlines of PS microspheres, representing an rotational flow in the center cross (b), and microscopy image of λ-DNA (c). The scale bars in (b) and (c) represents 200 µm, and 20 µm, respectively.
83
Figure 3.15 continued
(b)
(c)
84
λ = -2 λ = -0.2
Figure 3.16 Dimensionless vortex size versus the ratio of EP mobilty to EO mobility
λ (= µEP / µEO ) and two rotational flow patterns forλ = −0.2 and - 2 .
85 CHAPTER 4
POLYMER NANONOZZLE ARRAY BY SACRIFICIAL TEMPLATE IMPRINT (STI)
4.1 INTRODUCTION
In this chapter, a low-cost process, called Sacrificial Template Imprinting (STI), is introduced in detail, which is developed for massively production of desirable micro-
/nanofeatures (e.g., high aspect ratio nanostructures). As the demonstration, this technique is applied to the fabrication of a polymer nanonozzle array with high convergence ratios. Nanonozzle arrays fabricated by STI can provide both converging and diverging flow patterns, which are more beneficial than straight nanochannels in controlled delivery of drugs and genes. Basically, a polymer sacrificial template, instead of silicon, glass or metal mold insert, is used, which can be massively produced and easily removed after molding. These advantages make this technique to be low cost and high throughput. It also avoids structure damage or defects generation during the de- molding process without any mold release agents (due to the contamination for biological applications). This is very important in the fabrication of high aspect ratio nanofeatures.
In addition, since those sacrificial templates can be easily mass-produced with high quality, this STI method also solves the problems of high throughput, yield and massively
86 parallel production. In this process, no expensive lithography equipments or clean room facility are necessary.
The mother master is conically shaped nanotip array fabricated from coherent fiber-optic bundles by differential wet etching. A two-step replication is then applied to produce sacrificial templates using a PDMS female mold for transition. A thin layer of polymer with open channels is formed by controlling the spin conditions to ensure that the polymer layer thickness is slightly less than the height of the nanotips on the sacrificial template. The nanonozzle array is released by dissolving the sacrificial template in water. Each nanonozzle is 3µm high with the channel diameter on the sharp end as small as 80 nm.
Because of meniscus effect, nanonozzles have a volcano-shape aperture on the small end. Two types of different surface planization techniques (wet and dry planization) are introduced how to remove such features if they are not desired. This also provides other ways to extend this sacrificial template imprint. Other derivatives of sacrificial template imprinting (STI) are also discussed.
Limited by the geometry of nanotips on current sacrificial template, the nanonozzle layer produced is very thin (< 5 µm). Its nanostrucuture may not stand in aqueous working environment (e.g., biomedical applications) for long term due to structural and dimensional instability. In addition, it is difficult to obtain nanonozzles smaller than 50 nm in diameter. In the last session of this chapter, a new approach, called dynamic assembly, is introduced to solve this problem by growing silica on the internal surface of nanonozzles. This was realized via electrokinetic-induced dynamic surface
87 reactions. In conjuction with surface modification and silica synthesis on the channel surface, the channel size can be further regulated and the polymer structure can be reinforced. This dynamic assembly approach can also add other functionalities to the surface of the nanonozzles.
4.2 SACRIFICIAL TEMPLATE IMPRINT
4.2.1 Materials
Optic fiber bundles (Image guide, IGN-037/10) were purchased from Sumitomo
Electric, Inc. Buffered oxide etchant (BOE) was purchase from Ashland Chemical, Inc with a volume ratio of (NH4) F to HF equal to 7. The kit of PDMS precursor and curing agent (Sylgard 184, 10:1 weight ratio) was purchased from Dow Corning. Poly(vinyl
alcohol, PVA, M w = 30,000-70,000), Chitosan (Poly(D-glucosamine), Mr~150,000) and
glucose were purchased from Sigma-Aldrich. Poly (menthe methylacrylate, M w =
350,000) was purchased from Polysciences, Inc and dissolved in toluene with the concentration of 10-15 wt% and filtered with 1 µm PTFE filters.
4.2.2 Process description
The schematic of STI process is shown in Figure 4.1. Masters with an array of conically shaped tips were fabricated on the distal faces of a coherent fiber-optic bundle by differential wet etching (Pangaribuan, T., et al, 1992; Dam, T. H., et al, 1999). The conical shape tips were gradually formed in the buffered oxide etchant (BOE) due to the different solubility of the etching products ((NH4)2XF6, X=Si or Ge) from the core and
88 cladding materials. A two-step replication was then applied to produce polymer sacrificial templates. A PDMS mold was first served as the transition mold to generate inverted conical nanowells from the fiber-optic master by replica molding. An aqueous solution of a water soluble polymer was cast on the PDMS mold. After drying, the sacrificial template was peeled off and attached onto some flat substrate (i.e., a glass slide). A polymer solution (e.g.10-20 wt% PMMA in toluene) or resin (e.g. PDMS) was spun on the sacrificial template. A thin polymer layer with open channels was formed by controlling the spin conditions to ensure that the film thickness was slightly less than the height of the nanotips on the sacrificial template. After curing the resin or drying the solution, the nanonozzle array was released by dissolving the sacrificial template in water.
This sacrificial template imprinting (STI) technique is not limited to produce just polymer nanonozzle array but could become an attractive universal technique in massive fabrication of other micro/nano features, especially those with high-density or high aspect ratios or both without using any mold release agents.
4.2.3 Fabrication of optic-fiber nanotip
The coherent fiber-optic bundle (Image guide, IGN-037/10) has a diameter of 333
µm, composed of 10,000 individual optical fibers (the fiber density is ~ 107/cm2). Each individual optical fiber has a diameter of 3-micron, composed of a GeO2-doped silica core and fluorine-doped silica cladding. The fiber diameter can be reduced by simultaneously pulling and heating its distal end with a standard glass capillary tube pipette puller (Narishige Model PE-2000G, Sea Cliff, NY). This can further increase the
89 tip density to ~109/cm2. Fourteen bundles were grouped together to form a surface area near 3 mm2.
The image guide was embedded in an acrylic block (10mm×10mm×8mm) with one face exposed. The distal face was then polished successively with 30-, 15-, 3-, and
0.3 µm lapping films (General Fiber Optics, Fairfield, NJ). Sonication in DI water might be used, if necessary, to remove residual polishing material. Then the distal face of the image guide was immersed perpendicularly into a 100 ml buffered oxide etchant (HF to
(NH4) F equal to 1:7, v/v) for a set period of time at room temperature. CAUTION:
Hydrofluoric acid is extremely corrosive. The conical shape tips were gradually formed in the buffered oxide etchant (BOE). Finally, the whole image guide was rinsed and sonicated thoroughly in DI water before analysis. This helps to stop the etching process and remove the partially dissolved materials that might stick to the tip surface.
Four reactions are involved in this etching process.
XO2 + 6HF ÆH2XF6 + 2H2O (4-1)
H2XF6 + 2NH3 Æ (NH4)2XF6 (X=Si or Ge) (4-2)
The generated (NH4)2SiF6 and (NH4)2GeF6 are absorbed on the surface of the fiber and need to be dissolved away. Their different dissolving rates determine the relative etching rates between the core and the cladding. The etching process occurs in both radical and along-the-fiber directions. Eventually, the etching process reaches a steady state and excess etching time has no effect on the tip geometry. The final tip height, cone angle, and tip formation time can be expressed as
h = d1(S2−S1) tip 2S1 (4-3)
90 θ = 2tan−1( S1 ) S2−S1 (4-4)
t = d1 min 2S1 (4-5) where d1 is the diameter of individual optic fiber, Si is the dissolving rate, and the subscripts 1 and 2 denote core and cladding, respectively. The dimensions and geometry of the tips can be finely tuned through varying the differential etching rates (Puygranier,
B. A. F., et al, 2000). Desirable shape of nanotips can be obtained by adjusting factors, such as the doping ratio in pure silica, the initial dimensions of the fiber, the composition of etching solution (the volume ratio of NH4F to HF), the etching time, and temperature.
The nanotip formation procedure was observed by taking SEM images at certain etching stages (shown in Figures 4.2a, 2b, and 2c). Through these images, the minimum etching time to achieve the conically shaped probe can be determined. With equations 4-3 to 4-5, the etching rate for the core and the cladding materials can be calculated. Most etching processes were carried out at 25oC in buffered oxide etchant (BOE) with a volume ratio of (NH4) F to HF equal to 7. Under these conditions, the minimum etching time is 30 min. Those optical tips are around 5 µm in height with the sharp end around
50-100 nm in diameter and the conical top angle around 24o (shown in Figure 4.3a).
Through calculations, the etching rate for the core and the cladding materials are 2 µm/hr and 10 µm/hr, respectively.
The etching conditions, such as BOE composition and etching parameters (e.g., etching temperature) are two major ways which can be optimized to obtained desirable tip geometry. For this particular case, higher tips with sharper distal end are desirable.
91 From the model and equations 4-3 to 4-5, it can be found the geometry of nanotips is determined by the etching rate of the core material and the etching rate difference between the core and the cladding materials.
Like most of other chemical reactions, the effect of temperature is held an exponential relationship with the etching rate. According to equations 4-3 to 4-5, temperature affects the value of both S1 and (S1-S2). The temperature effect was plotted in
Figure 4.2d with y axis as cot (θ/2), equal to (S2- S1)/ S1. The larger the value of (S2- S1)/
S1, the higher and sharper the tips are. In Figure 4.2d, the tips became shorter and their conical angle became larger with the increasing of etching temperature. But different temperature doesn’t significantly amplify the etching rate difference between the cladding and core materials. Thus, optimal temperature individually is hard to achieve higher aspect ratio tips as expected.
Although optical nanotip array could be used directly as the mold itself, our recent experience indicated that they would not last in mass-production and probably face the same de-molding issues mentioned above. Therefore, a polymer sacrificial template
(Figure 4.3b) was used instead.
4.2.4 Fabrication of sacrificial template
A two-step replication was applied to produce polymer sacrificial templates. A
PDMS mold was prepared first as a transition mold with inverted conical nanowells from the fiber-optic master. An aqueous solution of a water soluble polymer was cast onto the
PDMS transition mold and dried at ambient conditions. Most sacrificial templates were made of polyvinyl alcohol aqueous solutions (5-10 wt%). Other materials with similar
92 characteristics (i.e., 1wt% Chitosan/1M acetic acid, Glucose aqueous solutions, 40 wt %) were also successfully applied to produce high quality sacrificial templates.
The two-step replication entails easy mass-production of sacrificial templates with high-fidelity replicated features except that the tip end is not as sharp as those on the original fiber-optic master (see Figures 4.3a and 3b, 3c). Since a two-step replication is involved, the replication accuracy of sacrificial template depends on the wetting between the mold and the casting fluid (i.e. between the optic fiber nanotips and the PDMS resin, or the cured PDMS and the aqueous polymer solution) and dimension changes during solution dry or resin cure. PDMS can spread out on the optic-fiber nanotip surface (which is mainly composed of glass) to ensure a good replicate quality. Thus, the replication inaccuracy mainly corresponds to the filling of fluid in PDMS nanowells and the drying process afterwards. PVA possesses characteristics of a surfactant due to the presence of a large number of hydroxyl groups. Its aqueous solution can easily wet the hydrophobic surface and completely fill inverted conical nanowells on the PDMS mold without tracking air. After drying, the hydrophilic PVA can be easily peeled off from the hydrophobic PDMS mold.
To verify this hypothesis, different solutions were tested to produce sacrificial templates with the same PDMS transition mold, as shown in Figures 4.3b-4f. For those materials (i.e., PVA, Chitosan, etc) with a large number of hydroxyl groups, all showed high-fidelity replicated features of the fiber-optic master (see Figures 4.3a and 3b, 3c). In contrast, for those without surfactant characteristic, such as Poly (methyl methacrylate)
(10wt% in toluene or in 1M acetic acid) and Poly (methyl acrylic acid, sodium) (10wt% in aqueous solution), the replicate conical tips were not shape enough, no matter aqueous
93 solution or organic solution was used (see Figures 4.3d, 3e, 3f). Some of these tips even have a rough tip surface because of the serious shrinkage during drying.
Relative humidity (RH) and temperature are two important factors during PVA solution drying. Both strongly affect the quality of polymer templates. Increasing the drying temperature leads to residual stresses while high relative humanity causes polymers to swell and elevates the structural instability. These phenomena can be explained by their influence on the mobility of polymer chains. Increasing drying temperature leads to more ordered (crystalline) conformations of polymer chains depending on interacting chemical functional groups, structure, and stereochemistry of the polymer. Drying too fast might lead to too much residual stresses inside the sacrificial template to stay flat on the PDMS surface. High relative humanity causes polymers to swell and elevates the mobility of individual polymer chains. The nanoscale structures turn to be instable and might not hold their shape because of their weak mechanical strength. Consequently, drying too fast (e.g., heating) or too slow (e.g. under high relative humidity) tends to cause distortion of the nanotips on the PVA sacrificial templates. In most cases, slow drying at arid ambient condition can ensure smooth PVA nanotips without noticeable residual stresses. Another concern for slow drying is about good replication quality. Slow drying can elevate the shrinkage and produce nanotips with smooth surface.
4.2.5 Fabrication of nanonozzle array
After drying, the sacrificial template was peeled off and attached onto a flat substrate (i.e., a glass slide). A polymer solution (e.g. 10-20 % PMMA in toluene) or a
94 liquid resin (e.g. PDMS) was then spun onto the sacrificial template. A thin polymer layer with open channels was formed by controlling the spin conditions to ensure that the film thickness was slightly less than the height of the nanotips on the sacrificial template.
The relationship of polymer film thickness and the spin speed is given in Figures 4.4. The desirable small end dimensions of nanonozzles can be finely tuned by using curves on that figure as the reference. For sacrificial templates with the height of 3-3.5 µm, the spin speed of 2,000-3,000 rpm was applied. After curing the resin or drying the solution, the nanonozzle array was released by dissolving the sacrificial template in hot water. This process avoids the de-molding challenge that commonly occurs in the conventional molding process. Although more steps are involved, they are much simpler and lower cost than silicon nanofabrication techniques.
Since the sacrificial template is made of a water-soluble polymer, its removal was fast, easy and safe, unlike reported sacrificial material removal approaches involving chemical etching or heating (described in the literature part). Based on the results of nine samples, the dissolving rate for the aperture part on the PVA sacrificial templates is around 0.6 µm/min, with PMMA layer coated on. The base parts dissolve much faster due to the large exposure area to water.
Figures 4.5a and 5b show the SEM images of two PMMA nanonozzle arrays from the sharp end. The surface of the polymer layer consists of an array of volcano-shaped nozzles, which is a result of the meniscus of polymer solution (or resin) on the PVA nanotips. Along the tip surface, surface tension is balanced with the gravity force. This meniscus effect was enhanced by the conical shape of those sacrificial nanotips.
95 Compared to liquid crawls along the vertical wall of a capillary, here surface tension only has to balance partial gravity (g*cosθ) along the outer surface orientation of the cones.
The final nozzle shape and channel diameter depend on the geometry of nanotips, viscosity of polymer solution or resin and the capillary effect. In Figure 4.5a, the height of each nanonozzle is 3 µm and the aperture size is less than 1 µm. The channel diameter is 80 nm on the sharp end and 1300 nm on the large end. Thus the convergence ratio of the 3 µm long channel is over 30. Nanonozzles in Figure 4.5b have a lower nozzle height and a larger channel diameter (200 nm) on the sharp end because of the use of a higher polymer concentration (i.e., higher solution viscosity) in the process. Using optic fibers with a smaller diameter and adjusting the etching solution composition, smaller nanochannels with different convergence ratios can be obtained. The small variation observed in Figures 4.5a and 5b results mainly from the difference of original optic fibers. This is verified by the distribution of individual optical fibers in images guide, shown in Figure 4.6. The size variation of optic fibers comes from the purposely disordered packing of optic fibers in the image guide to minimize cross talk in communication. Other factors such as replication accuracy and surface roughness of PVA template may also contribute to the channel size variation.
4.3 PLANIZATION OF APERTURE PART
When an aperture part of nanonozzle is inappropriate for a given application, it can be easily removed by planization techniques. Similar to the silicon based micromachining techniques, polymer planization can also be carried out in wet processes
96 (using liquid chemicals) or dry processes (using gases). They are called wet planization and dry planization, respectively.
4.3.1 Wet planization
In the wet planization process, liquid chemicals are used by dynamically spinning onto the surface of polymer substrate. The chemical must have desirable dissolvability of the polymer materials. Here, hydroxyethyl methacrylate (HEMA, monomer) was used since most of nanonozzle arrays were made of poly (methyl methacrylate acid, PMMA).
Compared to their molecular structure, methyl methacrylate (MMA, monomer) and
HEMA belong to the same group of monomers. Usually, many polymers can be dissolved in their corresponding monomers. Therefore, HEMA should be able to slowly corrode the PMMA surface. To accomplish wet planization, the process has to be quick and appropriate so that no further damage of the underneath polymer structure could happen. One easy way to fulfill this goal is to perform the wet planization by dynamic spin coating. HEMA is slowly dropped onto the spinning polymer substrate. Since the aperture part of nanonozzles dissolves much faster than the base polymer layer, the short contact time of dynamic coating of HEMA ensure the aperture part to be quickly dissolved while not long enough for further penetration to damage the underneath polymer structure. At a certain spin speed, an extra thin layer of HEMA is left on the polymer substrate. The retained HEMA in the PMMA substrate is then polymerized to form a thin layer of poly (HEMA) after curing HEMA under UV light. The rapid polymerization during dynamic coating can avoid the meniscus effect. Eventually a thin layer of poly (HEMA) is coated on the PMMA nanonozzle layer, which could
97 compensate the mechanical property loss due to the reduction of polymer thickness during planization. Morevoer, because the HEMA surface can be easily modified to graft other functional groups, this dynamics planization also provides a way to add different chemistry and charges on the outer surface of the polymer layer and the nanonozzle wall.
SEM images are shown in Figure 4.7 after spin HEMA on the PMMA nanonozzle surface with a spin speed of 3,000 rpm for 30 seconds. The surface of the final structure is much flat than the volcano structure without planization. The channel size correspondingly becomes larger due to the reduction of polymer thickness. In the shown
SEM image, the average channel size is around 700 nm. More irregular shape of channels was observed, indicating that there are certainly swelling-related dimensional changes during this wet planization process.
Beside the monomer/polymer wet planization, gentle solvents can also be used to planize the surface. Typically, planization is only performed to remove the aperture part.
No further compensation and surface modification is done.
4.3.2 Plasma planization
Plasma etching is another way to remove the aperture part. Since gases
(commonly oxygen) are used this time, plasma planization is actually a “dry” process.
Oxygen plasma is widely used to remove residual photoresist on the silicon substrate after dry etching, which is called “descum”. As one of the popular positive photoresists used in electronic beam lithography, the residual PMMA is left on the substrate after pattern transfer with reactive ion etching. Descum is one of the effective ways to remove the residual resist and clean the sample because PMMA could be etched pretty fast in
98 oxygen plasma. The removal rate depends on the energy input (e.g., RF power), gas components (e.g., oxygen flow rate) and operation temperature. The high RF power, high flow rate of oxygen will give a high removal rate. The dependence of the removal rate to temperature is pretty strong, i.e., it increases dramatically with temperature.
In this study, an inductively coupled plasma system (ICP, Oxford 130) was used to perform the plasma planization. ICP was used to ensure the uniformity of oxygen plasma. Oxygen was supplied at a flow rate of 40 sccm and RF power was set at 50 Watt.
The actual removal rate was 0.5 µm/min. To carry out this plasma planization, a thicker
PMMA layer was coated on purpose such that nanotips on the poly (vinyl alcohol) template were totally covered. But the top surface of the substrate is not even because of the meniscus effect. Figures 4.8a and 8b show the schematic and SEM image after a short time plasma planization on a thin polymer layer, respectively. The average channel size is
300 nm. Because of the anisotropic removal mechanism, the aperture and base have the similar removal rate. The initial surface morphology is roughly transferred during plasma removal. Therefore, same tiny aperture structures can still be found after plasma planization. When the coated PMMA layer is so thick that it completely eliminates the meniscus effect, that is, the initial surface is even, the resulting surface morphology is smooth after a long time plasma planization, as shown in Figure 4.8d. Therefore, plasma planization is more useful to open those channels by uniformly removing the polymer layer, rather than by selectively removing the aperture part only.
99 4.4 DERIVATION OF SACRIFICIAL TEMPLATE IMPRINTING
Sacrificial template imprinting does not need to include all three steps involved in the nanonozzle array fabrication and is not limited to produce nanonozzle arrays. With minor changes, the concept of sacrificial templates can be applied to generate many desired micro-/nanopatterns. For example, it can be used to easily obtain either negative or same replica from the mother mould when other fabrication techniques are not available.
In micro/nanofabrication, it is in principle true that any design can be fabricated into either positive or negative patterns. But in many cases, only one case (either positive or negative) is feasible in practice. Challenges come from many factors: the resolution of photoresists, replicate difficulties and so on. Additionally, many well-developed techniques are limited to fabricate patterns on silicon or glass. For example, EBL can easily generate nanowells or nanoholes but has difficulties to produce nanoposts or nanopillars. If nanowells or nanoholes are the desired patterns and plastic devices are required, sacrificial template imprinting can easily generate the negative or same replica with minor changes. Although PDMS can precisely replicate the negative pattern from silicon, it is not suitable in certain applications. For example, many hydrophobic molecules can adsorb and migrate into the PDMS matrix; many nonpolar solvents may swell PDMS devices. Also it is difficult to carry out polymerization within the PDMS channels because of its high oxygen permeable nature.
With the sacrificial template imprinting technique, patterns with the same or negative features on Si can be easily generated on the sacrificial template. They can also be transferred to a mold for further processing. Figure 4.9 shows the process to generate
100 PDMS microwells, starting with a PDMS pillar array mould. A PDMS pillar array mould is first hot embossed into a PMMA layer to produce microwells. Then a PVA aqueous solution is cast to generate the same pattern with PDMS mould. The PVA template with the micropillar array is finally cast with a PDMS precursor and cured to produce the
PDMS substrate with microwells. Another example is to obtain PDMS nanochannels, starting with PMMA nanochannels on a Si wafer written by EBL as shown in Figure
4.10. The photoresist SU-8 5 is employed to precisely replicate the same pattern by EBL.
A PMMA mould with nanochannels is obtained by using the PVA sacrificial template as the transition. These PMMA nanochannels can be used directly for nanofluidic studies.
The replication through the sacrificial template is not limited to produce PMMA or
PDMS patterns. In the demonstrations, these materials are used because they are widely employed in micro-/nanofluidics and the processes can be done rapidly.
4.5 DYNAMIC SILICA ASSEMBLY IN NANONOZZLES
4.5.1 Introduction
Limited by the height (~5 µm) of nanotips on the sacrificial template, the polymer nanonozzle array produced by STI is very thin (< 5 µm). Moreover, many commonly used polymers are structurally unstable due to their low glass transition temperatures and aqueous working environment. As a result, the applications of the nanonozzle membrane may be restricted by its low mechanical strength and dimensional stability. In addition,
101 the size (50 -100 nm) of nanotips on the template makes it very difficult to obtain nanonozzles smaller than 100 nm in diameter.
It is well known that self-assembly can create regular patterns at the nanoscale or even molecular scale with simple forms (thin films, particles, or fibers, Thurn-Albrecht,
T. et al, 2000). However, the self-assembly approach alone is not sufficient to scale up nanostructures with high aspect ratios over reasonably large areas without introducing significant defects. A hybrid process was developed in this study by growing silica on the internal surface of nanonozzles fabricated by the STI approach. This was realized via external force-induced dynamic surface reactions. Electrokinetic flow (EKF) was used to drive the molecules or their precursors into the nanonozzles and surface chemistry was used to control the growth of nanostructures on the channel wall. Such a combination provides an efficient way to further tune the channel size and reinforce the polymer nanostructure.
4.5.2 Experimental
4.5.2.1 Materials
Poly(allyamine hydrochloride,PAH, Mv = 720,000) and Tetramethylorthosilicate
(TMOS, monomer) were purchased from Sigma-Aldrich (St. Louis, MO). Poly(styrene sulfonate, PSS, Mw = 500,000) was purchased from Polysciences, Inc. Poly(styrene sulfonate) and poly(allylamine hydrochloride) solutions were prepared using DI water with 0.5 M NaCl adjusted pH value to 9.0 with sodium hydroxide (NaOH). The polymer concentrations mentioned later are calculated based on their repeat unit.
102 4.5.2.2 Surface modification
To promote surface silicification, a cationic polymer, polyallyamine hydrochloride (PAH), was used as the catalyst for the ionic silica condensation. It can be immobilized onto the outer surface as well as the internal area of the channel walls. Two different strategies were applied and no significant evidence was shown between the modification efficiency. One was carried out via amidation reaction under basic conditions as shown in Figure 4.11a. Briefly, clean PMMA or PET substrate was immersed in PAH solution (167 mg of PAH in 120 mL of water, PH=11.5) for 1 hour at room temperature. After rinsing with three aliquots of DI water, the as treated substrate was introduced into diluted hydrochloric acid solution (pH =2.2) for 30 min. The other approach was to hydrolyze the ester bond at first to introduce –COO- on the surface of substrate by cooking in 1 M aqueous NaOH for 15 min at 60oC (Figure 4.11b). PAH was then introduced onto the surface by layer-by-layer adsorption.
To achieve desirable concentration of PAH on the surface, layer-by-layer adsorption of polyelectrolyes was carried out after introducing the first PAH layer. The procedure was firstly introduced by G. Decher and now was widely used in the formation of polyelectrolyte multilayers (PEM). Multiple layers of PAH and PSS were deposited alternatively and the substrate stood in each polyelectrolyte solution for 30 min, with five-minute DI water rinses in between. The desired number of PAH layers depends on the stability of charge density and its catalyst efficiency. Typically, the results became way reproducible with at least three total layers of polyelectrolyte layers (including the initial one) and 3-7 total layers were applied in practice.
103 4.5.2.3 Dynamic assembly
Tetraethylorthosilicate (TEOS, Aldrich) was hydrolyzed under acidic conditions with the concentration of TEOS of 10-3 M (TEOS, 0.104 g; 37.5% HCl 4.88 g; DI H2O,
500g; the molar ratios are TMOS: HCl: H2O = 1:1:55.6) for 30 min. The hydrolyzed solution was then diluted by DI H2O with a ratio of 1:100 and continued hydrolyzing for another 30 min. The diluted solution was then immediately introduced into a two- compartment chamber separated by the nanonozzle array film. The reaction was carried out in the presence of a DC electric field (20-100V/cm), with the cathode immersed in the chamber facing the sharp end and the anode facing the large end. The reaction proceeded for 15 min, after which the nanonozzle array was taken out and rinsed thoroughly with DI water and dried.
4.5.3 Results and discussions
4.5.3.1 Dynamic assembly procedure
The schematic of dynamic assembly process is shown in Figure 4.12. The nanonozzle array was first treated by anchoring poly(allyamine hydrochloride) (PAH) onto the surface (see details in the experimental section). The role of PAH is two-fold: it enhances the electrokinetic flow (EKF) and thus moves the silica precursor into the conically shaped nanochannels and it also catalyzes silica condensation enhancing the surface reaction rate (Mizutani, T., et al, 1998; Patwardhan, S. V., et al, 2002). A dilute solution of silica source (10-5 M) was then introduced into the nanonozzle array layer by adding electric bias (20 V/cm-100 V/cm). The condensation reaction happened between
104 the deprotonated and neutral species in silica precursor to form silica, as shown in Figure
4.13. Polyamine (e.g., PAH) can accelerate this reaction by promoting the proton transfer between these two species. Different from the bulk silica formation process, this dynamic assembly process includes very dilute Tetraethylorthosilicate (TEOS) solution (e.g., 10-3-
10-5 M) to suppress homogeneous nucleation in the bulk and to promote heterogeneous nucleation at the surface (Aksay, I. A., et al, 1996; Trau, M. et al, 1997; Tarasevich, B. J., et al, 1996). After 30 min, the polymer layer is taken out and washed with copious of DI water and left to dry at the ambient condition.
4.5.3.2 Dynamic assembly in STI nanonozzle array
As a proof-of-concept, nanonozzle arrays of a nominal average diameter of ~200 nm were chosen as the template to conduct silica assembly. Figures 4.14a and 14c are the
SEM images of polymer nanonozzle arrays taken from the large end and the sharp end.
The diameters of the two ends of those conically shaped channels are 1300 nm and 200 nm, respectively. PAH was introduced on the internal surface of channel walls by surface treatment procedure addressed in the experimental section. After 30-minute running
TEOS (10-5 M) by electrokinetic flows induced by an 80 V/cm electric bias, the dynamic synthesis of silica reaction was stopped and samples were checked on both sides.
Figures 4.14b and 14d are the SEM images of the polymer nanonozzle arrays after the 30-minute dynamic assembly. From the large end, silica was clearly seen nicely deposited on both outer and internal surface of polymer nanonozzle array layer, as shown in Figure 4.14b. But the layer of silica on the outer surface is much thicker than that inside the channels. From the inserted picture in Figure 4.14b, it can tell that the diameter
105 of the large end is slightly reduced but not very pronounced. It can be clearly seen silica is grown deep inside the nanonozzle. Figure 4.14d showed the channel diameter on the sharp end after the dynamic assembly. The growth of silica on both the external surface and within the channel, in this particular case, leads to the average channel diameter being reduced from ~200 nm (Figure 4.14c) to ~50 nm (Figure 4.14d). The final structure is essentially a polymer/silica composite. Depending on the initial channel diameter, we believe that this technique has the potential to reduce the channel diameter to 10 nm or less by proper control of electrokinetic flows (EKF) and surface reaction. Moreover, it is also possible to add functionality to the polymer micro/nanofeatures in the future. Thus this approach would have the potential of a wide variety of multifunctional polymer based micro/nanodevices.
In the presence of the electric field, the interaction of the field with the electric double layer (EDL) near the charged channel surface induces the motion of the bulk liquid, or electroosmotic flow (EOF). The actual bulk flow of the liquid exists as the startup of electric bias. Such flow motions could become more obvious in 3D dynamic assembly when much higher electric bias (e.g., 100V/cm) was added in the two half-cells.
At the beginning, obvious liquid level difference can be built up with the liquid level on the anode side clearly higher than that on the cathode side. But half an hour later, the liquid level difference switches to the other direction with the liquid level on the cathode side clearly higher. The nanonozzle surface is initially positively charged because the top layer surface is PAH and therefore, electroosmotic flow drives liquid from the cathode to the anode. Simultaneously, the silica condensation reaction started on the surface of nanonozzles. Since the silica precursor itself is negatively charged, the silica assembly on
106 the channel will first gradually neutralize the surface charge and then make it become negative slowly. When the channel surface become negatively charged enough, the electroosmotic flow direction is switched to the opposite direction, that is the induced liquid motion became from the anode to the cathode.
4.5.3.3 Control of dynamic assembly procedure
An intrinsic difficulty associated with the assembly in submicro- or nanoscale channels is the constant supply of precursor solution to the interior of the channel. In the absence of flow during reaction, the reactant solution was drawn into the channel by capillary force and diffusion. The mass transfer limit leads to reactants depletion and cessation of silica growth within the channels. Furthermore, the growth of the silica on the external surface may block the channel entrance and prevent the diffusion of reacting precursor to the interior of the channel. Therefore, the first advantage of dynamic assembly is to introduce flows during the reaction. The flow of precursor provides always fresh precursor solution into the confined nanostructures to maintain the high reaction rate. Here electrokinetic flow, instead of pressure driven flow, is chosen for two purposes: one is the minimum request for the mechanical strength of polymer nanostructures and the other is the uniform concentration profile of precursor solution cross the nanochannels.
To accomplish electrokinetic flow, a direct current (DC) electric field was employed. The channel surface initially bears positive charges as result of the immobilization of PAH. In the presence of the electric field, the interaction of the field with the electric double layer (EDL) near the charged channel surface induces the motion
107 of the bulk liquid, or electroosmotic flow (EOF). The EOF drives the precursor solution through the channel and the reaction takes place dynamically. Significant EOF was observed for field strength as low as 12 V/cm and the field strength required to induce
EOF is much lower than that is needed by adsorption of charged surfactants on the wall
(Trau, M. et al, 1997). Meanwhile, silica precursor carries weak negative charge because of the present of the deprotonated specie, which led to a drift velocity of electrophoresis.
The actual motion of precursor was determined by both the bulk liquid motion (EO) and the drift velocity (EP) of silica reactant.
The silica formation involves a nucleation process that could occur by either homogeneous mechanism in the bulk precursor solution, or heterogeneous mechanism at the liquid-solid interface, depending on the local concentration of precursor (Bunker,
B.C., et al, 1994). With a high concentration of precursor, the bulk nucleation might not be suppressed efficiently that both types of nucleation happens and it is very likely that homogeneous nucleation occurs rapidly, leading to the formation of colloidal particles and particle aggregates in the bulk solution. When these colloidal particles deposit onto the surface through electrostatic interactions, it would lead to coarse particle aggregates on the surface and a non-uniform surface coverage. As deposition continues, the silica layers grow both vertically and laterally, leading to an increase in layer thickness as well as particle bridging over the pore. Similar observations have also been reported previously (Tarasevich, B.J., et al, 1996). In order to form a dense and smooth inorganic film, it is critical to promote heterogeneous nucleation while minimizing the homogeneous nucleation of particles in the solution. To achieve this, a low level of supersaturation is highly desirable during the process (Aksay, I. A., et al, 1996). Low
108 concentration of precursor solution could be one way to suppress homogeneous nucleation this but it can not be too diluted to obtain a desirable reaction rate at the interface. In order to promote the heterogeneous nucleation, and hence a dense silica layer growth on the interior of the channel surface, a uniform reactant concentration is essential throughout the reaction if local enriched region can not be achieved. Here comes the second advantage having electrokinetic flows in this dynamic assembly: the homogeneous reaction is suppressed by the short residential time of precursor and diluted concentration while heterogeneous reaction maintained because of the existence of the catalyst at the interface and similar concentration of precursor solution as that in the bulk.
The possible issue of blocking nanochannels can be avoided by the big nanoparticle aggregates. In practice, a diluted TEOS precursor solution (10-5 M) was used in our study to conduct the assembly.
The electric field strength is also important since it determine the flow rate and flow patterns inside the nanochannels. By adjusting the electric field strength, the reaction/nucleation rate of silica can be manipulated. For example, high field strength in nanochannels may produce significant local Joule heating, which can accelerate the reaction both in the bulk and at the interface of liquid and channel surface. Even at the low reactant concentration, the fast reaction rate quickly leads to high supersaturation, and consequently a rapid homogeneous nucleation. Moreover, the high field strength leads to a short residence time of the precursor solution within the channel, which is detrimental to the silica growth within the channels. Under the right combination of electric field strength and reaction time, the homogenous nucleation can be effectively suppressed during the entire reaction period, leading to a dense and uniform silica film.
109 Hence, there is delicate balance of the two reaction/nucleation rates to achieve substantial silica growth by the heterogeneous mechanism, while suppressing the homogeneous nucleation. Two key parameters control the silica growth in this dynamic assembly: the concentration of precursor and the electric field strength. They determine the nucleation/reaction rate, the type of nucleation reaction and their partition contribution to this dynamic assembly process. More precisely tuning of the dynamic assembly would produce more desirable nanostructures (e.g., smaller channel size, desirable channel geometry and high mechanical strength). To obtain the optimal operation conditions or correct guideline of this dynamic process, it is necessary to well understand the actual flow inside the nanochannels and also the mechanism of this dynamic assembly. Attempts were done within similar 2D converging microchannels and will be described in next chapter.
4.5.3.4 Mechanical property testing
The hardness of both the pristine and hybrid channel array was measured by atomic force microscopy (AFM) operated in the nanoindentation mode. A three-sided, pyramid-shaped diamond tip was used to indent the sample with a known force or load F.
The indented region was immediately imaged under the tapping mode to obtain the projected residual area A. The hardness is calculated by dividing the force by the area.
The results are shown in Figure 4.15. The pristine PETE has an average hardness of 239
MPa, while the hardness of the hybrid channel array is 712 MPa. After silica growth, the hardness increases by about 300%. The dynamic assembly of silica in the channel array provides the anticipated reinforcement to the polymer structures.
110 4.6 SUMMARY
In brief, sacrificial templates imprinting (STI) is introduced and applied to the fabrication of polymer nanonozzle array. A polymer sacrificial template, instead of its mother master, is used in the fabrication, which can be massively produced and easily removed after molding. The formed nanonozzles have volcano shape with half a micrometer high aperture part. Using wet or dry planizations, the aperture part can be removed if not desirable to the applications. Silica dynamic assembly is then introduced to further reduce the size of nanonozzles as well as reinforce their mechanical strength.
Electrokinetic flow is applied to assist this silica surface assembly by continuously providing precursors into nanonozzles and avoid the possible blocking of channels by aggregations. As the demonstration, the channel size was successfully reduced to 50 nm for a 200 nm nanonozzle and the nanostructure was enhanced.
111
Optical fiber bundle Nanotip array PDMS Mold Wet Replica etching molding
Sacrificial Casting Template Template 112 Nanonozzle Array removal Spin in water coating
Figure 4.1 Schematic of the Sacrificial Template Imprinting (STI) process for fabricating nanonozzle arrays with uniform, conically shaped nanochannels.
112
(a) (b)
15.0 kV 50000x 500nm 15.0 kV 25000x 1µm
(c) (d)7
6.5
6
5.5 /2) θ 5
Cot ( 4.5 y = -0.0432x + 6.7508 4
3.5 15.0 kV 20000x 1µm 3 0 102030405060
Temperature, OC
Figure 4.2 SEM images of nanotip formation study in BOE (NH4F/HF=7:1) at room temperature: (a) after 10 min; (b) after 20 min; (c) after 30 min and (d) the temperature effect on the nanotip geometry, resulting from the different etching rates of core and cladding materials.
113
C D
Continued
Figure 4.3 SEM images of: (A) a nanotip array from an optic fiber bundle; (B) a nanotip array cast by poly(vinyl alcohol) aqueous solution, 10 wt%; (C) a nanotip array cast by chitosan 1M acetic acid solution, 1.0 wt%; (D) a nanotip array cast by poly(methylacrylic acid, Sodium) aqueous solution, 10 wt%; (E) a nanotip array cast by poly(methyl methacrylate) toluene solution, 10 wt%; and (F) a nanotip array cast by poly(methyl methacrylate) acetic acid solution, 10 wt%.
114 Figure 4.3 continued
E F
115
10 9 20 wt% 15 wt% 10 wt% 8 7 m)
µ 6 5
116 4
3
Film Thickness ( Thickness Film 2
1 0
0 1000 2000 3000 4000 Spin Speed (rpm)
Figure 4.4 The spin speed versus thickness curves for PMMA/toluene solutions with different solid contents.
116
Figure 4.5 SEM images of poly(methyl methacrylate, PMMA) nanonozzle arrays formed with (A) 15 wt% and (B) 20 wt% PMMA solution in toluene.
117
0.5
Nanonozzles 0.4 optic fibers
0.3
F 0.2
0.1
0.0 0.00.10.20.30.40.50.60.70.80.91.01.11.21.31.41.5
Size, micron
Figure 4.6 Comparison of size variation of nanonozzles and original optic fibers. The solid circle represents for nanonozzles and hollow circle for optic fibers.
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(a)
HEMA PHEMA
PMMA
(b) (c)
Figure 4.7 The Schematic of (a) and SEM images of dynamic planization using hydroxyethyl methacrylate, HEMA: the landscape-scale view (b) and an individual channel (c).
119
(a) (c)
O2 plasma O2 plasma
(b) (d)
Figure 4.8 The Schematic and SEM images after dry planization by oxygen plasma etching: a short period removal with a thin PMMA layer (a-b) and a long period removal with a thick PMMA layer (c-d).
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Solution Casting (a) PDMS male mold PMMA female mold
Hot Embossing Solution Casting
Resin Casting PDMS female mold PVA sacrificial mold
(b) (c)
(d) (e)
Figure 4.9 The derivatives of sacrificial template imprinting I—obtain negative pattern with the same material: schematic of the process (a) and SEM images of PDMS male mold (b), PMMA female replica (c), PVA sacrificial mold (d) and PDMS female mold (e).
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(a) PMMA nanochannels on Si substrate
Solution Casting
PDMS mold
Resin Casting
SU_8 mold
Solution Casting
PVA sacrificial mold
Solution Casting
PMMA nanochannels
Continued
Figure 4.10 The derivatives of sacrificial template imprinting II—obtain same pattern starting with EBL patterns: schematic of the process (a) and SEM images of PMMA photoresist nanochannels on Si substrate (b), PDMS male mold (c), SU-8 nanochannels (d), PVA sacrificial template and (e) PMMA nanochannels (f).
122 Figure 4.10 continued
(b) (c)
(d) (e)
(f)
123
(a)
O O NH3+Cl-
O PAH, pH=11.5, 60min HN + - NH3 Cl O pH=2.2 , 30min O
O HN NH3+Cl-
+ - O O NH3 Cl
O HN PAH, pH=11.5, 60min OH O + - O NH3 Cl
O pH=2.2 , 30min HN OH + - NH3 Cl
(b) O O
O 1 M NaOH O_
o O O 60 C, 15 min O_ O
O O 1 M NaOH - O O
O o 60 C, 15 min O
O O-
Figure 4.11 Strategies to introduce polyallyamine hydrochloride (PAH) on PMMA and PET surface: through amidation reaction under basic conditions (a) and hydrolyzing the ester bond under strong base solution (b).
124
125
Figure 4.12 Schematic of dynamic assembly of silica assisted by electrokinetics.
125
(a)
Si - Si OH OR + OH - - Si OH + OH Si O
(b)
Si O- + Si HO
- Si O Si + OH
Figure 4.13 Reaction mechanism of silica formation reaction: (a) the hydrolysis of
Tetraethylorthosilicate (TEOS) precursor to form Si(OH)4 at pH 2-7 and the condensation of the deprotonated and neutral species to form silica.
126
(a)
(b)
Continued
Figure 4.14 Scanning electron microscopy (SEM) images of polymer nanonozzle array before (a) & (c) and after dynamic assembly (b) & (d): (a) & (b) on the large end, (c) & (d) on the small end.
127 Figure 4.14 continued
(c)
(d)
128
900 800
700
600
500
400
300
200
Hardness (MPa) 100 0 PET PET/silica
Figure 4.15 Comparison of hardness of PET and PET/silica polymer layers.
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CHAPTER 5
NANOPARTICLES AND SINGLE DNA DYNAMICS IN CONVERGING/DIVERGING FLOWS
5.1 INTRODUCTION
In the previous chapter, the sacrificial template imprinting (STI) process and its application to produce conically shaped nanonozzles are described. In conjunction with silica synthesis on the channel surface by dynamic assembly, the channel size can be further reduced and the polymer structure can be reinforced.
To investigate the mechanism of this dynamic assembly process, similar 2D studies were carried out in converging microchannels in this chapter, but with the small end scaled up to 20 µm in diameter (a scale ratio of 100). Both converging and diverging flows were investigated and the dynamic complexation was carried out to simulate the
3D dynamic assembly process in nanonozzles.
The nanonozzles can provide two important flow patterns: converging flow and diverging flow. In the second part of this chapter, particle transport through nanonozzle arrays was investigated in both flow directions with electrolyte suspensions. The testing nanoparticles include rigid Fluoresbrite (YG) nanosphere suspensions (40-200nm) and flexible bacteriophage λ-DNA (48.5 kbp). In comparison, 2D electrokinetic-induced
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flows were also carried out in converging channels with the small end of 5 µm in diameter, to investigate the transport phenomena observed in the 3D nanonozzle array.
5.2 2D MICROFLUIDIC STUDY
To investigate the mechanism of the dynamic assembly process described in the previous chapter, 2D electrokinetics-induced flows and dynamic complexation were studied using the same procedure and same geometry as in the 3D dynamic assembly, except that the channel size was scaled up to a level that the small end is 20 µm in width
(a scale ratio of 100). Correspondingly, larger nanospheres are used to maintain the ratio of dparticle/dnozzle closer to that used in 3D studies. Other materials and experimental conditions were kept almost the same as in 3D experiments unless otherwise specified.
5.2.1 Experimental
5.2.1.1 Materials
Suspensions of Fluoresbrite Yellow Green (YG) nanospheres were obtained from
Polysciences, Inc. The nanosphere suspensions with the solid content of 0.00265% were made by diluting stock suspensions (2.65% solid) by a factor of 1000 in demineralized distilled water (pH = 6.5).
5.2.1.2 Fabrication 2D microfluidic channel
2D glass microfluidic channels were fabricated by photolithography followed by wet etching. In brief, the glass slides were first spin-coated with hexamethyldisilazane
(HMDS) solution (4000 rpm for 30 seconds) and baked at 110 oC for 2 min to improve
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the adhesion. AZ 5214 positive photoresist (PR) was then coated (3000 rpm for 45 seconds) and baked at 105oC for 1 min. After exposed at 70 mJ/cm2 dose with i-line using the MJB-3 aligner, samples were developed in the MF 319 solution (Shipley, Co.) for 45 seconds. Hard baking may be necessary if stripe-off of photoresist occurs early in the following wet etching step. Finally, wet etching was carried out to transfer channels onto a glass substrate by immersing samples in the same BOE (7:1) etchant used in
Chapter 4.
2D PDMS microfluidic channels with the same geometry of nanonozzles were fabricated by first patterning the microfluidic structure on the silicon wafer using photolithography. Negative photoresist, SU-8 50 (Microchem, Inc) was used to fabricate channels with the smallest dimension of 20 µm following the protocol provided by the vendor. PDMS channels were then made by casting the mixture of 10:1 ratio (by weight) of PDMS precursor and curing agent (Sylgard 184, Dow Corning) on the patterned SU-8 structure using soft lithography. The mixture was degassed in vacuum for at least half an hour and then cured overnight at 65oC in an oven. Image of 2D PDMS microscale converging channels is shown in Figure 5.1.
5.2.1.3 Charecterization
The electrophoretic (EP) mobility of nanoparticles was measured using a
ZetaPALS zeta potential analyzer (Brookhaven Instruments Company, Inc). The electroosmotic (EO) mobility of different solid surfaces was characterized using a BI-
EKA streaming potential analyzer (Anton Paar, Inc, Austria).
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5.2.1.4 Flow Imaging
Microfluidic imaging techniques mentioned in Chapter 3 are applied in this section. PDMS or glass chips with 2D converging or straight channels was mounted onto the stage of an inverted epifluorescence microscope (TE 2000S, Nikon, Japan), using the same experimental setup as in Chapter 3. Negatively charged polystyrene microspheres
(700nm, and 3.0 µm, C=0.00265 wt%) were used as the tracer for flow visualization with
10 and 40x objective lens. The streamlines of the flow pattern were generated by compounding the video graphs.
5.2.2 Results
5.2.2.1 Flow profile inside the converging channels
To understand the dynamic assembly process, the first step is to map the velocity profile in converging/diverging flows. Both PDMS/glass hybrid and glass converging channels were used with fluorescently labeled PS microspheres of 3 µm in diameter as the tracer. In PDMS/glass hybrid converging channels, a few particles were observed to flow backward while the majority of particles migrated forward to the anode. Since the glass surface at the bottom has a higher zeta potential than other surfaces made of PDMS, the flow field in these 2D microchannels is not symmetric in the vertical direction.
Consequently, bi-directional flows were observed everywhere in the imaging window inside the converging microchannels because the fluorescent microscopy illuminates more than a single plane in the vertical direction. To check how the non-uniformity affects the backflow, complete glass channels were used for comparison. With the glass
133
channels, two clear vortices with a compressed ellipse shape were symmetrically distributed in regions close to the channel walls. The vortices were highly squeezed in the transverse direction to the flow, as shown in Figure 5.2a. The upper and the lower vortices rotated clockwise and counterclockwise, respectively. The zeta potential of glass is reported to be around four times higher than that of PDMS (Sylgard 184, Dow
Corning). Since the only change comes from the replacement of PDMS with glass, the large vortices formation is believed to be the result of the enhancement of electroosmotic
(EO) flow in that region.
Since vortices were observed in the converging channels for flows induced by electrokinetics, the electrokinetic effects were investigated by carrying out pressure- driven flows in the same converging channels with the same PS suspensions. With a similar particle migration speed or particle Reynolds number (Re~1.0), particles in this pressure-driven converging flow always followed the streamline inside the channels as
duρ shown in Figure 5.2b. Here, the Reynolds number for particles is defined as Re= , η where d is the diameter of the small end of the converging channel, ρ and η is the density and viscosity of the suspension, respectively, and u is the velocity of tracer particles at the small end of the converging channel. We choose particle Reynolds number for comparison because the motion of particles is the main focus of this study and it can be easily measured. It should be noticed that the liquid flow rate is substantially different from the motion of particles in the electrokinetics-induced flows while particles have a similar velocity as the fluid in pressure-driven flows in our experimental conditions.
134
To ensure that the suspensions used here are “dilute” enough and can be considered as the Newtonian fluid, the viscosities of various PS suspensions were measured using MCR300 rheometer (Anton Paar, Austria) with a 50 mm parallel plate fixture. With the solid content in the range of 1/100-1/1000 of their stock concentration, the viscosity of suspensions remained unchanged with the shear rate larger than 10 (1/s), very close to the viscosity of water (0.001 Pa⋅s) as shown in Figure 5.3. The scattering of data in the low shear rate range resulted from reaching the measurement limitation of the torque. Considering the shear rate in our system is much higher than the upper limit
(>100 1/s), the PS microsphere suspensions used in this study belong to the “dilute” suspensions, and could be treated as the Newtonian fluid.
The pressure-driven flows were carried out across a broad range of Reynolds number (from 1.0 to 500). No clear vortices were observed, which is quite different from the observation in non-Newtonian fluids with the similar geometry (e.g., 2 wt% PEG solution, see Kang, K, 2004). The observations in pressure-driven converging flows suggest that the vortices inside the channels observed in the electrokinetics-induced converging flows are related to the electrokinetic effects.
To further investigate the mechanism of vortex formation, electrokinetics-induced flow in straight channel with the channel width similar to the small end of the converging channel, i.e., 20 µm, was also carried out. When the similar field strength (i.e., 60V/cm), nanoparticles were observed following the streamline inside without any vortices as shown in Figure 5.2c. This result implies that the channel geometry plays an important role on the vortex formation in electrokinetics-induced flows.
135
5.2.2.2 Flow profile at the outlet
In addition to inside the converging channels, vortex flows were also observed outside the small end in PDMS/glass hybrid channels. When smaller rigid particles (e.g.,
40 nm PS nanospheres) or λ-DNA solutions were used, two large and symmetric vortices were observed at the exit of the converging channels as shown in Figures 5.4a and 4b.
The upper and the lower vortices rotated clockwise and counterclockwise respectively, with had the same rotation direction as the two vortices inside the channel. The majority of particles followed the streamlines, passing through the small end while particles in the adjacent regions were trapped in the two vortices.
The flow profile at the outlet of a straight channel is quite different from that of a converging channel. The straight channel size applied here was 20 µm in width, the same dimension as the small end of the converging channel (i.e., 20 µm).When a similar electric bias was applied (i.e., 60V/cm), nanoparticles followed the streamlines in the sudden expansion region without any vortices as shown in Figure 5.4c.
In the pressure-driven flow in the converging channels, Figure 5.4d shows that the fluid followed the streamline at low Reynolds number, in comparable to the value in the electrokinetics-driven flow and no lip vortices were observed at the exit. Until Reynolds number was higher than 10, as shown in Figure 5.4e, the fluid escaped from the small end carries a high momentum that a low pressure regions was formed around the lip with high pressure regions in the expanded channel corner. At higher flow rates (e.g., Re=100-250), vortices were expanded to the corners of the expanded channels (Figure 5.4f). The higher the momentum it gained inside the converging channel, the larger the affected regions were. The lip vortices had their centers in the middle of the vortices and more particles
136
were observed near the edge of the vortices than at the center. On the contrary, in the electrokinetics-induced converging flows, the two lip vortices were not center symmetric and their centers were offset to one side (the solid edge side of the exit). More particles were trapped at the eccentric center of vortices and the vortices were slightly extended inside the small end. With the growth of vortices and more trapped particles, particles started to aggregate and deposit on the solid surface, both inside and outside of the small end of the channel.
5.2.2.3 Flow profile in the diverging and straight channels
When the electric bias is reversed, particles migrate in the diverging direction.
With the same field strength (i.e., 60 V/cm) as in the converging direction, particles followed the streamline in the diverging direction without any vortex (Figure 5.5a). When the field strength was increased to 120 V/cm, particles started circulating at the two edges of the entrance (the small end) and a great number of particles were trapped in these vortices as shown in Figure 5.5b. An interesting observation here is that the two vortices had the same rotation direction as those inside and outside the converging channels. For comparison, pressure-driven flows were also carried out in the diverging direction. No recirculation motion of particles was observed in the same geometry, even at a much higher Reynolds number as shown in Figures 5.5c and 5d. This indicates that the motion of particles at the small end is not simply a result of high momentum of nanoparticles.
The electrookinetic effects have played an important role.
At the entrance of a straight channel, the motion of particles in the sudden contraction flow was quite similar to that in the diverging flow case. When a low field
137
strength was applied (i.e., 60 V/cm), particles followed the streamlines. After increasing the field strength, particles started recirculating and aggregating at the entry as shown in
Figures 5.5e and 5f.
5.2.3 Discussions and force analysis
5.2.3.1 Electrokinetic forces
Polystyrene nanoparticles used in the experiments carry negative charges. In electrokinetics-induced flows, such particles experience electrophoresis (EP) at the presence of electric bias and its quantity is expressed as electrophoretic mobility. The EP mobility of various nanoparticles used in the study is listed in Table 5.1.
Since converging channels are made of either polymers (e.g., acrylic) or silica
(during and after the dynamic assembly), their surfaces also carry weak negative charges
- - derived from weak acids, e.g., -COO or intermediate acid, e.g., SiO3 . In solutions, not all of the acidic groups are dissociated and the charge density is strongly dependent on the pH value of the solution. Besides, the dissociation reaction occurs randomly along the polymer chains, which could readily change due to the variation of local conditions. But when the surfaces are modified by grafting of strong polyelectrolytes, e.g., poly(allyamine hydrochloride) and poly(styrene sulfonate), which are completely dissociated, the charges are permanent and their charge density is constant on the entire surface. Zeta potential of various polymer substrates is plotted versus pH value in a broad range as shown in Figure 5.6.
138
The channel surface charge induced electroosmosis also plays an important role in the electrokinetics–induced flow. In an ideal electrokinetic flow, the similitude between the velocity field and the electric field is satisfied. The velocity components of both electrophoresis and electroosmosis are linearly proportional to the local electric field strength. The superposition of these two components can be expressed as
u P = (µ EP − µ EO )E (5-1)
where µ EO is the electroosmotic (EO) mobility, µ EP is the electrophoretic (EP) mobility, and E is the local electric field vector. EOF could be in the same direction as EP (i.e., positive surface charge with negatively charged particles, or vice versa) or in the opposite direction (i.e., same charge on nanoparticles and channel surface). Specifically in our experiments, the EO component of the velocity is in the opposition direction to that of the
EP component when the surface of nanonozzles is negatively charged, and in the same direction of the EP component when the surface of nanonozzles is positively charged
(e.g., in the dynamic assembly).
The ratio of EP mobility and EO mobility can affect the actual flow patterns (refer to Chapter 3, Electrokinetic Interactions Section). In most cases, the particles used in the
2D converging microfluidic study were 3 µm microspheres with an EP mobility of -0.78
×10-8 m2/(sV). The EO mobility of the PMMA surface is -1.01 ×10-8 m2/(sV) at pH=6.5, calculated from equation (2-3) with the zeta potential data from Figure 5.6. Since the ratio of EP and EO is 0.77, which is close to the range discussed in Section 3.3.2.2, both
EP and EO effects have to be considered in cases discussed in this chapter.
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The electric field strength ( E) is equal to the potential gradient (E = −∇φ ), while the electric potential (φ ) at steady state can be obtained by solving the Laplace equation in the converging channel.
∇ 2φ = 0 (5-2)
Since the microchannels were made of polymer or glass, which are insulators for electric currents, the Neumann boundary condition is applied at the channel wall. The Laplace equation here implies the continuity of the electric field. In nanonozzles or 2D converging channels, the electric field strength (E) varies along the conical channels due to the change of the cross section area. Figure 5.7 shows the calculated electric field strength in a 2D converging geometry. When the electric bias is from the large end to the small end (i.e., the converging direction), the electric field strength (E) jumps from the bulk value (E0) to a higher value (EL). Along the contracting channels, the strength of the electric field increases parabolically to a higher value (EH) near the outlet and then descends drastically until reaching the bulk value (E0) outside the small outlet. For a nanonozzle with a converging ratio of 15, the magnitude of the electric field at the small end (EH) is around 225 times higher than that at the large end (EL). A great gradient of the electric field strength exists in the converging channel.
In an ideal electrokinetic flow, charged particles flow along the electric field lines.
From our experimental results, the motion of nanoparticles far away from the channel wall followed the streamline (Figure 5.2). Figure 5.8 provides the experimental result of lateral velocity profile at the centerline along a 2D converging channel. The shape of the velocity profile matches the calculated result from equations (5-1) and (5-2). It has a similar profile as the electric field strength along the converging channels (Figure 5.8).
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However, this similitude between velocity and electric field does not be held everywhere in the converging channels. In the presence of electroosmotic flow (EOF), induced motion of the bulk liquid actually exists as soon as the startup of electric bias.
The EOF flow could be in the same direction as the negatively charged nanoparticles (if positive surface charge) or in the opposite direction (if negative surface charge). For negatively charged converging channels and nanoparticles, the actual EOF is in the opposite direction of EP, that is, the flow is actually in the diverging direction when the electric bias is added in the converging direction. No matter the migration direction of particles and the sign of channel surface charge, there is always a net flow caused by
EOF with a plug-like velocity profile. This net flow would induce an extra pressure difference opposite to the EOF direction in the channel. Velocity of fluid (V) can not be calculated from the equation (2-3). Instead, it could be calculated by solving following equations combining with equation (5-2).
1 V • ∇V = []− ∇p +η∇ 2 V − εk 2ψ∇φ (5-3) ρ
∇ • V = 0 (5-4)
∇ 2ψ = k 2ψ (5-5) where φ , ψ , 1/ k , ε , ρ and η are the electrostatic potential, surface potential, Debye layer length, permittivity, the density and viscosity of the solution, respectively.
When two ends of channels are closed and not connected to balance this pressure difference, it would lead to a liquid level difference, resulting in an imposed pressure difference. These two types of pressure difference (both induced and imposed) provide a
“backward” velocity component with a parabolic velocity profile. Combining with the
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forward “plug-like” velocity profile caused by electroosmotic flow, there could be different velocity profiles, such as a bi-directional laminar flow (Figure 5.9a) in a straight channel with a uniform cross-section area and a fixed surface charge (Lettieri, G.-L., et al, 2002, and Daiguji, H., et al, 2005). Usually the pressure-driven flow is predominant near the center of the channel while the flow direction is reversed to the EOF direction in regions close to the wall. The actual net flow direction (if not zero) depends on the relative magnitude of two types of flows. The converging/diverging geometry would produce a velocity component in the transverse direction, leading to the appearance of recirculation flow as shown in Figure 5.9b. The whole glass channel has a higher surface zeta potential than the PDMS/glass hybrid channel, which enhances this recirculation flow and exhibits a pair of pronounced vortices (Figure 5.2a).
5.2.3.2 Viscous drag and hydrodynamic interactions
Since the charged particles could gain a high velocity inside the conically shaped channels, viscous drag and hydrodynamic interactions also need to be considered, especially near the small end. As shown in both pressure-driven and electrokinetics- induced flows, nanoparticles can gain high momentum inside the converging channels. In the pressure-driven flow, both particles and fluid migrate in the same direction and have almost the same velocity. But in the electrokinetics-induced flow, the fluid is slowly moving in the opposition direction of particles because of EOF. Charged particles are accelerated when they pass through the converging channels because of the existing high electric field gradient. These high speed particles meet with the nearly still surrounding liquid outside the small end. A sudden deceleration of a large amount of nanoparticles
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could result in very large drag forces on the fluid and the excited liquids could exert additional hydrodynamic forces onto adjacent nanoparticles. By generating and reacting to the local velocity of the fluid, colloidal particles experience hydrodynamic interactions. When such interactions are near the edge of the exit, extra hydrodynamic interactions with the channel wall would occur.
Hydrodynamic interactions are very complicated and are still not fully understood. Only several limiting cases have been investigated, e.g., well-separated particles moving slowly through a viscous fluid. This well-known case is described by
Stokeslet analysis, which gives the simplest relation of a drag coefficient to express the viscous drag effect:
ξ = 6πηd (5-6) where η is the viscosity of the fluid and d is the diameter of the particle. Stoke’s equation is widely used in the calculation of drag forces. But this equation is only accurate with “diluted” particles and cases when Reynolds number is less than 1.0 because of the ignorance of high order terms in its approximation scheme. Few cases for many-body systems such as colloidal suspensions were studied and no simple equation of hydrodynamic interactions can be found in such complicated cases except some empirical equations. Stoke’s equation is often overused in many cases for rough estimation of the hydrodynamic interactions. For example, EP and EOF mobilities are widely used in electrokinetics-induced flow simulation while their expressions are derived by assuming that the drag forces follow the Stoke’s equation. In reality, pronounced hydrodynamic interactions are expected in cases where a large difference of velocity exists between
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particles and the surrounding liquid or solid edge and when a large number of particles are moving in a constrained geometry.
The hydrodynamic interactions at the small end of the converging channel are believed to have important contributions to the formation of backflow or vortices (Figure
5.4a and 4b). The different flow behavior observed in the converging channel and diverging/straight channel can be explained by the different hydrodynamic interactions at the small end. The momentum of particles is much lower at the entry of the diverging and straight channel than in the converging channel because of the lack of acceleration.
Therefore, the hydrodynamic interactions are not strong enough to generate vortex in those two cases unless the bulk field strength (E0) becomes high enough such that particles could gain a strong initial momentum. In the converging channel, fluid driven by electroosmosis actually flows in the diverging direction. Because the diverging direction offers less flow resistance (i.e., less back pressure flow) than the converging direction and straight channels, the electroosmosis-induced diverging flow has a higher flow rate than the converging flow and flow in the straight channel (Singhal, V., et al,
2004). Consequently, the fluid velocity in the converging channel is also higher than that in the other two types of channels. With the velocities for both particles and fluid high, the hydrodynamic interactions between particles and liquid in the converging channel are much more pronounced than in the diverging and straight channels the. This explains the different experimental results observed in the converging channel and in the other two types of channels. Near the small end, both the diverging channel and straight channel experience similar hydrodynamic interactions. Therefore, it is not surprising that a similar flow behavior is found in these two cases at the small end as shown in Figure 5.5.
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5.2.3.3 Dielectrophoresis(DEP)
Because of the presence of a highly non-uniform electric field inside the converging channel, dielectrophoresis may also be important, especially near the small end. Dielectrophoresis is the motion of particles produced by the induced dipole moments between particles and their surrounding medium, resulting from the non-uniformity of the electric field. Most dielectrophoresis studies relied on purely alternating current (AC), but most recently, some direct-current (DC) dielectrophoresis studies were reported in the highly non-uniform electric field using microfabricated patterns with electrodes very closer to each other (Chou, C. F., et al, 2002). With DC dielectrophoresis, the effective dipole moment for a spherical particle is given by (Pohl, H. A., 1978),
⎛ ε − ε ⎞ p = 2πε ⎜ p m ⎟r 3E (5-7) DEP m ⎜ ⎟ ⎝ ε p + 2ε m ⎠
Here, ε is the permittivity and p and m represent particle and medium, respectively, r is the radius of particle and E is the electric field strength. The dielectrophoresis force acting on a spherical particle is given by
FDEP = (p DEP • ∇)E (5-8)
Substituting equation (5-6) into equation (5-7) yields,
⎛ ε − ε ⎞ F = 2πε ⎜ p m ⎟r 3∇(E • E) (5-9) DEP m ⎜ ⎟ ⎝ ε p + 2ε m ⎠
From equation (5-9), the motion of particles could be away or towards the regions of the high electric field intensity, depending on the polarity degree of particles and medium. If the induced dipole moment of the particles is greater than that of the fluid, the particles exhibit “positive dielectrophoresis” and will move towards the regions of the
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high electric field intensity. On the contrary, if the surrounding liquid is more polarizable, the particles exhibit “negative dielectrophoresis” and will move away from the high-field regions. For the system employed in our study, the dielectric constants of the medium
(water) and nanoparticles (polystyrene) are 78.5 and 2.55, respectively. The corresponding “induced dipole moment” is negative, according to equation (5-7).
Therefore, “negative dielectrophoresis” must be present if significant. This means that nanoparticles would always move away from the high field strength region.
When the particle suspension is sufficiently dilute, that is, the interactions between particles can be ignored, the motion caused by dielectrophoresis can be constitutively related to the gradient of the dot product of the electric field strength via a
factor called dielectrophoretic mobility, uDEP (Cumming, E. B., et al, 2000),
u DEP = −µ DEP∇(E • E) (5-10)
The dielectrophoretic mobility is a function of the properties of both particles and surrounding medium, and is expressed as
⎛ ε − ε ⎞ µ = r 2ε ⎜ m p ⎟ / 6η (5-11) DEP m ⎜ ⎟ ⎝ ε p + 2ε m ⎠
Unlike the electrokinetic flow, the dielectrophoretic motion is along the electric field gradient. Two typical motions exist when DEP is dominant. One is called
“dielectrophoretic streaming”, which happens when DEP overcomes diffusion but is slightly weaker than electrokinesis. In this situation, dielectrophoresis can concentrate and rarefy particles. The other is called “trapping”, when DEP becomes stronger, overcoming even electrokinesis. Particles in this regime are trapped to form large
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aggregates. The condition for dielectrophoretic trapping can be expressed as (Lapizco-
Encinas, B. H., et al, 2004),
µ ∇(E • E) DEP • E > 1 (5-12) µ EP − µ EO (E • E)
Substituting equations (5-11) and (2-4) into the expression on the left side of equation (5-
12), the ratio of dielectrophoresis to electrokinesis can be approximately expressed as
ε r 2 ε − ε ∇(E • E) r 2 ∇(E • E) m m p • E ≅ (5-13) 6ε m (ζ p − ζ S ) ε p + 2ε m (E • E) 12(ζ p − ζ S ) E
A rough estimation of dielectrophoresis effect can be done using equation (5-13).
The value of E2 in the converging channel is firstly obtained from the calculated electric field strength (E) using equation (5-2), as shown in Figures 5.10a and 10b. The ratio of the dielectrophoretic mobility to the electrokinetic mobility for different polymer substrates is then obtained by equation (5-13). The ratio along the centerline of the converging channel is plotted in Figure 5.10c. From Figure 5.10c, it can be found that the
µ ∇(E • E) ratio ( DEP • E ) is less than 1.0 in most regions of the converging µ EP − µ EO (E • E) channel. This means that electrokinetic effects dominate where the local field strength is relatively low. The location of vortices inside the converging channel falls in these regimes. But dielectrophoresis becomes significant towards the small end. With the increase of the field strength and the gradient of the dot product of the field strength itself, the value of this ratio at the small end can become larger than the unit. For 40 nm
PS nanoparticles, the ratio is over unit when the field strength reaches 20V/cm, which is smaller than the field strength used in experiments (e.g., 60V/cm).
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Considering the fact that both dielectrophoresis and hydrodynamic interactions result from the great electric field and velocity gradient at the small end, these two factors are coupled together here. It is difficult, if not impossible, to isolate them and measure how much each of them contribute to the motion of particles near the small end. The lack of a simple mathematic expression of hydrodynamic interactions makes the quantitative estimate difficult. Besides, when hydrodynamic interactions become dominant, the simple expression of dielectrophoretic mobility by equation (5-11) is not held any more.
The estimation of actual DEP effect based on equation (5-12) can be done only if the mobility data are obtained from other ways (e.g., experimental measurements). As pointed out before, “negative dielectrophoresis” should be performed in our cases. That is, nanoparticles should move away from the high field regions (i.e., the small end). This does not match our observation of the recirculation motion of nanoparticles there. To completely understand the mechanism of motions of nanoparticles in this converging channel, more work, especially advanced models and simulation tools, should be done.
At this point, we can only conclude that both effects are important and responsible for the phenomena observed outside the small end.
In conclusion, four major effects exist: dielectrophoresis, electrokinesis (including electrophoresis and electroosmosis), and the hydrodynamic interactions control the electrokinetics-induced motions in the converging channels,. The velocity of a charged particle inside the nozzle can be expressed as
u = uEP + u EO + uDEP + u HI (5-14)
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Since DEP is of second order of the local electric field while EP and EO are of first order, they play different roles in different locations because of the continuous field strength change inside the converging geometry. Various flow regimes exist in different locations along nanonozzles, where electrokinesis, dielectrophoresis, and hydrodynamic interactions are present together but contribute differently to the motion of species. In general, the regimes inside the converging channels can be divided into three zones: Zone
I starts from the large end, where DEP is negligible and electrokinesis dominates because of the low magnitude of the local electric field strength. The regions with the stretched vortices can be included in Zone I. Zone III is located near the small end, where the magnitude of the electric field is much larger than the bulk field strength and DEP overtakes electrokinesis and dominates the motion of particles. Right outside the small end, the hydrodynamic interactions and DEP are important. Between these two ends is
Zone II, which is the transition regime, where both DEP and electrokinesis play significant roles on the particle motion. These three zones are shown in Figure 5.7.
5.3 2D DYNAMIC COMPLEXATION
5.3.1 Experimental
5.3.1.1 Materials
Suspensions of Fluoresbrite Yellow Green (YG) nanospheres were obtained from
Polysciences, Inc. 40 nm nanoparticles were used to maintain the dparticle/Dnozzle ratio
(=0.0025), close to that used in the 3D dynamic assembly process. In the latter case, the nanonozzle size is 200nm at the small end. A reasonable value for the average size of
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silica nanoparticles in the precursor is about 0.5 nm. This gives the dparticle/Dnozzle ratio
(=0.0025) similar to that in the 2D case. The nanosphere suspensions with the solid content of 0.00118% and 0.0118%were made by diluting stock suspensions (2.65% solid) by a factor of 2000 and 200 with demineralized distilled water (pH = 6.5), respectively.
5.3.1.2 Fabrication 2D microfluidic channel
2D polymeric (e.g., PET and PMMA) microfluidic channels were fabricated by first patterning the microfluidic structure on the silicon wafer by photolithography.
Negative photoresist, SU-8 50 (Microchem, Inc) was used to fabricate channels with the smallest dimension of 20 µm used in section 5.2. The PDMS mold was then made transfer the SU-8 pattern by soft lithography as described in section 5.2.1.2. The pattern on PDMS (as the mold) in this case was the negative replica of microfluidic channels.
Using PDMS as the soft mold, a hot embossing step was applied to fabricate polymer channels. The mold and polymer samples were sandwiched between two glass plates and hot embossing was carried out in a benchtop press with 9-inch wide heating platens
(Carver, Inc, IN). The operation temperature of hot embossing for PET and PMMA was
160 oC and 180 oC, respectively.
5.3.1.3 Experimental setup and imaging
The experimental setup used here is similar to the description in section 3.2 and section 5.2. Electric bias (60V/cm) was applied to two wells, which is connected to the two ends of the converging channel through large microchannels (5 mm long and 500 µm wide). Suspension of 40 nm nanoparticle was fed from the well connected to the large end. The dynamic complexation process was recorded by a 12-bit high-resolution
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monochrome digital camera system (CoolSNAP HQ, Roper Scientific®) under the control of the Metamorph® imaging software (Version 6.0, Universal Imaging CorporationTM,
Downingtown, PA). The extent of complexation was quantified by measuring the fluorescence intensity along the channel wall.
5.3.1.4 Polyelectrolyte modification
In 2D PMMA or PET converging channels, the surface modification was done before sealing with a polymer film of the same material but without surface modification.
This leaves the bottom side surface for clear microscopic observation. PAH was grafted onto the other three surfaces by reaction and/or adsorption with copious of reactant (see section 4.5.2.2).
5.3.2 Results and discussions
5.3.2.1 Polyelectrolyte distribution
In the 3D dynamic assembly, surface modification was done by immersing the whole polymer layer in the polyelectrolyte solutions. Both the internal and external surfaces of nanonozzles were modified simultaneously. PAH diffused into channels from both ends. But in 2D dynamic complexation, open channels were immersed into copious polyelectrolyte solutions for layer-by-layer deposition.
To check the difference, two pieces of PMMA substrate were done for PAH modification using these two methods, respectively. One piece was simply immersed into the bulk solution and its surface was modified using the protocol described in section
4.5.2.2. The other piece was reversibly sealed by either a PDMS block or a piece of
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scotch tape. Fluorescence labeled PAH was used this time and the fluorescence intensity was checked after the surface modification, which represented the amount of immobilized PAH on the substrate. No significant difference was found between those two methods, which indicated that uniform PAH immobilization was achieved in both approaches. Therefore, in the 2D experiments, the channel surfaces were modified first and then sealed for the further dynamic complexation study.
5.3.2.2 2D dynamic assembly process
To achieve a uniform deposition on the channel surface, the concentration of the nanoparticle suspension is critical in dynamic complexation. This is also true in the 3D dynamic assembly. In Figures 5.11a and11b, the solid content of nanoparticles in suspension was about 0.00118 wt% and the suspensioin was continuously supplied. The fluorescence intensity represents the number of nanoparticles immobilized on the solid surface during dynamic complexation. In Figure 5.11a, four curves were generated from different locations on the converging channel surface, labeling by letters (a-d) with the point “a” located closest to the small end. It is clearly shown that the dynamic complexation started from the small end and extended gradually towards the large end.
After a various period of precomplexation, there was a rapid growth stage, followed by a much slower growth stage on all four curves, indicating that two different deposition rates existed during the dynamic complexation. These two stages are named as “primary complexation” and “secondary complexation”, respectively. Beause of this two-stage complexation, the early started complexation near the small end would be caught up by that near the large end, and eventually the complexation along the whole channel surface
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would reach the equilibrium in sequence. As shown in Figure 5.11a, the first three curves reached the same level of fluorescence intensity around 40 seconds.
To better understand the evolution of this surface complexation process, the fluorescence intensity was measured along the whole channel surface at different time intervals as shown in Figure 5.11b. It again shows that dynamic complexation started from the small end. Before reaching the plateau value, the fluorescence intensity decayed rapidly from the small end to the large end. Most of the channel surface (i.e., between x/L=0.2 and x/L=0.7) had a similar level of fluorescence intensity after reaching the plateau value. This indicates a uniform coating along the converging channel surface, confirmed by the fluorescence microscopic image (Figure 5.12).
To estimate the complexation difference at these two stages, a constant growth rate is assumed in both stages. The results from Figure 5.11a are rearranged by normalize the fluorescence intensity to a new variable, θ (instant fluorescence itensity/equilibrum fluorescence intensity), which can be considered as the extent of complexation. The slopes for the best fitting lines represent the complexation rate as shown in Figure 5.13.
The ratio of the primary deposition rate and the secondary deposition rate falls in the range of 2.5~3.6. The variation of the complexation rates (both primary and secondary complexation) comes from the different deposition extent of previous deposition step
(various initial complexation and primary complexation) and how far away the complexation is to its equilibrium state.
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5.3.2.3 Dynamic complexation mechanism
The complexation between PAH and negatively charged PS colloid nanoparticles comes from the electrostatic interactions. The channel surface is initially positively charged from the polyelectrolyte, PAH, but its surface charge will slowly become negative when the negatively charged nanoparticles or silica precursors are dynamically assembled on the surface. The results given in Figure 5.11 indicate that the complexation does not start evenly everywhere. Instead, it starts from the small end and then extends gradually towards the large end. This oriented dynamic complexation leads to an extra zeta potential gradient along the converging channel surface. The non-uniform zeta potential would enhance the dissimilitude between the flow field and the electric field.
The inhomogeneous surface charge distribution and its induced flows were considered by several researchers in electroosmotic flows. A step change of zeta potential was used to simulate the surface defects in microchannels (Anderson, J. L. et al, 1985,
Long, D. et al, 1999). Further studies using undulated surface and surface with stripes of alternating charge found that either a complex recirculating flow or a more regulated rotational flow could be generated. Among all of these studies, a constant electric field was applied (Ajdari, A. et al, 1996 and Stroock, A. D., et al, 2000). Recently, simulations considering geometries with a gradual change of both the surface charge and the cross section of the channel were reported, which is close to our dynamic complexation study.
Lubrication theory or a series of long-and-narrow channel models were applied (Ghosal, et al, 2002 and Brotherton, M. C., et al, 2004). In both studies, the characteristic length scale for the variation of the cross-section was much larger than its characteristic width and the surface charge pattern was pre-specified and remained unchanged. These
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conditions are substantially different from our complexation study in both 3D and 2D converging channels (length/width < 15). However, some useful insights can be applied to explain our experimental observations.
Similar to the previous fluidic study, vortices initially exist both outside the small end and inside the converging channel. But this time, the rotation direction is reversed because the channel surface is positively charged initially (because of PAH layer), as shown in Figure 5.14a. These vortices are believed to cause the initial trapping of nanoparticles on the solid surface, especially at the small end. As mentioned previously, the center of the two vortices outside the small end are offset to the side of the solid cape and part of the vortices could also be extended into the small end. These lip vortices create low motion regions close to the small end and these regions should be the first place where many particles are aggregated and deposited on the solid surface. This initial complexation leads to the following deposition inside the converging channel. In our 3D dynamic assembly, there was a large aperture of ring shape outside the small end of nanonozzles after silica formation (Figure 4.14). The vortices and the initial complexation at the small end must have played an important role there.
Given that nanoparticles first deposit at the small end as the complexation starts, zeta potential at that location is changed. At first, the surface becomes less positively charged and such changes continue when more and more nanoparticles deposit.
Eventually, the surface would become negatively charged. During this process, the surface zeta potential at that location becomes different from the adjacent unaffected surface, leading to an inhomogeneous surface charge distribution. With such a zeta
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potential gradient, recirculation flows would appear and form a new vortex near the small end (shown in Figure 5.15).
When the second vortex is formed near the downstream region because of the zeta potential gradient along the channel, a stagnation point (or a stagnation plane if considering the depth direction and in the 3D flow) is formed at the location where it meets with the vortex already existing inside the converging channel as shown in Figure
5.15b. With this vortex pair, a relatively isolated region would appear near the channel surface. Some of the nanoparticles trapped into the recirculation flow would fall into the stagnation region. The slow motion in the stagnation region provides a much longer residence time to the nanoparticles than at other locations, leading to the primary complexation between the PAH layer and nanoparticles. With more particles being deposited, the zeta potential profile on the channel surface keeps changing. The interfacial plane of the step gradient would gradually move towards the large end, leading to the motion of the vortex interface, or the frontline of the primary dynamic complexation.
With the interfacial plane moving towards the large end, the complexation continues taking place in the region between the interfacial plane and the small end inside the converging channels because of the vortex. Compared to the primary stagnation deposition, the complexation rate caused by vortex deposition is much slower and is considered as the secondary complexation stage. Two effects make this secondary complexation slower. First of all, nanoparticles are brought near the surface only by the recirculation flow after the primary complexation. The motion of nanoparticles though much slower than in the major flow, is much faster than the stagnation region. Secondly,
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the channel surface behind the complexation front becomes less positive or even weakly negative after the primary complexation. Such surface is less effective for further deposition as the complexation approaches to its saturation state. Therefore, the interfacial plane (i.e., the stagnation region) is the most efficient location for complexation and the dynamic process in other regions are much slower.
The vortex pair caused by inhomogeneous zeta potential along the channel surface and the EOF/back pressure flow interaction forms the unique condition for the dynamic complexation. When the pressure-driven flow is carried out alone in the converging channels, the lip vortex can also be generated at the small end and a few nanoparticles were also deposited there (Figures 5.4f). But the lack of vortices inside the converging channel leads to no continuous complexation inside the channel because of the high velocity of nanoparticles. Similar situation is also held for straight channels. The fixed dimensions of cross section in the straight channel can not generate vortices inside even when electrokinetic-induced flows are applied. Deposition was observed only near entry area (Towns, J. K., et al, 1992).
5.3.2.4 Effect of feed pattern on complexation
Figure 5.11b shows two unusual peaks, one at x/L=0.1 and the other at x/L=0.75.
Both locations a very high fluorescence intensity at the very early stage of complexation.
The fluorescence images in Figure 5.12 indicate that those unusual peaks come from the deposition of the clusters of nanoparticles, analogous to large silica nanoparticles observed in the bulk reaction of silica assembly. Since clusters of nanoparticles are more easily formed in concentrated suspensions than in diluted ones, diluted suspensions
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would be more desirable for uniform complexation. Besides, the local recirculation flow or vortices there could also be less pronouced with dilute suspensions, which would slow down the deposition rate of nanoparticles near the small end. The zeta potential gradient would also be built up more slowly. Therefore, when more diluted particle suspensions is employed, more uniform coating would be expected.
In Figures 5.11, the solid content of nanoparticle is about 0.00118 wt% of the suspensions and the suspensioin is continuously supplied. When this diluted particle suspension was employed, a uniform surface coating was achieved. But the drawback of this continuous feeding is that dynamic complexation takes place slowly and the process could be time consuming. One alternative approach to speed up the process is to change the feeding pattern of nanoparticles. Instead of continuously supply of the suspensions, single or multiple dosages of more concentrated suspensions were used.
Figures 5.16a and 16b show the results after a single dose (1 µl) of nanoparticle suspension with a higher solid content (0.0118 wt%). The complexation close to the small end could be achieved the same level as in the continuous feeding case. But it became slow in other regions because of the quick exhaust of nanoparticles after the pulse of feed (Figure 5.16b). Therefore, the deposition curves at the four locations are gradually flattened but with different fluorescence intensity (Figure 5.16a). When the same dosage of the nanoparticle suspension was feeded every one minute, the deposition would continued as shown in Figure 5.17a. Compared to the complexation of continuous feeding, this multiple dosage feeding could locally accelerate the dynamic complexation and deposit more particles at specific locations, like at the small end. However, the complexation becomes more non-uniform than in the continuous feeding as shown in
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Figure 5.17b. More nanoparticles were deposited close to the small end and the number of nanoparticles along the channel surface decayed rapidly (close to an exponential decay) towards the large end. It implies that the majority of deposition occurred in the initial complexation stage. In the pulse feeding, there is not enough time to conduct the primary complexation along the channel surface in most regions before the concentration of nanoparticles became too low.
Because of the rapid deposition at the small end, the channel size at that location could be quickly reduced, which leads to a rapid increase of the local electric field. The electric field strength is inversely proportional to the cross-section area, which leads the value of E • E proportional to the fourth power of the size of the channel. If the channel size is reduced to half of its original value, the velocity component from electrokinetics is four times higher than its original value and the local dielectrophoresis would increase sixteen times. Therefore, with the reduction of channel size, the local electric field is also significantly changed in nanoparticle complexation. This geometric enhancement of the field strength will bring most of nanoparticles quickly to the small end in each dosage, resulting in a local high concentration of nanoparticles around that region. This could further enhance the nanoparticle trapping. With more and more dosages provided, particle trapping most likely becomes the main source for the continuous deposition of nanoparticles adjacent to the small end. In regions far away from the small end, this effect will be much less pronounced and the nanoparticle deposition is still believed to be the result of complexation.
The high concentration suspension used in this multiple-dosage feed could easily lead to the formation of nanoparticle aggregates, which were observed in the
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experimental images (Figure 5.18) and shown as small but sharp peaks on the fluorescence intensity curves (Figure 5.17). The presence of aggregations could attract more particles to these sites. Large aggregations could even change the local flow field, forming additional local recirculation flow, which robs nanoparticles from the adjacent region. As a result, the aggregation at these sites would grow rapidly and form nanoparticle clusters. Small clusters would lead to a rough surface after complexation while large clusters could block the entire channel during complexation. There are many ways to avoid the large cluster formaton and one simple way is to shorten the dosage interval and reduce the suspension concentration in each dose. The shorter the dosage interval is, the closer to the continuous feeding and the more uniform complexation can be achieved.
Even though a non-uniform deposition takes place in the multiple dosage feeding process, it can still be useful as it can rapidly form a thick colloid layer near one end of the channels. For 3D dynamic assembly in nanonozzles, this non-uniform complexation could reduce the size at the small end of nanonozzles much more than at the large end.
This helps increase the converging angle of naonozzles (which is the cone angle if conically shpaed nozzles are used). Such nanostructures could be more desirable for applications where both a high separation efficiency and a high flux are required based on size exclution, e.g., immunoisolation in drug delivery.
5.4 TRANSPORT IN 3D NANONOZZLE ARRAY
Polymer nanonozzles with uniform distributed conical channels can provide two important flow patterns: converging and diverging flow. The electric field enhanced
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particles transport was studied in both flow patterns with rigid nanospheres and flexible
DNA molecules as model materials. Different from the dynamic complexation study,
, nanoparticles used here have a much larger ratio of dparticle/dnozzle which results in the hindered migration. Similar to the dynamic assembly study, 2D flows were also carried out to help explain the transport phenomena observed in the 3D nanonozzle array. This time, the microchannels were scaled up to 5 µm at the small end (a scale ratio of 25).
Correspondingly, large nanospheres were used to maintain the ratio of dparticle/dnozzle closer to that used in the 3D studies. Other materials and experimental conditions were kept as much the same as in the 3D experiments.
5.4.1 Experimental
5.4.1.1 Materials
Polycarbonate track-etched membranes (average pore size: 3 µm and 200 nm) were purchased from GE Osmonics. Suspensions of fluoresbrite Yellow Green (YG) nanospheres were obtained from Polysciences, Inc. These nanosphere suspensions with the solid content of 0.00265% were made by diluting the stock suspensions (2.65% solid) by a factor of 1000 in demineralized distilled water (pH = 5.5). Bacteriophage λ-DNA
(48.5 kbp) was purchased from New England Biolabs. TE10 (Tris-HCl, 1mM EDTA,
10mM NaCl, pH=8), β-mercaptoethanol, glucose and sucrose were purchased from
Sigma-Aldrich (St. Louis, MO) and used as received. YOYO-1 fluorescent dye was obtained from Molecular Probes, Inc. Fluorescently labeled λ-DNA solutions were prepared using the same protocol as described in Chapter 3.
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5.4.1.2 Fabrication of 2D microfluidic channel
2D PDMS converging channels with the size of 5 µm at the small end were made by soft lithography using Si mold. Si mold were fabricated by photolithography and dry etching. Briefly, microfluidic structure was first patterned on a silicon substrate by photolithography, same as the description in section 5.2.1.2. The pattern was then transferred onto the silicon substrate by deep reactive ion etching (DRIE) process using an induced coupled plasma (ICP) system (Oxford 130, Oxford Instruments, Inc). The plasma etching step was performed in a mixture of SF6 (40-80 sccm) and O2 (3-8 sccm) gases under the cryogenic condition. 2D PDMS channels were then made by casting the mixture of 10:1 ratio (by weight) of PDMS precursor and curing agent (Sylgard 184,
Dow Corning) using soft lithography. The mixture was degassed in vacuum for at least half an hour and then cured overnight at 65oC. Images of 2D PDMS converging channels are shown in Figure 5.1b and Figure 5.22a.
5.4.1.3 3D experimental setup
A polymer layer with the nanonozzle array was clamped between two half acrylic cells with a silicone O-ring seal as shown in Figures 5.19a and 19b. Each cell chamber has a volume of 7 ml. The donor half-cell is filled with an aqueous testing suspension containing charged permeates while the acceptor half-cell initially contains only the distilled water or buffered solutions. All aqueous solutions were prepared with analytical grade reagents and DI water, and filtered through a 1.0 µm syringe filter (Acrodisc,
Gelman Sciences). Triton-X 100 (Sigma-Aldrich, MO) was used to facilitate the filling of nanochannels with aqueous solutions and to stabilize the polystyrene nanospheres. An
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external electric field (20 V/cm) was applied to enhance the transport of electrolyte species.
5.4.1.4 Characterization
The transport results of nanospheres were obtained by periodically measuring the fluorescence intensity of solutions (using Cytofluour) in both half-cells. A standard curve was firstly established to convert the fluorescence intensity to the nanoparticle concentration of suspensions.
To image the 3D converging/diverging flow, the entire chip was mounted onto the stage of an inverted epifluorescence microscope (TE 2000S, Nikon, Japan). The dynamics of λ-DNA conformation were observed with a 100x/1.3 NA oil immersion objective lens.
5.4.2 Transport of rigid nanospheres
For PS nanospheres with a diameter of 40 nm (dparticle/dnanonozzle=0.20), the cumulative transport in the permeate cell is shown in Figure 5.20a. The amount of transported particles in the acceptor half-cell remained almost constant in the converging flow after a slight increase at the very early stage. On the other hand, the particle concentration in the donor half-cell decreased as a function of time. This off-balance suggests that rigid nanospheres met a traffic jam inside the conical nanochannels and were rapidly stacked, building a large transport barrier to block the flow of the following nanospheres. The transport phenomena in the diverging flow, however, is completely different. The permeate concentration in the acceptor half-cell kept increasing, following
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a Fickan’s profile (i.e., the first order diffusion). Correspondingly, the particle concentration in the donor half-cell decreased exponentially at the beginning. The concentration of nanospheres decreased faster in the converging direction than in the diverging direction in the donor half-cell. But after around 7 hours, the situation was reversed with more nanoparticles left in the converging case, suggesting that nanochannels started to be stacked by the colloid rigid particles.
Similar experiments were also carried out using polycarbonate track-etched
(PCTE) membranes with the average pore size of 200 nm and 1 µm, close to the channel dimensions at the two ends of the nanonozzles. For the 200 nm track-etched membranes, there was little permeate flux in the first 10 hours. The particle concentration in the acceptor half-cell then increased rapidly. After more than half of nanoparticles have left the donor half-cell, the particle concentration in the acceptor half-cell and the permeate flux become nearly the same in both track-etched membrane and nanonozzles (Figure
5.20b). Such a delay in the track-etched membrane suggests a long residence time of nanoparticles in torturous nanochannels. Our nanonozzles, on the other hand, show a very short residence time. For the 1 µm track-etched membrane, the permeate concentration increased rapidly in the acceptor half-cell from the beginning. The nanoparticles seemed to move almost as freely as in the bulk solution.
To further explore the hindered transport through the nanonozzle array, the cumulative flux of nanospheres with various sizes was investigated in the diverging flow and plotted in Figure 5.21a. Different shapes of transport curves are shown, from Fickan to linear when the size of nanospheres changes from 40nm to 84nm, which is about half of the size of the nanonozzles at the small end. The partition coefficients of these
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nanospheres are calculated based on the data in Figure 5.21a. Here the partition coefficient is defined as the ratio of the diffusion coefficient in the nanonozzles to its value in free solution (= Dnozzle/Dbulk). The value of the diffusion coefficient in the free solution is estimated from the diffusion coefficient of nanospheres transporting through
3.0 µm-in-diameter track-etched membrane. The hindered factor is defined as the ratio of the diameter of the diffusing particles to the diameter of the nanochannels based on the small end (λ = dparticle/dnozzle). The partition coefficient of various colloid particles is plotted versus the hindered factor in Figure 5.21b. When λ = 0 (dparticle< Dnozzle/Dbulk = 1, and when λ approaches 1.0, Dnozzle/Dbulk must be zero. This has been verified by the transport result of 200 nm nanospheres through the equal sized nanonozzles, i.e., nanospheres were nearly 100% retained when λ = 1.0. The other two hindered factors shown in Figure 5.21b are 0.20 (40 nm) and 0.42 (84 nm), respectively. The partition coefficient is much less than unity and decays rapidly with the increase of the hindered factor. This result agrees with those from other researchers on hindered diffusion study (Beck, R. E., et al, 1970). Renkin correlated the hindered transport results by an empirical equation through fitting the hindered diffusion data from various species (Renkin, E. M., 1954), D h = (1− λ) × (1− 2.10λ + 2.09λ3 − 0.95λ5 ) (5-15) Dbulk Here, Dh and Dbulk represent the diffusion coefficient in the hindered diffusion and free diffusion, respectively. The first part of the equation on the right side represents the geometrical effect and the second part represents the hydrodynamic interactions on the analyte due to the proximity of the channel wall. 165 Different from the results of 200 nm and 40 nm nanospheres, the flux curve of 84 nm nanospheres (λ = 0.42) conforms to a linear relationship with time (i.e., zero order diffusion), similar to the single file diffusion (SFD) phenomena. The actual partition coefficient of the 84 nm nanoparticles is slightly higher than the prediction of equation (5-14). This suggests that nanospheres with 84 nm in diameter probably followed the sequential pass through the nanochannels due to small channel size and the greatly constrained Brownian motion. To investigate the hindered transport phenomena in the 3D nanonozzles, 2D experiments were conducted under the microscopy with converging channel size scaled up to 5 µm at the small end. Different from the cases in dynamic assembly, particles used here are much “bigger”, to maintain the same hindered factor. For example, charged and fluorescent-labeled PS nanoparticles with the diameter of 2 µm and 1 µm were used to maintain the dparticle/dnozzle ratios (=0.40 and 0.20, respectively) close to those used in 3D transport experiments (Note: with nanonozzles of 200nm at the small end, the dparticle/dnozzle ratios for 84 nm and 40nm nanospheres are 0.42 and 0.20, respectively). The force analysis discussed in Section 5.2.3 is still valid here. That is, various flow regimes exist in different locations along the nanonozzles, where DEP and electrokinesis are present together but contribute differently to the motion of species. The observed nanoparticle migration through nanonozzles can be explained by considering the combination of dielectrophoresis, electrokinesis (including electrophoresis and electroosmosis), and the hydrodynamic interactions. Because of the larger ratio of dparticle/dnozzle, the hydrodynamic interactions between the wall and particles could play a more dominant role here. 166 With the field strength the same as in the previous section (e.g., 20V/cm), no obvious vortices were observed in the diverging case with the hindered factor equal to 0.20 (dp=1.0 µm). As the result, no serious aggregation was found (shown in Figure 22c). But when the same size microspheres were used in the converging direction, obvious vortices were observed along the walls both inside and outside the channels, leading to the rapid buildup of traffic jam inside converging microchannels and a fast increase of the transport barrier. Particles were rapidly trapped in the vortices and aggregated around the small end. The actual transport pathway was gradually narrowed or even blocked by the stacked particles, as shown in Figure 22b. This difference explained the different transport behavior observed in the 3D nanonozzle array, i.e., hindered effects became more serious in the converging direction than in the diverging direction at the same hindered level. But when the hindered factor becomes higher (0.40, when 2 µm-diameter microspheres were used), the serious aggregation would occur even in the diverging case. However, particles were packed much loosely this time at the small end. 5.4.3 Transport of semi-flexible DNA For large flexible molecules (i.e., λ-DNA), different migration behaviors were observed. λ-DNA (unstained) used in this study has a radius around 0.7 µm under its supercoiled configuration; after being fully stretched, it (unstained) has a contour length of 16.3 µm and a hydrodynamic radius of 2 nm. To study the conformation dynamics of λ-DNA in the two classical flows, a dilute solution (~ 0.03 µg/ml, about 10-3 of the critical concentration at which the macromolecules completely fill the space without overlapping) of λ -DNA in Tris-EDTA buffer was used and labeled with fluorescent dye. 167 The channel array was placed in a nanofluidic platform as shown in Figure 5.23a. The DNA solution was loaded to the cathode side, while the anode side was loaded with the buffer solution only. Due to the negative charge, the DNA migrates from cathode to anode. The exit of DNA from the channel was captured using a Nikon inverted epi- fluorescence microscope mounted underneath the platform, as shown in Figure 5.23b. Figures 5.24a and 24b show the snapshots of the DNA movement through the nanonozzle array in both converging and diverging directions. The amount of DNA molecules at each snapshot indirectly reflects the migration rate. At each instantaneous moment, significantly fewer DNA molecules were observed through nanonozzles in the diverging direction than in the converging direction. Compared to the size of nanonozzles at the small end (200 nm), the coiled λ-DNA molecules have a much larger radius of gyration. Because of the deformable nature of the DNA chain, DNA molecules could be stretched to long, worm-like shape with a much smaller radical size. Different from the rigid PS nanospheres, these DNA molecules could perform slide motion and rapidly pass through the nozzles. The migration rate of DNA depends on the dimensions and geometry of nanochannels, the physical characterization of DNA, and the electric field strength. The degree of DNA stretching depends upon the flow strain rate and the relaxation time of the molecule conformation. The 2D study provides a direct evidence of DNA stretching even though the microscale channels reduce the contribution of hydrodynamic interactions near the channel surface to the DNA deformation. Figure 5.25a shows the dynamic process of an individual λ-DNA molecule being stretched to different extents in the converging channel. In the converging direction, acceleration is always present along the DNA 168 migration path. Such acceleration ensures stretching of DNA molecules for easy entry and to maintain their stretching configuration inside the channel. The gradual converging geometry provides a high extensional rate along the length direction and a similar residence time on each streamline. Therefore, more uniform stretching of DNA chains was observed in the 2D converging channels, compared to the results in the single cross- slot geometry (refer to Chapter 3). From this observation, we may conclude that DNA molecules kept their stretched and worm-like configuration all the time when passing through 3D nanonozzles. Because its stretching configuration with a diameter of only 2 nm, DNA molecules could conduct sliding motion inside the nanonozzles and meet with only very low resistance. In the diverging direction, on the contrary, deceleration occurs along the DNA migrate path. A positive velocity gradient exists only at the entry of nanonozzles. Consequently, DNA molecules experience only gentle stretching when entering the constrained channels and meet immediate deceleration in the expansion region. The head of the polymer chain moves slower than its tail, helping the molecular chain to coil back. Such dynamic deformation was verified in the microscale 2D diverging flows, as shown in Figure 5.25b. As mentioned earlier, this dynamic relaxation process is expected to be limited by the geometry restrain in nanonozzles. The strong interactions between the DNA chain and the channel surface would hurdle DNA from further relaxing to its super- coiled configuration. Therefore, DNA molecules experience a higher steric hindrance resistance and a longer hindrance time inside nanonozzles. Such differences between the two migrating directions explain the different migrate behaviors (e.g., numbers of DNA in each snapshot) mentioned previously (Figure 5.24). 169 Different from the stretching in the cross-slot microfluidic geometry, the 2D converging geometry used here provides DNA stretching all the way along the channel. The relaxation of polymer chain is delayed until it escapes the converging region. The 2D converging channel itself could be a useful tool to achieve uniform DNA stretching, which is desirable in the study of the dynamic complexation between DNA molecules and other materials (e.g., proteins and functional nanoparticles). Multi-functional species could also be grafted onto the DNA chains in such flow geometry. 5.5 SUMMARY A 2D dynamic complexation study was carried out using the same procedure and same geometry in the 3D dynamic assembly. The system was scaled up to microchannels with 20 µm in diameter at the small end (scale ratio is 100). Generally speaking, electroosmotic (EO), electrophoretic (EP), dielectrophoretic (DEP), and hydrodynamic interactions all play important but different roles in the dynamic assembly process, depending on the locations inside the channels. EP is always important inside or outside the converging channels while EO becomes dominant inside the channels. DEP and hydrodynamic interactions are more important in the regions near the small end. As a result, vortices are observed both inside (close to the channel surface) and outside (at the small end of the converging channels). The interactions between EO and the induced back pressure flow are responsible for the formation of internal vortices, while DEP and hydrodynamic interactions are the reasons for the external ones. The dynamic complexation was also carried out and quantified by recording and measuring fluorescence intensity along the channel wall. The results exhibit that the 170 complexation starts from the small end of the nozzle and extends gradually towards the large end. Continuous feeding of diluted suspensions offers a uniform deposition on the surface, while multiple dosages feeding can result in a rapid reduction of channel size. A two-stage complexation phenomenon was observed, which is believed to be associated with the dynamic development of a vortex pair in the channel. The primary complexation occurs when a non-uniform zeta potential surface, forms a local stagnation region between two vortices for fast complexation. The secondary complexation is a result of slow deposition caused by circulation flows in the vortex. The electric field enhanced particle transport in both 3D and 2D polymer nanonozzle arrays was studied with rigid nanospheres and flexible DNA molecules. The diverging flow shows a self-clean function while nanonozzles are easily clogged in the converging flow for rigid colloid nanospheres. With the increase of analyte size, the hindered transport becomes apparent until completely blocking when the channel size is closer to the analyte size. But for flexible polymers (such as DNA), the converging flow could stretch the DNA chains to achieve easy pass of molecules with the equilibrium size much larger than the channel size. Such nanonozzle arrays may serve as a useful platform for biomolecule separation and sequencing, controlled drug and gene delivery. 171 8 2 Species ζ , mV µ EP , ×10 m /(sV) 40 nm PS -69.18 -5.41 84 nm PS -46.72 -3.68 200 nm PS -46.92 -3.67 700 nm PS -53.31 -4.17 2 µm, PS carboxyl-modified* -89.78 -7.02 3 µm PS -9.95 -0.78 λ-DNA -5.10 -0.40 * The solid content is 0.0053%, 1/500 from the stock suspension. Table 5.1 Zeta potential and mobility of Fluoresbrite Yellow Green (YG) polystyrene micro-and nanospheres and λ-DNA. The solid content of micro/nanospheres is 0.00265%, 1/1000 from the stock suspensions unless is specified mentioned and the concentration of λ-DNA is about 0.03 µg/ml. 172 (a) 100 µm (b) Figure 5.1 Images of 2D converging channels. The width of the small end is 20 µm in (a) and 5 µm in (b). 173 (a) (b) (c) Figure 5.2 The flow characteristics inside the converging channel by electrokinetics- induced flow (a), pressure-driven flow (b), and electrokinetics-induced flow in a straight channel (c). 174 (a) 0.01 700 nm, diluted 100 folds, run#1 700 nm, diluted 100 folds, run#2 700 nm, diluted 1000 folds, run#1 700 nm, diluted 1000 folds, run#2 0.001 Viscosity, PaS 0.0001 1 10 100 1000 Shear rate, 1/s 0.1 (b) 40nm, diluted 100 folds 40nm, diluted 1000 folds 40nm, diluted 10000 folds 0.01 Viscosity, PaS 0.001 0.0001 1 10 100 1000 Shear rate, 1/s Figure 5.3 The rheology data of PS suspensions of different dilute folds: (a) 700 nm, diluted 1/100~1/1000 of the stock concentration and (b) 40nm, diluted 1/100~1/10000 of the stock concentration. 175 (a) (b) (c) (d) (e) (f) Figure 5.4 The flow characterization outside the converging channels in the electrokinetics-induced flow (a, b): 40 nm PS nanospheres (a) and λ-DNA molecules (b), outside the straight channel with the same size of the small end of the converging channel (c), and in the pressure-driven flow (d, e, f): Re=1.0 (d), Re= 10 (e) and Re=250 (f). The arrows indicate the migration direction and the scale bars represent 20 µm. 176 (a) (b) (c) (d) Continued Figure 5.5 The flow characteristics of electrokinetic-induced flows outside the small end of the diverging channels (a, b): E= 60 V/cm (a) and 120 V/cm (b), and pressure-driven flows outside the small end of the diverging channels (c, d): Re= 100 (c) and Re= 500 (d), and straight channels (e, f): E= 60 V/cm (e) and 120 V/cm (f). The scale bars present 20 µm. 177 Figure 5.5 continued (e) (f) 178 30 20 10 0 -10 -20 -30 Zeta Potential, mV PSS modified PMMA -40 PAH modified PMMA PMMA Glass slide -50 -60 34567891011 pH Figure 5.6 Zeta potential of various polymer substrates. 179 180 Figure 5.7 The electric field strength profile in a 2D converging channel and zone division based on the dominating forces. 180 14000 converging flow (experimental) converging channel (predicted) 12000 10000 m/s 8000 µ straight channel (20 µm wide, predicted) 6000 Velocity, straight channel (1300 µm wide, predicted) 4000 2000 0 -1.5 -1.0 -0.5 0.0 0.5 x/L Figure 5.8 The experimental data of lateral velocity profile measured along the centerline of a 2D converging channel. The solid lines are the simulated curves for both converging and straight channels with the channel size the same as the two ends of the 2D converging channel. The small end of the converging channel is located at x/L=0 and the large end is at x/L=-1.0. 181 (a) EOF PDF Velocity Velocity (b) EOF PDF Figure 5.9 Different velocity profiles including a bi-directional flow in a straight channel (a) and the vortices in a converging channel (b) after the superposition of the electrokinetic-induced flow (plug flow) and the induced pressure-driven flow (parabolic velocity profile). 182 (a) (b) Continued Figure 5.10 The contour of E2 in the converging channel: (a) the whole image and (b) the local image near the small end and (c) the ratio of dielectrophoretic mobility to electrokinetic mobility along the centerline of PDMS, untreated and PAH modified PMMA 2D converging channels. Particles used here are 40 nm PS nanospheres. The small end of the converging channel is located at x/L=0 and the large end is at x/L=-1.0. 183 Figure 5.10 continued (c) 6 4 PDMA surface (E*E)/Ell δ )ll EO PMMA surface µ - EP µ /( 1 PAH surface DEP µ 0 -2 -1 0 1 x/L 184 (a) 1500 a b c d 1000 a bc d 500 Fluorescence Intensity 0 0 1020304050 Time, seconds (b) 3000 0 sec 15 sec 30 sec 2500 35 sec 40 sec 45 sec 2000 1500 1000 Fluorescence Intensity 500 0 0.0 0.1 0.2 0.3 0.4 0.5 0.6 0.7 0.8 0.9 1.0 x/L Figure 5.11 The growth curves (a) and the concentration profile along the converging channel surface (b) in dynamic complexation process with the solid content of 10-5 in the suspension. The fluorescence intensity near the channel surface indicates the amount of nanoparticles deposited. In (a), letters (a-d) label different locations on the converging channel surface, starting from “a” on the small end, and in (b), the small end of the converging channel is located at x/L=0 and the large end is at x/L=1.0. 185 (a) (b) (c) (d) (e) Figure 5.12 Snapshots of dynamic complexation using 40 nm fluorescence-labeled PS nanospheres (10-5 solid content) in 2D PAH-modified converging channel by continuous feeding. The elapse time has a unit of second. The scale bar represents 100 µm. 186 1.0 k2a=0.024 0.9 0.8 0.7 k2c=0.023 0.6 θ 0.5 0.4 k1a=0.088 k1c=0.057 0.3 a c 0.2 0.1 0.0 0 1020304050 Time, second Figure 5.13 The extent of two-stage dynamic complexation. θ is equal to the instant fluorescence intensity/equilibrium fluorescence intensity. The slopes, k1 and k2 represent the complexation rates for primary and secondary complexation, respectively. Here only curves “a” and “c” are shown for clear image purpose. 187 (a) (c) (b) Figure 5.14 The flow characterization of the startup of dynamic complexation with converging channel surface initially positively charged (a). As comparison, flow patterns at the outlet (b) and inside (c) a negatively charged converging channel are also shown. The arrows indicate the migration direction and the scale bars represent 20 µm. 188 (a) (b) ζ + ζ + 0 x/L 1.0 0 x/L 0.1 (c) (d) ζ + 0 x/L 0.1 0 x/L 1.0 ζ − Continued Figure 5.15 The schematic of dynamic complexation mechanism: (a) startup with the surface uniformly positively charged, (b) the small end becomes less positively charged, (c) vortex pair appear with the small end becoming negatively charged, (d) two zones are formed with the negatively charged zone expands and the positively charged zone retreats, and (e) completely negatively charged zone. The formation of vortex pair induced by the zeta potential gradient is explained by the schematic in (f) and exhibited by the streak image of dynamic complexation experiment (g). 189 Figure 5.15 continued (e) ζ + x/L 0 1.0 ζ − (f) ζ (g) Vortex I Stagnation point Vortex II 190 1500 (a) a b c d 1000 a b c 500 Fluorescence Intensity d 0 0 102030405060 Time, seconds (b) 3000 0 sec 2500 10 sec 20 sec 30 sec 40 sec 2000 50 sec 1500 1000 Fluorescence Intensity 500 0 0.0 0.1 0.2 0.3 0.4 0.5 0.6 0.7 0.8 0.9 1.0 x/L Figure 5.16 The growth curves (a) and the concentration profile along the converging channel surface (b) in dynamic complexation process by single dosage feeding with the solid content of 10-4 in the suspension. The fluorescence intensity indicates the amount of nanoparticles deposited. In (a), letters (a-d) label different location on the converging channel surface, starting from “a” on the small end and in (b), the small end of the converging channel is located at x/L=0 and the large end is at x/L=1.0. 191 (a) a 4000 a b b c d e f 3500 d 3000 c 2500 2000 1500 d Fluorescence Intensity Fluorescence 1000 e f 500 0 0 100 200 300 400 500 600 700 Time, seconds (b) 4000 0 sec 3500 1 min 3 min 5 min 3000 7 min 2500 2000 1500 Fluorescence Intensity 1000 500 0 0.0 0.1 0.2 0.3 0.4 0.5 0.6 0.7 0.8 0.9 1.0 x/L Figure 5.17 The growth curves (a) and the concentration profile along the converging channel surface (b) in dynamic complexation process by multiple dosage feeding with the solid content of 10-4 in the suspension. The fluorescence intensity indicates the amount of nanoparticles deposited. In (a), letters (a-f) label different location on the converging channel surface, starting from “a” on the small end and in (b), the small end of the converging channel is located at x/L=0 and the large end is at x/L=1.0. 192 (a) (b) (c) (d) (e) Figure 5.18 Snapshots of dynamic complexation using 40 nm fluorescence-labeled PS nanospheres in 2D PAH-modified converging channel by multiple dosage feeding. The elapse time has a unit of second. The scale bar represents 100 µm. 193 (a) - + Nanonozzle Array (b) Figure 5.19 Experimental setup for transport study in 3D nanonozzle arrays: schematic (a) and experimental setup (b). 194 (a) 100 90 80 70 60 50 40 30 Percentage Transport 20 10 0 0 5 10 15 20 25 30 35 40 45 50 55 Time, hour Donor, diverging Acceptor, diverging Donor, converging Acceptor, converging Continued Figure 5.20 Plots of electrokinetically hindered transport of 40 nm PS nanospheres (a) in a nanonozzle array with the small end diameter of 200 nm: circular symbols for nanospheres transport from the small end to the large end; triangle symbols for nanospheres transport in the opposite direction; (b) in a nanonozzle array with the small end diameter of 200 nm (reverse triangle), 200 nm track-etched membrane (square) and 1 mm track-etched membrane (circle). 195 Figure 5.20 continued (b) 100 90 80 70 60 50 40 200nm STI 200nm PCTE 30 1 µm PCTE Transport Percentage 20 10 0 0 102030405060 Time, hour 196 (a) 100 90 40 nm 80 84 nm 200 nm 70 60 50 40 30 Transport Percentage 20 10 0 0 5 10 15 20 25 30 35 40 45 50 55 60 Time, hour (b) 1.0 0.8 2 3 5 D/Dbulk = (1-λ) x (1-2.10λ+2.09λ -0.95λ ) bulk 0.6 (prediction of Renkin's equation) /D 0.4 nozzle D 0.2 0.0 0.0 0.2 0.4 0.6 0.8 1.0 λ (=d /d ) particle nozzle Figure 5.21 Plot of electrokinetic hindered transport of various sizes of PS nanospheres in nanonozzle arrays with the small end diameter of 200 nm (a) and the hindered factors of different size PS nanospheres (b). The line is the prediction of Renkin’s equation. 197 (a) (b) (c) (d) Figure 5.22 Microscopic images of 5-µm converging channel: before experiment (a), after hindered migration study in the converging direction (b) and in the diverging direction (c, d). The hindered factors in (b, c, and d) are equal to 0.20, 0.20, and 0.40, respectively. The scale bar represents 20 µm. 198 (a) Nanonozzle Array Nanofluidic Support film edge Platform Donor side Acceptor side (b) Microscope with camera Figure 5.23 Experimental setup (a) and diagram of DNA migration through channel array under electric field (b). 199 (a) 130 100 (b) 130 100 Figure 5.24 Snapshots of λ-DNA molecules passing through nanonozzle array in the converging (a) and diverging (b) directions, respectively. 200 (a) (b) 5 µm Figure 5.25 Different stretching extents of λ-DNA in the 2D converging channel (a) and diverging channel (b). The solid dots in the cartoons represent the positions of DNA molecules in each image, but not drawn in scale for clear purpose. The scale bar presents 5 µm. 201 CHAPTER 6 CONCLUSIONS AND RECOMMENDATIONS 6.1 CONCLUSIONS The purpose of this study is to fabricate polymer nanofluidic devices, which can provide better understanding of fluid transport at the micro-/nanoscale and tools for highly valuable biomedical applications, such as cell immunoisolation and drug/gene delivery. Single or multiple cross-slot microfluidic platforms were designed to generate elongational, shear and rotational flows by using different surface charge patterns on the channel surfaces and various electric bias configurations in the liquid storage reservoirs. Fairly homogeneous and two-dimensional flows can be readily generated by electrokinetics in low-viscous fluids (e.g., aqueous solutions), which were successfully identified using fluorescence labeled polystyrene microspheres as the tracer. The conformational evolution of single λ-DNA molecules in the electrokinetic flow were investigated and it was found that the initial conformation and the residence time of DNA molecules played important roles in determining the extent of DNA stretching. The strong interactions between charged analytes and channel surface made it difficult to 202 generate pure rotational and shear flows other than the elongational flow in the single cross-slot design. By extending the cross-slot design to five cross-slots, the situation is significantly improved because of the independence of flow patterns to the particle charge density, especially for shear and rotational flows. However, non-uniform stretching and various configurations of DNA still exists in those flows. A new design, nanonozzles, can provide uniform and controllable DNA stretching because of its continuous contraction geometry. A low-cost approach, Sacrificial Template Imprinting (STI), was developed to massively produce the polymer nanonozzle arrays. Each nanonozzle is 3 µm high with the channel diameter on the sharp end as small as 80 nm. Using the polymer sacrificial template successfully avoids any structure damage or defects during the de-molding. In conjuction with the surface modification and silica synthesis on the channel surface, the channel size can be further reduced and the polymer structure can be reinforced. The 2D converging microfluidics was studied to investigate the mechanism of dynamic assembly. As a result, vortices are observed both inside (close to the channel surface) and outside (at the small end) of the converging channels. The interactions between the electroosmotic flow and the induced back pressure flow are responsible for the formation of internal vortices, while dielectrophoresis and hydrodynamic interactions are the reasons for the external ones. The dynamic complexation was also carried out in the 2D microfluidics and was quantified by recording and measuring the fluorescence intensity of 40 nm PS nanoparticles deposited along the channel wall. The results exhibit that the complexation starts from the small end of the nozzle and extends gradually towards the large end. 203 Continuous feeding of diluted suspensions offers a uniform deposition on the surface, while multiple dosages feeding can result in a rapid reduction of channel size at the small end. A two-stage complexation phenomenon was observed, which is believed to be associated with the dynamic development of a vortex pair in the channel. The primary complexation occurs in a local stagnation region between two vortices resulting from the non-uniform zeta potential surface. The secondary complexation is a result of slow deposition caused by circulation flows in the vortex. Polymer nanonozzle arrays can provide two important flow patterns: converging flow and diverging flow. The electric field induced particle transport was studied in both 3D and 2D nozzles with rigid nanospheres and flexible DNA molecules. The diverging flow shows a self-clean function while nanonozzles are easily clogged in the converging flow for rigid colloid nanospheres. With the increase of the analyte size, the hindered transport becomes apparent until the channel is completely blocking when the channel size is closer to the analyte size. But for flexible polymers such as DNA, the converging flow could stretch the DNA chains to achieve easy pass of molecules with the equilibrium size much larger than the channel size. 6.2 RECOMMENDATIONS 6.2.1 Electrokinetic-driven flows For the single converging/diverging flow, the flow patterns could be either pure extension (when the channel edge follows the flow streamline), shear or recirculation flows and the flow patterns could be controlled by adjusting the geometry, converging 204 angle and surface properties. When multiple channels are connected in series, an entropic ratchet microdevice can be formed. A periodic asymmetric zig-zag potential energy is generated, considering the non-linear velocity profile (Figure 6.1). Array-type nanofluidic circuits can also be designed to achieve high throughput. Current microscopy imaging technologies have to illuminate a large volume of liquid in both lateral- and vertical-directions. This makes local imaging a very challenge task. The large background reflection from the wall and overshadow particles also make it difficult to study the hydrodynamic interactions and electrostatic interactions, which could be dominant in nanofluidics. Confocal laser scanning microscopy could improve the imaging quality, and its major drawback – i.e., the slow scan speed –has recently been largely improved with the application of the micro-pinhole or micro-slit spin-disk technique. This new technique could raise the scan speed to 1,000 fps, capable of catching the motion of high-speed tracers. It should be able to carry out the 3D PIV measurements in microfluidics – for instance, the 3D images of DNA deformation and dynamic complexation. The completed velocity profile mapping could be very useful for providing more precise explanation of the behavior of nanoparticles in various flow patterns. The 3D images of DNA deformation can provide more accurate data when interactions (hydrodynamic interactions, electrostatic interactions, etc) are considered in the DNA deformation study. Converging/diverging fluidic channels can also used as genomic or proteomic biosensors. The special potential profile in the converging channels could be applied to manipulate and immobilize biomolecules at the local region, which could dramatically increase the detection sensitivity. Examples include performing single-DNA mapping or 205 even sequencing with nanochannels consisting of a converging region and an elongation region. The converging region is used to stretch the DNA for the full exposure of active sites for transportation and anchoring with the fluorescently tagged nucleotides. The elongation region is used for the detection and identification of the DNA sequence. Decoding could be completed by hybridizing small gene fragments onto the major chain of genes with a known sequence. Fluorescent tags (e.g., DNA binding proteins and peptide nucleic acids, or PNAs), or quantum dot tags can be applied to stain specific gene sites. By detecting the population of illuminated signals from the probes complexed on the molecular chains, the length and sequences of the DNA can be determined by performing multiple runs with different tags. The results could also be verified by measuring the mobility change before and after the hybridization. The 2D nanochannels can facilitate single-molecule stretching and single-molecule sequencing. With multiple nanochannel platforms, statistically significant results can be obtained. Both DNA tag techniques and the velocity of DNA migration in the channels play important roles in the resolution of DNA mapping. There are several tag techniques (including a bar-code technique) available and worth to be investigated to find a robust way with low coefficient variations (CVs). 6.2.2 Dynamic complexation Electrokinetically driven flows often produce a plug-like velocity profile. In conjunction with externally tunable surface forces at the nanometer scale, 2D micro- and nanonozzles could overcome the Brownian motion and relaxation forces of biomolecules and nanoparticles. These molecules and particles could be caged in the fluid or near the 206 solid surface with a desirable shape and orientation and moved along a pre-specified “assembly line” with controlled velocity and displacement. Together with synthetic chemistry and biological complexation, the dynamic conjugation platform would allow continuous production of well-defined, multifunctional 3D biomimetic nanostructures and devices through polymer-biomolecule and polymer-nanoparticle-biomolecule conjugation. Such biologically active nanoscale structures and devices may greatly enhance the clinical realization of extensive genomics and proteomics research results for the treatment of cancers and chronic, infectious, parasitological and central nervous diseases, as well as vaccine delivery. Examples for research include the complexation of polymer-DNA conjugates, chromosomal DNA packaging, and fast hybridization of DNA (Figure 6.2). Efficient complexation of macromolecular therapeutics, such as DNA and proteins using polymers and/or nanoparticles, has promising applications in non-viral vector gene therapy. The structure of the biomolecule-cationic-carrier complex greatly affects the biomolecular protection and compaction, as well as its tissue bioactivity, cellular entry, intracellular transport, nuclear membrane penetration, and sustained release. The large size ratio of DNA with the carriers makes its complexation difficult to control, which often results in extremely low efficiencies to the target site. 2D homogenous flows (including extensional and shear flows) could be generated by electrokinetic forces with the proposed converging geometries in section 6.2.1. The fluid velocity can be manipulated readily by varying the applied electrical field, surface charge density of the solid, and solution properties. The presence of EP responses can be used to manipulate the flow field. The nonlinear electric gradients inside the converging channels 207 provide a good environment for stretching DNA. This active nanofluidic device can be used for high-speed complexation by binding biomolecules with polymers and nanoparticles. Polyelectrolytes, such as PEI, can be used as the condensing regents for gene therapies. Dynamic complexation is determined by the local reaction rate (the local concentration of DNA and nanoparticles), the residential time and the conformation change of polyelectrolytes. One or several such converging channels can be assembled together to achieve a desirable complexion ratio. The hybridization of DNA may also benefit from the stretching configuration of the semi-flexible molecular chains. Current DNA hybridization is a time-consuming process and is limited by molecular diffusion. Recent scientific discoveries have exhibited that the internal hybridization sequences could be accelerated greatly after unfolding and stretching the DNA molecules under shear flow (Haber, C. et al, 2000). The deformation of molecules helps the extra exposure of complementary targeting sites for easy accessory. With the converging nanofluidic channels and arrangements, the extent of DNA stretching can be controlled by the applied electric field, the geometry and the numbers of a series of the same units. The unfolded DNA super structures may reduce the steric barrier significantly and shorten the diffusion distance, while electrophoresis may enhance the contact of nucleic acid probe with DNA molecules. The possible weak Joule heating close to the thrust region may also be helpful for the hybridization. The hybridization can be carried out with well-known nucleic acid probes, for instance, peptide nucleic acids (PNA). Synthetic PNA analogs containing a neutral 2- (aminoethyl)glycine backbone are linked by peptide bonds. PNA oligomers bind highly sequence-specifically. The binding is quite independent of the ion strength of solution 208 and could be grafted easily with biotin or amino acids. Through the biotin-streptavidin affinity binding, quantum dots or other tags could be attached to the PNAs. 6.2.3 Nanotip and nanonozzle drug/gene delivery Polymer nanonozzle and nanotip produced by Sacrificial Template Imprinting can be very useful for genes/drugs delivery. The array configuration of both nannozzles and nanotips may allow the simultaneous injection into a large population of cells, and/or the delivery of a variety of samples into a single cell. The microfluidic platform integrated with the electrokinetics-driven flow can eliminate the need of extruders in microinjection. The polymer-based structures and highly uniform and precise arrays provide a low cost, easy, safe and precise approach for drug/gene delivery. 6.2.3.1 Nanonozzle array based drug/gene delivery In nanonozzles, the conically shaped flow channels provide a great potential gradient when an electric bias is applied. This results in an efficient way to either accelerate the rigid carriers (e.g., liposome particles, quantum dots (Qdots) or gold particle conjugated genes) to a high momentum or to stretch a flexible gene containing biomolecules into a long, worm-like shape, with the radical size in several nanometers being such that genes can be delivered into cells by temporarily breaking through the cell membrane. Since the radical sizes of carriers and stretched genes are comparable to the natural pores on a cell membrane, the damage to the cell is significantly minimized. When integrated onto a microfluidic platform, it is quite feasible to inject diverse samples 209 sequentially into the same cell at a desirable rate, frequency and location. The response of a single cell to the dosage and time can be evaluated readily. The concept of a nanonozzle array cell patch can be used in two different ways for the delivery of drugs or genes into cells and/or tissues, as shown in Figures 6.3a and 6.3b. In Figure 6.3a, cells are placed at a short distance away from the nanonozzle patch surface, such that they would experience a very low electric shock during electrophoresis. The distance of cells to the small end of nanonozzles is controlled by placing a spacer between them. Drug/gene delivery relies on the momentum gained in the converging channels. Since this method relies on the electrokinetic (EK) movement of particles, the technique can be called an “EK Gun.” In this case, a high voltage and large electrophoresis time can be applied, since cells are not subject to any significant electric field strength. In Figure 6.3b, cells are in contact with the outlet of the nanonozzle cell patch. Around the outlet area, the cell membrane will experience very strong, localized electrophoresis where an electric bias is applied. This, together with the electrophoretic mobility gained in the converging channels, would deliver the drug/genes into cells. Lower voltage and short duration (i.e., pulses) should be used in the strategy to avoid cell lysis. For small genes or drugs, conjugated rigid particles (e.g., quantum dots and gold nanoparticles) can be used as the carriers. While traveling inside the converging nanonozzles, these carriers are able to gain a high enough momentum to overcome the cell membrane resistance, so drugs and/or genes can be delivered into cells. After optimizing the operation parameters, this method can be gentle enough that the cell membrane is able to completely recover after a short period of delivery time. Large genes have long and flexible polymer chains. They often present in a supercoiled configuration 210 with the radius of gyration in micrometers. Their flexible chains can be stretched with external forces to form long and worm-like “nanowires” – with the radius of gyration around 2 nm – which can easily pass through the intrinsic pores on a cell membrane. The great velocity gradient inside the converging channel provides enough stress to stretch gene molecules from their equilibrium coiled conformation to their stretched conformation. The extent of stretching depends upon both the flow stress and the relaxation time of the molecule conformation. 6.2.3.2 Nanotip array based drug/gene delivery Nanotips also can be used to deliver molecules, such as nucleic acids, proteins or other chemicals (e.g., drugs), to the nucleus, or maybe even to carry out cell surgery (shown in Figure 6.4). Nanotips, after being conjugated with drugs/genes, can penetrate into cells or even cell nuclei with a gentle force. The aperture of nanotips can carry a specific dosage of drugs/genes. A short penetration of the tiny nanotips would not cause permanent damage to cells, while drugs/genes would be left inside by dissolving in the surrounding medium when the rest of the nanotip is pulled from the cell membrane. The appropriate material and shape of nanotips are critical for both efficient delivery and a low cell lysis rate. The polymer nanotips made using the STI approach can be 50-300 nm in diameter and 3-10 µm long. Their small, conically shaped nanotips are important for gentle invasiveness and successful insertion into the cell. More importantly, the nanotip cell patch does not require the pullout step. Drug/genes are left inside the cell when the tip end of nanotips dissolves in the cell medium. Drug/genes can be bound onto nanotips by 211 either directly conjugating on the surface of the nanotips (Figure 6.4c) or precipitating as the aperture of the nanotips (Figure 6.4b). The selection of binding strategies will vary with the materials to be delivered. In general, genes can be conjugated by a covalent bond or van der Waal forces (e.g., avidin-biotin affinity binding) because of their simple and similar structures. Drugs can be fabricated as a part of the aperture. Besides, different drugs can be loaded by embedding them at different locations of nanotips, as shown in Figure 6.4b. The geometry of nanotips and their accurate manipulation can prevent lethal damage to the host cells, while the short and shallow penetration is enough for delivery into the cell or even its nuclei. Like nanonozzle cell patches, nanotip arrays also allow simultaneous treatment of a large population of cells. The dosage of drugs/genes can be controlled by the total amount of samples and the penetration depth of the nanotips. If necessary, multiple dosages or various delivery materials can also be injected into the targeted cells. 6.2.4 Flow assisted organization of nanostructures In Section 4.5 and Chapter 5, 3D and 2D micro/nanofluidics and dynamic assembly were studied and some phenomena can be used to generate novel nanostructures by flow assisted self-organization. If the goal of dynamic assembly is to enhance the mechanical strength of nanostructures and to reduce the channel size, a uniform deposition along the entire surface is desired. However, this dynamic assembly concept can be extended to generate various useful nanostructures. Instead of being the final products, the nanochannels themselves may serve as a template in this case. 212 6.2.4.1 Nanostructures with multiple functions In some applications, local surface modification is desirable. The change of local zeta potential has already been shown in Section 5.4 with multiple dosage feeding. Beside the various feeding approaches, other methods, like periodic electric pulse, can also achieve the same result. To enhance the non-uniform deposition of silica or nanoparticles, the non-uniform deposition of PAH on the channel surface is necessary. In principle, this can be carried out in the same way as in the multiple dosage or periodic electric pulse feeding. The first material with a non-uniform deposition could affect the following depositions in two ways: the small end will have the largest quantity of deposition of the second polyelectrolyte (e.g., PSS) because of the non-uniform deposition of the first material. When PSS has saturated the first layer at the small end, other regions are not saturated yet. This makes the deposition of the third layer (e.g., another layer of PAH) much less efficient in those regions because of the non-uniform deposition of the second layer. With such layer-by-layer deposition, the non-uniformity deposition of polyelectrolyte could be amplified after multiple-layer deposition. This could form a surface with two regions of different surface properties. If the functional groups in one region are finally terminated (e.g., by grafting other materials or by changing the deposition conditions, like pH), this non-uniform surface modification could be performed continuously in the second region. Nanostructures with multiple functional surfaces could eventually be achieved in this way, as shown in Figure 6.5a. Polyelectrolytes are not the only candidates for this approach, especially for the top layer. Other materials, like various functional biomaterials, quantum dots, carbon nanotubes can be grafted on the surface of different regions to form useful nanostructures. One example 213 is to graft functional group as the active recognition sites at the small end for the targeting purpose. If the initial nanostructures are not necessary for the final product, the polymer material can be dissolved to release multiple functional hollow nanostructures. 6.2.4.2 Drug/DNA loading in naocapsules The phenomena of hindered transport can also be a useful tool for drug/gene loading in nanostructures. As mentioned in Section 5.5, species with a large hindered factor are easily clogged when the migration is carried out in the converging direction. This could be useful for loading multiple drugs with structures as shown in Figure 6.4b. The possible procedure is proposed and schematically shown in Figure 6.5b. Materials, which are able to degrade or dissolve in certain conditions, can first be clogged at the small end as the cap of the nanocapsule. Various drug materials could be sequentially loaded into the nanonozzles through the converging flow. To control the quantity of each drug, the periodic dosage loading mentioned early can be applied. If used for targeted drug delivery, recognizing sites (e.g., specific proteins with functional groups) can also be grafted to the surface of cap or the small end of nanonozzles. Because flexible DNA can be stretched to pass through the converging channel, the loading of DNA in nanonozzles can be performed by the diverging flow. As described in Section 5.4.3, deceleration occurs after DNA molecule enters the nanonozzle. This helps the molecular chain to coil back while the strong interactions between polymer and channel surface would hurdle the relaxation. Combining with the strong steric effect, DNA molecules could be stuck in nanonozzles, as shown in Figure 214 6.5c. These loaded gene nanocapsule can be further integrated into artificial nanorobots, mimicking virus for efficient delivery of therapeutic genes. 215 (b) (a) converging angle α (c) (d) (e) Figure 6.1 Various converging/diverging geometries and their combinations (a-c); schematic of polymer conformation transition in converging/diverging geometry (d), Electric field profile in a series of converging channels (e). 216 (a) (b) (c) Figure 6.2 Schematic of dynamic DNA complexation or hybridization (a), DNA conjugates with diverse functional groups or nanoparticles (b), replica of helix structure (c). 217 Coiled Particles conjugated DNA with genes Stretched DNA Cell Cell 218 Cell Cell Nanonozzle Array Nanonozzle Array (a) (b) Figure 6.3 Schematic of two strategies of polymer nanonozzle array cell patch for gene/drug delivery (a) for drugs and small genes; (b) for large genes. 218 (a) (b) I Drug III Nanotip array Cell Drug II Cell nucleus Drug I II (c) III Binding site Conjugated gene Figure 6.4 Schematic of procedure for nanotip array drug/genes delivery (a) and the compositions of nanotips for the delivery of drugs (b) and genes (c). 219 (a) (b) Specie I Drug I Specie II Drug II Specie III Drug III (c) Recognition site Specie IV DNA Figure 6.5: Schematic of flow assisted organization of nanostructures: (a) multiple functional nanostructure, (b) drug and (c) gene loading in nanocapsules. 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However, scientists are able to observe the dynamic deformation of long DNA with electrophoresis when non-Newtonian fluid (e.g., linear polymer solutions). There are two types of motion for DNA molecules: one is “worm motion” and “hooked and sliding motion”. In “worm motion”, DNA migrates in polymer fluid like worms, the stretching out and drawing back motions are repeated in turn. In this type of motion, DNA keeps its so-called “I-shape” configuration. Scientists believe “worm motion” happens when DNA migrates in gaps between polymer molecular structures. In “hooked and sliding motion”, DNA molecules collide with a 229 polymer obstacle. The polymer structure provides the support spot for the sliding motion. Because of exerted electrophoretic force, DNA chain is extended into a “U-shape” configuration and then slides around the obstacle like a pulley. Because of the possible multiple entanglement points, other different shapes of configuration could also be found. The frequency of these two different configurations and motions depends on the size of DNS molecules and polymer and also the concentration and size of polymer solution. The “hooked and sliding motion” could be more often to be found when large DNA molecules, large and branched polymer or concentrated polymer solutions are used. In another words, entanglements could be more serious under these conditions. For the same reason, much diluted DNA concentration is employed in our single DNA dynamic deformation study. The entanglements among DNA molecules were minimized there. DNA conformation dynamics shown has the similar results obtained by others and described above. The purpose to carry out these experiments is to get practical experience for ourselves and make comparisons with Newtonian fluid for DNA deformation study described in Chapter 3 and Chapter 5. The reagents used here are T2 phage DNA and 1.0 wt% Poly (ethylene glycol, PEG, Mw = 4×106 Dalton) solution. The electric field strength is 60V/cm. The concentration of T2 DNA is 0.03 µg/ml, with the YOYO-1 dye added at a ratio of 1 dye per 5 base pair. Figure A.1 shows the snapshots of DNA experienced “worm motion” (a) and “hooked and sliding motion” (b). A.2 DNA DYNAMIC DEFORMATION IN FOUNTAIN FLOW As I mentioned at the beginning of this appendix, the deformation of DNA can be conducted under non-uniform flows. When liquid is driven by pressure into a dry 230 microchannel, the flowfront of the liquid is actually a fountain flow: the fluid penetrates to a finite height, stops, and falls back forming an outward intrusion at some intermediate height above the base. At the interface and in the backflow, great velocity difference exist, which, in principle, has the ability to deform DNA molecules. However, there are several technical challenges to observe deformation in fountain flow. First of all, depletion of DNA and liquid happens at the liquid-air interface. Few DNA molecules were observed at the interface than in the bulk when same concentration of DNA solution (e.g., 0.03 µg/ml) in previous deformation study is applied. But the concentration can not be too high to lead to obvious entanglements between different DNA molecules. Secondly, light scattering at the interface makes it difficult for imaging. Images taken by epi-fluorescence microscopy become fuzzy and no much detailed information of DNA dynamics can be obtained. Besides, at the liquid-air interface, the excited light may be refracted at the interface or into the liquid again, which would lead to higher total excitation light intensity than in the bulk liquid. As the result, high rate of photo-bleaching would happen at the interface. This local photo-bleaching make the interface imaging a real challenge. Thirdly, the imaging domain is limited to around 100 µm for clear DNA deformation image even when large DNA molecules (e.g., λ, T2, or T4 DNA) are used. The capture of the flowfront in the image domain is really painful. Deformation images shown in Figure A.2 were taken using a DNA solution with a concentration of ~0.3 µg/ml. A broad range of concentration for λ-DNA was tested and it has to balance between obtaining good image quality (not too bright to see anything) and capturing enough number of DNA molecules at the interface. It can be clearly seen a 231 DNA depletion zone close to the channel wall behind the flowfront. DNA molecules are found in major body of fountain flow. But because of the technique limitation of the second issue mentioned above, the images are pretty fuzzy. A better spin-disk laser confocal microscopy mentioned in the recommendation section might be helpful to improve the image quality since much thinner layer is illuminated and the imaging speed could also be highly improved with the spin disk device. To solve the third issue and make the flowfront more controllable, small channels (like slit) was used in experiments. The channel is only around 5 µm in depth, which provided high flow resistance so that the flow front could be much easier to be captured in that small domain. 232 (a) Continued Figure A.1 Snapshots of two typical motions for T2 DNA in 1 wt% PEG solution inside straight microchannel: (a) “worm-like” motion and (b) “hooked and sliding” motion. 233 Figure A.1 continued, (b) 234 Figure A.2 Images of flowfront in a fountain flow with λ-DNA solution (0.3 µg/ml). 235 APPENDIX B PRELIMINARY STUDY ON “ELECTROKINETIC GUN” As mentioned in the recommendation part, using non-uniform field could be attractive strategies to deliver materials (genes and drugs) into cells. So far, all of the experiments were performed without barriers on the small end. Preliminary study was discussed here when big insulated barriers are placed close to the small end. In the first example, air bubble is used as the insulate barrier. Air bubble is trapped on the small end. The liquid film covering the air bubble could conduct current so that a closed circuit is still formed in the whole channel. PS microspheres were placed on the large end side and continuous DC electric bias was added to drive microspheres towards the small end to shot the trapped air bubble. The size of air bubble increases, which is normal for most electrokinetic-driven. This is widely believed to be the result of local Joule heating happened when electric bias is added. When particles hit the air bubble, there are two possibilities for the motion of high momentum of particles: one is that particles break through both the liquid film and air bubble trapped in and the other is that particles only break in the liquid film and then 236 escape. Figure B.1 is a series of snapshots of two nanoparticles passing through an air bubble trapped at the small end of the converging channel. With these images, it is hard to tell whether these two high momentum nanoparticles enter the air bubble or not. But when particles tried to escape from the air bubble, trace of liquid was dragged from the liquid film and particles, as shown in Figures B.1 (c-e) and (g-j). This indicated that those nanoparticles at least were shot into the liquid film. Combining the electric force and liquid dragging, nanoparticles struggled with the thin liquid film for around 0.13 seconds and its moving track was twisted before they completed escape from the liquid film. In the second example, living cells (NIH 3T3) are used. Because the dimensions of the small end of the converging channel is close to the size of single cell, when cell suspension is loaded from the small end by weak vacuum, cells can be trapped at the small end. Figure B.2a present this trapping state. In that image, it probably traps more than a single cell but a cluster of cells at the small end. When electric pulse is added (100V/cm), materials from that cluster move away from the small end. This makes us believe the cells probably dead after the electric pulse. There are two possibilities counted for the death of cells: during the trapping and after the electroporation. In ongoing experiments, intermediate vacuum and electroporation parameters are used. New designs with multiple converging channels are used. The small end of channels is 5 µm in diameter. With this new design, single cells are easier to be trapped and studied. 237 (a) (b) (c) (d) (e) (f) Continued Figure B.1 The series of snapshots of charged nanoparticles’ motion in the converging channel with an air bubble trapped at the small end. (a-f) is for an individual nanoparticle and (g-l) is for the second one. The second particle enters the image domain in (d). The scale bar is 100 µm. 238 Figure B.1 continued, (g) (h) (i) (j) (k) (L) 239 (a) (b) Figure B.2 Cells trapping and electroporation study: (a) cells are trapped at the small end of converging channel and (b) after the electroporation. The scale bar is 20 µm. 240 APPENDIX C CD-LIKE MICROFLUIDIC PLATFORM FOR ENZYME-LINKED IMMUNOSORBENT ASSAY (ELISA) This appendix presents a compact disk (CD) microchip that performs enzyme- linked immunosorbent assay (ELISA) for detecting rat IgG from a hybridoma cell culture. Centrifugal and capillary forces were used to control the flow sequence of different reagents involved in the ELISA process. Each step of the ELISA process was carried out automatically by controlling the rotation speed of the CD. C.1 PROCEDURES A five-step flow sequencing CD (Figure C.1) was used. The first-antibody (affinity purified antibody goat anti-rat IgG (H+L)) and the BSA blocking solution were carried out off-chip by applying the first antibody (2.5 µg/ml) to the detection reservoir (reservoir 2). The antibody was allowed to adsorb onto the surface of this reservoir. After a 30-minute incubation, the excess antibody was removed by washing solution. The BSA solution was then added to block all excess protein-binding sites on the surface of the microchip. 241 After a 15-minute incubation and washing off the excess protein, the antigen/sample, washing, 2nd antibody, and substrate solutions were loaded into their corresponding reservoirs. The CD was mounted onto the motor plate. The rotation speed of the CD was set to 360 rpm (± 15 rpm) to release the sample solution (rat IgG) from reservoir 3 (containing to-be-assayed antigen) into reservoir 2 for the binding process of antigen-antibody. To ensure the immunoreaction to reach the equilibrium, a 15-minute incubation was chosen between two adjacent assay steps (According to the literature, several minutes of incubation are sufficient to reach equilibrium of the immunoreaction in a microchannel having a dimension similar to that of reservoir 2). After the incubation, reservoir 2 was washed with washing solution (from reservoir 4) at a rotation speed of 560 rpm (± 30 rpm). Based on previous experience, 3 times the volume of the washing solution is sufficient to displace the existing water-based solution in reservoir 2. The quantity of washing solution was therefore set at about 3 times the volume of reservoir 2 in the CD. The conjugate solution (affinity purified antibody horseradish peroxidase (HRP) labeled goat anti-rat IgG (H+L), in reservoir 5) was released into reservoir 2 at a rotation speed of 790 rpm (± 35 rpm) to let the enzyme-labeled secondary antibody bond to the primary antibody. After incubation, reservoir 2 was washed with washing solution (in reservoir 6) at a rotation speed of 1190 rpm (± 55 rpm). The substrate solution (3-(p- hydroxyphenyl)-propionic acid, HPPA, in reservoir 7) was released at a rotation speed of 1280 rpm (± 65 rpm) into reservoir 2. Immediately after the release of the substrate, the disk was stopped and detection was carried out using an inverted fluorescence microscope (Nikon ECLIPSE TE2000-U). 242 A 100W mercury light source with a 335/20 nm filter and a dichroic mirror was used as the excitation source. The fluorescence signal was obtained through a dichroic mirror and a 405/40 nm filter. Images were recorded with a 12-bit high-resolution monochrome digital camera system (CoolSnap HQ). The intensity of the fluorescence was analyzed using the Fryer Metamorph Image Analysis System. C.2 RESULTS AND DISCUSSIONS The fluorescence intensity level was checked along the whole detection microchannel (16 mm long). From the results showed in Figure C.2a, the signal is quite uniform from x/L=0.3 to x/L=0.8. The maximum variation is from both ends of the microchannel, about 30% higher than the average. The end effect is believed to be the result of copious reagents aggregates from the reservoirs, especially during the immobilization of the first antibody. In this case, such variation should be avoided so that the detection spot can be randomly picked up. However, this phenomenon could also be used in the future to improve the resolution of detection by concentrating reagents right surrounding the detection spots if their positions are fixed. During the detection, a special holder was used so that images were captured at the same region for each microchannel to avoid errors from signal variation. The signal (intensity of the fluorescence) variation in the same microchannel is within 5%. Figure C.2b shows the calibration curves of the microchip with fluorescence signal. The data were obtained from three independent measurements with different microchannels. Result in the microchip has a similar signal as that of the microtiter plate within the same detection range. The detection limit is 5 mg/L (31nM) of the rat IgG (MW ~ 160,000). 243 Since the concentration of the rat IgG from the hybridoma culture is typically in the range of 1 to 100 mg/L, this microfluidic platform is expected to be suitable for practical measurement. 244 CD center (a) (b) 5 4 3 2 1 7 6 (c) 1600 1400 1200 1000 800 600 rpm speed, Spin 400 200 0 0123456 Reservoir number Figure C.1 (a) Schematic of 5-step flow sequencing CD (reservoir 1 − antigen/sample, 2,4 − washing, 3 −2nd antibody, 5 − substrate, 6 − detection region, and 7 − waste), (b) a CNC-machined CD having four microfluidic platforms with the same design as shown in (a), and (c) the burst frequency of reagents from each reservoir. 245 (a) 1000 800 600 400 Fluorescence Intensity 200 0 0.0 0.1 0.2 0.3 0.4 0.5 0.6 0.7 0.8 0.9 1.0 x/L (b) 1000 900 800 700 600 500 400 300 Fluorescence Intensity 200 100 110100 IgG Concentration, µg/ml Figure C.2 (a) The signal variation checking along the whole microchannel and (b) the calibration curves of rat IgG conducted on the microchip. 246 APPENDIX D CRITICAL CONCENTRATION FOR DILUTE POLYMER SOLUTIONS All the flow-induced deformation theory (coil-stretch transition) for DNA dynamics addressed in Chapter 3 and Chapter 5 is valid only when DNA solution can be considered as the “dilute solution”. Depending on the concentration, polymer solutions can be divided as “dilute”, “semi dilute” and concentrated”. When the concentration of a polymer solution is far below the critical value, each polymer molecule can be considered in a coil state and have no interactions with other polymer molecules. On the other hand, if the concentration of a polymer solution is far above this value, many polymer molecules entangle heavily with each other and form a network with their polymer chains. Polymer solutions with the concentration around the critical value are classified as the “semi-dilute” polymer solution. In literature, there are several equations to calculate this critical concentration, or called overlap concentration in some cases. Equations (A-1~A-3) are from Doi and Edwards, des Cloizcaux and Jannink, and Graessley, respectively. Mw C * = (A-1) 4 3 3 π r g N A 247 Mw C * (A-2) = 3 ( 2 r g ) N A Mw C * = 3 (A-3) 8 r g N A where Mw is the molecular weight of polymer, rg is the radius of gyration of the polymer at the coil state, and NA is the Avogadro’s number, respectively. For λ-DNA, it is composed of totally 48, 502 base pairs (Mw=31.5×106 Dalton) and its radius of gyration at the super-coiled state is around 0.7 µm. The values of C* are about 0.036 mg/ml (Equation A-1), 0.054 mg/ml (Equation A-2) and 0.019 mg/ml (Equation A-3), respectively. 248