advancement of interpolyelectrolyte complexes for the

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Spring 2018

Advancement of Interpolyelectrolyte Complexes for the Delivery Advancement of Interpolyelectrolyte Complexes for the Delivery

of Genetic Material of Genetic Material

Keith Hampton Parsons University of Southern Mississippi

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Advancement of Interpolyelectrolyte Complexes for the Delivery of Genetic

Material

by

Keith Hampton Parsons

A Dissertation

Submitted to the Graduate School,

the College of Science and Technology

and the School of Polymer Science and Engineering

at The University of Southern Mississippi

in Partial Fulfillment of the Requirements

for the Degree of Doctor of Philosophy

Approved by:

Dr. Charles L. McCormick, Committee Chair

Dr. Alex S. Flynt

Dr. Sergei I. Nazarenko

Dr. Derek L. Patton

Dr. Daniel A. Savin

____________________ ____________________ ____________________

Dr. Charles L. McCormick

Committee Chair

Dr. Jeffrey S. Wiggins

Department Chair

Dr. Karen S. Coats

Dean of the Graduate School

May 2018

COPYRIGHT BY

Keith Hampton Parsons

2018

Published by the Graduate School

ii

ABSTRACT

This dissertation focuses on the development and advancement of

interpolyelectrolyte complexes (IPECs) and block ionomer complexes (BICs) for the

delivery of genetic material, namely RNA, to cells, both human and insect. RNA

interference (RNAi) provides a powerful tool for disease treatment and the elimination of

crop pests at the genetic level. Therefore, development of successful delivery vehicles for

its effector molecules, small interfering and double stranded RNAs (siRNA and dsRNA),

is imperative. IPECs and BICs show the most promise as RNAi vectors, and thus this

work focuses on ascertaining the structure-property relationships affecting RNA delivery

as well as applying such insights toward enabling RNAi in crop pest insects that remain

highly resistant to such treatment.

In Section I, BIC-siRNA interactions and effectiveness in cell transfection are

reported. Aqueous RAFT polymerization was used to prepare a series of hydrophilic-

block-cationic copolymers in which the cationic block statistically incorporates

increasing amounts of neutral, hydrophilic monomer such that the number of cationic

groups remains unchanged but the cationic charge density is diluted along the polymer

backbone. Reduced charge density decreases the electrostatic binding strength between

copolymers and siRNA with the goal of improving siRNA release after targeted cellular

delivery. However, lower binding strength resulted in decreased transfection and RNA

interference pathway activation, leading to reduced gene knockdown. Enzymatic siRNA

degradation studies with BICs indicated lowered binding strength increases susceptibility

to RNases, which is the likely cause for poor gene knockdown.

iii

Section II discusses how RNAi-based technologies are ideal for pest control as

they can provide species specificity and spare non-target organisms. However, in some

pests biological barriers prevent use of RNAi, and therefore broad application. In this

study we tested the ability of a synthetic cationic polymer, poly-[N-(3-

guanidinopropyl)methacrylamide] (pGPMA), that mimics arginine-rich cell penetrating

peptides to trigger RNAi in an insensitive animal–Spodoptera frugiperda. Polymer-

dsRNA interpolyelectrolyte complexes (IPECs) are efficiently taken up by cells, and can

drive highly efficient gene knockdown. These IPECs also trigger target gene knockdown

and moderate larval mortality when fed to fall armyworm larva. This effect was sequence

specific, which is consistent with the low toxicity we found to be associated with this

polymer. A method for oral delivery of dsRNA is critical to development of RNAi-based

insecticides. Thus, this technology has the potential to make RNAi-based pest control

useful for targeting numerous species and facilitate use of RNAi in pest management

practices.

iv

ACKNOWLEDGMENTS

I would like express my sincerest gratitude to my research advisor, Dr. Charles

McCormick not only for granting me the opportunity to pursue research in his

laboratories, but also for convincing me to come to USM and join the Polymer Science

department in the first place. In addition to his support and guidance throughout my

graduate career, I am particularly grateful to Dr. McCormick for fostering a research

environment that encourages independence and critical thinking while allowing for

freedom to pursue varying research avenues of my choosing. I believe that these qualities

have allowed me to develop into a better scientist than I thought possible, and for that, I

am truly grateful. I would also express my gratitude to Dr. Alex Flynt, who not only

opened his laboratories to me, but also mentored me in the development of my skills in

cell biology. His patience, encouragement, and expertise have allowed me to employ

skills and techniques within my research to which I would not have otherwise had access.

I am truly thankful for his guidance, which has been absolutely crucial to the success of

my graduate career. I would also like to thank the rest of my graduate committee, Dr.

Sergei Nazarenko, Dr. Derek Patton, and Dr. Daniel Savin, for their support and

guidance.

I am thankful for the support and friendship of all past and current McCormick

group members. I am especially grateful to Dr. Chris Holley and Dr. Brooks Abel for

their mentorship, without which I would have struggled significantly and would have

been unsuccessful. I would also like to specifically thank Dr. Wenming Wan, Dr. Marco

Oliveira, Phil Pickett, Gabbie Munn, and Kaden Stevens. I am also extremely grateful to

Dana Froelich, without whom none of this, even the day-to-day operations of the

v

McCormick group, would be possible. I greatly appreciate her friendship and emotional

support throughout my tenure here.

I thank Dr. Reid Bishop and Dr. Vijay Rangachari for allowing me the use of their

respective facilities as well as for their aid in solution DSC and CD spectroscopy

respectively. I thank Dr. William Jarrett for his support and patience in the use of NMR

and for his assistance in technical maintenance problems I have had with various

instruments.

Although there are too many to thank, I am deeply grateful for all of my friends

who have supported me throughout my graduate career. Thanks to them, my time in

Hattiesburg has been fun and memorable. I will always cherish our experiences together.

Additionally, I am grateful for the continuous love and support that I have received from

my family, who have always been my biggest advocates.

Finally, I would like to acknowledge the financial support that has made this

research possible: NSF EPSCoR EPS-0903727, NSF-MCB 1616725, and Mississippi

INBRE (NIH-NIGMS) P204M103476.

vi

DEDICATION

This work is dedicated to my family members who are no longer with us: my

grandparents, Ellis and Lee McCormick and Elise Parsons, and my great aunt, Helen

Grisham. Their love and support has made me the man I am today.

vii

TABLE OF CONTENTS

ABSTRACT ........................................................................................................................ ii

ACKNOWLEDGMENTS ................................................................................................. iv

DEDICATION ................................................................................................................... vi

LIST OF TABLES ........................................................................................................... xiii

LIST OF ILLUSTRATIONS ........................................................................................... xiv

LIST OF SCHEMES........................................................................................................ xxi

LIST OF ABBREVIATIONS ......................................................................................... xxii

CHAPTER I - INTRODUCTION ...................................................................................... 1

1.1 RNA Interference ...................................................................................................... 1

1.2 Barriers to RNA Delivery ......................................................................................... 3

1.3 IPECs: Physicochemical Considerations .................................................................. 7

1.4 Block Ionomer Complexes ..................................................................................... 11

1.5 IPECs and BICs for Nucleic Acid Delivery ........................................................... 13

1.6 RAFT Polymerization ............................................................................................. 25

CHAPTER II – OBJECTIVES OF RESEARCH ............................................................. 30

CHAPTER III - EXPERIMENTAL ................................................................................. 33

3.1 Block ionomer complexes consisting of siRNA and aRAFT-synthesized

hydrophilic-block-cationic copolymers II: The influence of cationic block charge

density on gene suppression.......................................................................................... 33

viii

3.1.1 Materials .......................................................................................................... 33

3.1.2 Polymer Synthesis ............................................................................................ 34

3.1.2.1 Synthesis of poly(HPMA-stat-APMA) macroCTA ................................. 34

3.1.2.2 Synthesis of poly[(HPMA-stat-APMA)-block-(HPMA-stat-

DMAPMA)] copolymers (P100, P75, P50, P25, and P0) ............................. 35

3.1.2.3 Copolymer functionalization with folic acid ............................................ 36

3.1.3 Formation of hydrophilic-block-cationic/oligonucleotide complexes .... 36

3.1.3.1 Preparation of copolymer-dsDNA complexes for solution

differential scanning calorimetry ...................................................................... 36

3.1.3.2 Preparation of copolymer-siRNA complexes for gene suppression 37

3.1.4 Cell Culture ...................................................................................................... 37

3.1.5 Gene Suppression of Human Survivin ............................................................. 37

3.1.6 Fluorescence Microscopy ................................................................................ 38

3.1.7 Cell Fractionation............................................................................................. 39

3.1.8 Copolymer Cytotoxicity................................................................................... 40

3.1.9 Characterization ............................................................................................... 40

3.2 Guanidinium-functionalized Interpolyelectrolyte Complexes Enabling RNAi in

Resistant Insect Pests .................................................................................................... 44

3.2.1 Materials .......................................................................................................... 44

3.2.2 Synthesis of pGPMA ....................................................................................... 45

ix

3.2.3 In vitro Transcription of dsRNA ...................................................................... 45

3.2.4 Polymer Characterization ................................................................................. 46

3.2.5 Light Scattering ................................................................................................ 47

3.2.6 Polymer-dsRNA Binding Assay ...................................................................... 48

3.2.7 Gene Suppression in Sf9 Cell Culture ............................................................. 48

3.2.8 Cell Viability Assay ......................................................................................... 49

3.2.9 Confocal Microscopy ....................................................................................... 50

3.2.10 Larval Feeding Experiments .......................................................................... 50

CHAPTER IV – RESULTS AND DISCUSSION............................................................ 52




4.1.1 Overview .......................................................................................................... 52

4.1.2 Synthesis of hydrophilic-block-cationic copolymers with varying cationic

block charge density ................................................................................................. 55

4.1.3 Hydrophilic-block-cationic copolymers with varying charge density form

stable, neutrally-charged complexes ......................................................................... 58

4.1.4 Reduced cationic block charge density decreases oligonucleotide stabilization

................................................................................................................................... 59

4.1.5 Reduced cationic block charge density reduces complex binding strength ..... 60

x

4.1.6 Reduced charge density diminishes gene knockdown efficacy ....................... 62

4.1.7 Cellular delivery and RNAi pathway activation decrease with reduced

cationic block charge density ................................................................................ 64

4.1.8 Enzymatic degradation rates increase as cationic block charge density

decreases ................................................................................................................... 67



4.2.1 Overview .......................................................................................................... 69

4.2.2 pGPMA synthesis and IPEC characterization ................................................. 71

4.2.3 pGPMA-dsRNA IPEC transfection and gene suppression in lepidopteran cell

culture ....................................................................................................................... 75

4.2.4 pGPMA-dsRNA IPEC gene suppression in lepidopteran larvae after oral

ingestion .................................................................................................................... 78

CHAPTER V – CONCLUSIONS .................................................................................... 81






APPENDIX A – SUPPLEMENTARY INFORMATION ................................................ 84

xi

A.1 Block ionomer complexes consisting of siRNA and aRAFT-synthesized



A.1.1 Discussion of experiments pertaining to control copolymer PO .................... 84

A.1.2 Supplementary Figures ................................................................................... 85

A.2 Guanidinium-functionalized Polymer Carriers Enable RNAi in Insensitive Crop

Pest Arthropods ............................................................................................................. 91

A.2.1 Primers Used in this Study .............................................................................. 91

A.2.2 Supplementary Figures ................................................................................... 92

APPENDIX B – INFLUENCE OF Z-GROUP HYDROPHOBICITY IN VISIBLE

LIGHT-MEDIATED AQUEOUS RAFT POLYMERIZATION ..................................... 96

B.1 Overview ................................................................................................................ 96

B.2 Experimental .......................................................................................................... 99

B.2.1 Materials .......................................................................................................... 99

B.2.2 Light Source .................................................................................................... 99

B.2.3 General Procedure for Blue Light-Mediated aRAFT Polymerizations ........ 100

B.2.4 Isolation of TTC Byproduct .......................................................................... 101

B.2.5 Characterization ............................................................................................ 101

B.3 Results and Discussion ......................................................................................... 102

B.3.1 Trithiocarbonate Byproduct .......................................................................... 102

xii

B.3.2 Effect of Byproduct Precipitation on Polymerization ................................... 105

B.4 Conclusions .......................................................................................................... 112

APPENDIX C RIGHTS & PERMISSIONS ................................................................... 114

REFERENCES ............................................................................................................... 116

xiii

LIST OF TABLES

Figure 1.5 pH-dependencies of the degree of conversion (ϴ) in the reaction between an

T20 oligonucleotide and poly(ethylene oxide)-block-polyspermine (black dots) and the

degree of ionization (α) of poly(ethylene oxide)-block-polyspermine (white dots). The

shift in pH between α-pH and ϴ-pH curves is shown by a horizontal arrow. .................. 10

Table 4.1 Molecular weight (number average), dispersity (Ð), composition, and dn/dc

values for macroCTA and chain-extended polymers ........................................................ 57

Table 4.2 The hydrodynamic radii (Rh), ζ-potential, and percent complexed siRNA for

siRNA and copolymer-siRNA complexes ........................................................................ 59

Table 4.3 ζ-potential measurements of pGPMA-dsRNA complexes at varying weight

ratios and pH ..................................................................................................................... 74

Table 4.4 Dynamic and static light scattering measurements of pGPMA-dsRNA

complexes at varying pH (weight ratio = 8) ..................................................................... 74

xiv

LIST OF ILLUSTRATIONS

Figure 1.1 . Mechanism of RNA interference3 ................................................................... 2

Figure 1.2 Endocytosis and endosomal escape of siRNA. ................................................. 5

Figure 1.3 Schematic representation of IPEC formation .................................................... 8

Figure 1.4 Schematic representation of stoichiometric and nonstoichiometric IPECs ....... 8




shift in pH between α-pH and ϴ-pH curves is shown by a horizontal arrow.13 ............... 10

Figure 1.6 BIC formed from polyelectrolyte homo- and block copolymers. Red and green

indicate oppositely charged polyelectrolyte segments. Blue indicates neutrally charged

hydrophilic segments. ....................................................................................................... 12

Figure 1.7 Schematic representation of the BIC formed between poly(ethylene oxide)-

block-poly(sodium methacrylate) and poly[4-vinylpyridine-stat-N-ethyl-4-

vinylpyridinium bromide]. (A) The contour length of the polycation chain is shorter or

equals that of the sodium methacrylate segment; (B) the polycation chain is much longer

than the sodium methacrylate segment.13 ......................................................................... 12

Figure 1.8 Schematic model for charged block length-dependent recognition to form

BICs and BIC self-assembly.37 ......................................................................................... 13

Figure 1.9 Enzymatic degradation of small interfering ribonucleic acid with RNase A in

the presence and absence of (HPMA258-b-DMAPMA13)46 ............................................... 15

Figure 1.10 BIC formation from siRNA and (HPMA-s-APMA)-b-DMAPMA copolymers

functionalized with folic acid.18 ........................................................................................ 16

xv

Figure 1.11 Fluorescent microscope images of small interfering RNA (siRNA; cyanine-3

and fluorescein (FAM) labeled) delivery to KB cells (A, C, E) and A549 cells (B, D, F).

Lipofectamine (A, B; + control), unconjugated (N-(2-hydroxypropyl)methacrylamide315-

stat-N-(3-aminopropyl)methacrylamide13)-b-N-(3-

dimethylaminopropyl)methacrylamide23, (HPMA315-stat-APMA13)-b-DMAPMA23 (C, D;

− control), and folic acid conjugated (HPMA315-stat-APMA13)-b-DMAPMA23 (E, F).

Nuclei were stained with 4′,6-diamidino-2-phenylindole (blue). For clarity, FAM

fluorescence is not shown. Scale bars = 50 μm.18 ............................................................. 17

Figure 1.12 (A) Structure of pAcF. (B) Schematic illustration of pAcF- mutated anti-

HER2 antibodies, S202-pAcF Fab, Q389-pAcF IgG, and A121-pAcF IgG. (C) Synthetic

scheme for the antibody−polymer conjugates.62 ............................................................... 18

Figure 1.13 Confocal microscopy of internalization of siRNA mediated by S202-Fab-P1.

HeLa (A,C,E) and SKBR-3 (B,D,F) cells were treated with buffer (A,B), 200 nM S202-

pAcF Fab + 200 nM P1 +50 nM siRNA-FITC (C,D), or 200 nM S202-Fab-P1 + 50 nM

siRNA- FITC (E,F) for 4 h. Cells were then stained with Hoechst 33342 (blue) and ER-

Tracker (red) and imaged with a Leica 710 confocal microscope. Bar = 10 μm.62 .......... 18

Figure 1.14 Cellular uptake of FITC-labeled HPMA-b-GPMA copolymers.64 ................ 19

Figure 1.15 Scheme of polyelectrolyte-membrane protein interaction.12 ......................... 20

Figure 1.16 HPMA-b-L-Glu copolymers form α-helices at low pH, allowing them to

embed themselves in cell membranes.71 ........................................................................... 21

Figure 1.17 Mean residue ellipticity as a function of pH for poly[HPMA220-b-(L-

Glu56)].71............................................................................................................................ 21

xvi

Figure 1.18 RAFT-mediated synthesis of diblock copolymers consisting of a cationic

poly(dimethylaminoethyl methacrylate) (DMAEMA, x=58) block and an endosomolytic

polyampholyte block incorporating DMAEMA and propylacrylic acid (PAA) in

equimolar ratios, and butyl methacrylate (BMA) (y~70)72 .............................................. 22

Figure 1.19 (a) Formation of SS-PAAs/DNA polyplexes that are stable in the

extracellular environment. (b) Intracellular reduction of the disulfide linkages in the

polymer of the polyplex. (c) Dissociation of DNA from the degraded polymer.89 .......... 24

Figure 1.20 IPECs formed from oligo-DNA and PDMAEA dissociate because of self-

catalyzed hydrolysis in water, whereas those formed from quaternized PDMAEA do

not.90 .................................................................................................................................. 25

Figure 4.1 ASEC-MALLS of poly(HPMA-stat-APMA) macroCTA and subsequent chain

extensions with DMAPMA and HPMA (P100-P0) ......................................................... 57

Figure 4.2 Differential power thermograms for copolymer-dsDNA complexes. Samples

shifted along Y-axis for clarity. ........................................................................................ 60

Figure 4.3 3. α- and θ vs. pH curves for (A) P100, (B) P75, (C) P50, (D) P25, (E) P0, and

(F) P2 and their respective complexes with PSS .............................................................. 62

Figure 4.4 RT-qPCR analysis of down-regulation of human survivin mRNA by

copolymer-siRNA complexes. mRNA expression normalized to Lipofectamine. ........... 63

Figure 4.5 Corrected total fluorescence of siRNA labelled with AlexaFluor594 delivered

via copolymer complexes. Samples not belonging to same letter grouping were found to

have statistically significant variance via Tukey analysis. ............................................... 65

xvii

Figure 4.6 Relative radio-labelled siRNA content after copolymer complex delivery and

cell fractionation. Higher gradient numbers correspond to heavier fractions. siRNA

content normalized to fraction 1 for each complex. ......................................................... 66

Figure 4.7 (A) Molar ellipticity of siRNA before and 20 minutes after addition of

Riboshredder RNase blend. (B) Enzymatic degradation of free and copolymer-complexed

siRNA with Riboshredder RNase blend as monitored by the normalized disappearance of

the CD band at 212 nm. .................................................................................................... 68

Figure 4.8 (a) Structure, number average molecular weight (Mn), and dispersity (Ð) of

pGPMA. (b) SEC trace of pGPMA (c) Gel electrophoresis of pGPMA-dsRNA IPECs.

Numbers indicate polymer/dsRNA weight ratio. (d) Proposed morphological changes in

IPEC structure between pH = 7.4 and pH = 10. ............................................................... 73

Figure 4.9 Sf9 cells treated with Cy5-labeled dsRNA (red) complexed with pGPMA after

(a, top row) 24 h or (b, bottom row) 48 h. Nuclei were stained with DAPI (blue). Scale

bars = 5 μm. (c) Cell viability assay of pGPMA after 48 h employing polymer

concentrations identical to the indicated weight ratios used in IPECs. Cell viability was

determined relative to the untreated control. Error bars represent the standard deviation

from triplicate experiments. .............................................................................................. 76

Figure 4.10 (a) Expression of CDC27 determined by RT-qPCR in Sf9 cells following

incubation with pGPMA complexed with either CDC27- or control-dsRNA. Numbers

indicate polymer/dsRNA weight ratios. Values are normalized to CDC27 expression in

respective control (KIF-dsRNA-treated) samples. Errors bars represent SEM. (b)

Expression of CDC27 determined by RT-qPCR in Sf9 cells following incubation with

CDC27 dsRNA complexed with either pGPMA (8x) or Lipofectamine 3000. Values are

xviii

normalized relative to respective untreated controls. Error bars represent SEM. (c) RT-

qPCR quantification of CDC27-dsRNA transfected by pGPMA, Lipofectamine 3000, or

untreated control. Values are relative to zero. Error bars represent SEM. For plots (a-c),

groupings indicated with asterisks (*) were found to be significantly different after Tukey

analysis. ............................................................................................................................. 78

Figure 4.11 (a) Expression of V-ATPase mRNA in midgut tissue from 2nd instar fall

armyworm larvae fed with pGPMA complexed with either V-ATPase dsRNA or GFP

dsRNA determined by RT-qPCR. Letters indicate individual animals. Days between

feeding and harvesting are indicated in parentheses. Values are normalized to V-ATPase

expression in control sample. Errors bars represent SEM. (b) Percent survival of 2nd and

3rd fall armyworm larvae fed pGPMA complexed with dsRNA targeting V-ATPase (N =

25) or control dsRNA (N = 31). (c) Image of fall armyworm larval gut after feeding with

pGPMA complexed with dsRNA targeting GFP or (d) sfV-ATPase. Scale bars = 2 mm.

........................................................................................................................................... 80

Figure A.1 1H NMR spectra of P100, P75, P50, P25, P0, and macroCTA .................... 85

Figure A.2 UV-Vis spectroscopy of conjugated folic acid P100, P75, P50, P25, and P0

copolymers. ....................................................................................................................... 86

Figure A.3 Agilent 2100 Bioanalyzer electropherograms for copolymer-siRNA

complexes, free siRNA, and ladder. Uncomplexed siRNA concentration determined from

area of peak at ~39 s. Peak at ~35 s corresponds to 4-nucleotide marker. ....................... 87

Figure A.4 . Potentiometric titration curves for (A) P100, (B) P75, (C) P50, (D) P25, (E)

P0, and (F) P2 and their respective block ionomer complexes with polystyrene sulfonate

(PSS). ................................................................................................................................ 88

xix

Figure A.5 Confocal fluorescence microscopy images of KB cells treated with

AlexaFluor594-tagged siRNA (red) delivered via (A) P100, (B) P75, (C) P50, (D) P25,

and (E) P0. Nuclei were stained with DAPI (blue). Scale bars = 10 μm. ......................... 89

Figure A.6 Cell viability assays of P100, P75, P50, P25, and P0 after 48 and 72 hours.

The cell viability was determined relative to KB cells. Error bars represent the standard

deviation from triplicate experiments. .............................................................................. 90

Figure A.7 1H NMR spectra of pGPMA polymerization aliquots taken at (a) t0 and (b) 19

hr. ...................................................................................................................................... 92

Figure A.8 Hydrodynamic size histograms for pGPMA-dsRNA IPEC (8x weight ratio) at

pH = 7.4 (black) and pH = 10 (red) and free dsRNA (pH = 7.4, green) as determined by

dynamic light scattering (DLS) at 90° scattering angle. ................................................... 93

Figure A.9 Sf9 cells treated with free Cy5-labeled dsRNA (red) after (a, top row) 24 hrs

and (b, bottom row) 48 hrs. Nuclei were stained with DAPI (blue). Scale bars = 5 μm. . 94

Figure A.10 Expression of CDC27 determined by RT-qPCR in Sf9 cells following

incubation with free pGPMA at identical concentration as used for 8x IPEC. Values are

normalized to respective untreated controls. Error bars represent SEM. ......................... 94

Figure A.11 Survival of fall armyworm larvae after ingestion of pGPMA. Animals were

directly fed masses indicated on the left y-axis (black line). The percentage of animals

viable after feeding on right y-axis (grey line). (N = 4) ................................................... 95

Figure B.1 Home-built photo-reactor used for visible light-mediated aRAFT

polymerizations ............................................................................................................... 100

Figure B.2 Structures of monomers and RAFT agents used in this study. ..................... 102

xx

Figure B.3 1H NMR spectra of ethyltrithiocarbonic acid, bisethyltrithiocarbonate, and

TTC byproduct. ............................................................................................................... 104

Figure B.4 FT-IR spectra of ethyltrithiocarbonic acid, bisethyltrithiocarbonate, and TTC

byproduct. ....................................................................................................................... 105

Figure B.5 Mn vs. conversion plots, pseudo-first order kinetic plots, and photos of final

polymerization solutions for visible light-mediated aRAFT polymerization of MAPTAC

using (a) ImET and (b) CETPA as CTA. [M]0:[CTA] =75 for both reactions. ............. 106

Figure B.6 ASEC traces for MAPTAC macroCTAs and their chain extensions to form

MAPTAC-b-MAPTAC using (a) ImET and (b) CETPA as CTA. ................................. 107

Figure B.7 Percent transmittance and monomer conversion as a function of time for

reaction of ImET with MAPTAC and ImET alone in the presence of blue light. .......... 108

Figure B.8 Mn vs. conversion plots and pseudo-first order kinetic plots for visible light-

mediated aRAFT polymerization of HPMA using ImET as CTA. [M]0:[CTA] = (a) 200,

(b) 400, (c) 500, (d) 1000. ............................................................................................... 110

Figure B.9 ASEC-MALLS traces of visible light-mediated aRAFT polymerization of

HPMA using ImET as the RAFT agent targeting [M]0:[CTA] = 1000. ......................... 112

xxi

LIST OF SCHEMES

Scheme 1.1 Mechanism of RAFT polymerization ........................................................... 26

Scheme 1.2 Mechanism of light-mediated RAFT polymerization ................................... 28

Scheme 4.1 Synthetic pathway for the preparation of poly[(HPMA-stat-APMA)-block-

(HPMA-stat-DMAPMA)] copolymers and subsequent complexation with siRNA. ....... 55

Scheme B.1 Mechanism of light-mediated RAFT polymerization .................................. 97

Scheme B.1 Proposed mechanism for byproduct precipitation during visible light-

mediated aRAFT polymerization of MAPTAC using ImET as CTA. ........................... 103

xxii

LIST OF ABBREVIATIONS

AIBN Azobisisobutyronitrile

APC Antibody-Polymer Conjugate

APMA N-(3-aminopropyl)methacrylamide

aRAFT Aqueous Reversible Addition-Fragmentation Chain Transfer

ASEC Aqueous Size Exclusion Chromatography

ATP Adenosine Triphosphate

ATRP Atom-Transfer Radical Polymerization

BIC Block Ionomer Complex

BMA Butyl Methacrylate

CD Circular Dichroism

CEP 4-Cyano-4-[(ethylsulfanylthiocarbonyl)sulfanyl]pentanoic acid

CPP Cell-Penetrating Peptide

CTA Chain Transfer Agent

CTF Corrected Total Fluorescence

DAPI 4’,6-diamidino-2-phenylindole

DCC Dicyclohexylcarbodiimide

diNHS-FA di(N-Hydroxysuccinimide)-activated Folic Acid

DLS Dynamic Light Scattering

DMAEA N-(2-dimethylaminoethyl)acrylate

DMAEMA N-(2-dimethylaminoethyl)methacrylate

DMAP 4-Dimethylaminopyridine

DMAPMA N-(3-Dimethylaminopropyl)methacrylamide

DMF Dimethylformamide

DMSO Dimethylsulfoxide

dn/dc Change in refractive index with respect to change in concentration

DNA Deoxyribonucleic Acid

DSC Differential Scanning Calorimetry

dsDNA Double Stranded DNA

dsRNA Double Stranded RNA

EDTA Ethylenediaminetetraacetic Acid

EPR Enhanced Permeation and Retention

FBS Fetal Bovine Serum

GPMA N-(3-guanidinopropyl)methacrylamide

HEPES 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid

HPLC High Performance Liquid Chromatography

HPMA N-(2-hydroxypropyl)methacrylamide

IPEC Interpolyelectrolyte Complex

MALLS Multi-Angle Laser Light Scattering

MeOH Methanol

miRNA Micro RNA

mRNA Messenger RNA

MTT 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide

N:P Nitrogen-to-Phosphate Ratio

xxiii

NHS N-Hydroxysuccinimide

NMP Nitroxide-Mediated Polymerization

NMR Nuclear Magnetic Resonance

PAA Poly(Acrylic Acid)

PAAm Poly(Amido Amine)

PBS Phosphate Buffered Saline

PCR Polymerase Chain Reaction

pDNA Plasmid DNA

PEG Polyethylene Glycol

PSS Poly(Sodium Styrene Sulfonate)

PVDF Poly(Vinylidene Fluoride)

RAFT Reversible Addition-Fragmentation Chain Transfer

RBC Red Blood Cell

RDRP Reversible-Deactivation Radical Polymerization

RI Refractive Index

RISC RNA-Induced Silencing Complex

RNA Ribonucleic Acid

RNAi RNA Interference

RT-qPCR Reverse Transcription Quantitative Polymerase Chain Reaction

SEC Size Exclusion Chromatography

SFM Serum-Free Medium

siRNA Small Interfering RNA

SLS Static Light Scattering

TAE Tris, Acetic Acid, EDTA

Tris Tris(Hydroxymethyl)aminomethane

TTC Trithiocarbonate

UV Ultraviolet

UV-Vis Ultraviolet-Visible Light Spectroscopy

V-501 4,4’-Azobiscyanovaleric Acid

V-ATPase Vacuolar ATPase

1

CHAPTER I - INTRODUCTION

1.1 RNA Interference

RNA interference (RNAi) refers to post-transcriptional gene regulation in which

small RNA duplexes, like microRNA (miRNA) and small interfering RNA (siRNA),

along with their associated enzymes, destroy or inhibit the translation of genetic

transcripts like messenger RNA (mRNA), resulting in gene suppression, or

“knockdown.”1 Discovered by Fire and Mello in 1998,2 RNAi has prompted extensive

investigation over the last two decades not only because it offers insight into cellular

regulatory mechanisms, but also for the promising genetic manipulation applications it

provides us as researchers. The phenomenon has been demonstrated to occur in

vertebrates, invertebrates, and plants,3 and while the specific proteomic machinery can

vary across different species, the fundamental mechanism is consistent.

The mechanistic details of RNAi have been reviewed elsewhere,3 but the

simplified process (Figure 1.1) begins with the processing of double stranded RNA

(dsRNA) by Dicer to form 21-25 nucleotide (nt) siRNA with 2-nt 3’ overhangs and 5’

phosphates. The dsRNA can be generated in the nucleus as miRNA or can be introduced

to the cytoplasm exogenously as siRNA, produced synthetically or as a result of viral

infection. The siRNA duplex consists of a guide strand that is complementary to the

targeted mRNA as well as a passenger strand that is removed once the siRNA is loaded

into the RNA-induced silencing complex (RISC). Once loaded, the RISC performs

“cellular surveillance,” binding with targeted mRNA through sequence-specific

hybridization with the siRNA guide strand. The mRNA is then either sliced, or other

enzymes are recruited to the RISC to induce translational suppression without slicing,

2

which can then lead to deadenylation and degradation of the targeted mRNA. Active

RISCs can then be recycled to continue cellular surveillance.

Figure 1.1 . Mechanism of RNA interference3

Control of RNAi through the introduction of synthetic siRNA or dsRNA to cells

offers great promise for a variety of genetic applications. Clinically, RNAi could be used

to suppress genes critical to cancer proliferation or as treatment for other genetic diseases.

More recently, the high RNAi activity in insects, combined with increasing pest

3

resistance to traditional pesticides, has prompted its use in pest control.4,5 However, there

exist a number of barriers to successful RNA delivery.

1.2 Barriers to RNA Delivery

While the specifics vary drastically between humans and insects, there are four

broad categories of barriers to successful RNAi that must be considered in the context of

this work: RNA protection, cell transfection, endosomal escape, and payload release from

the delivery vehicle. For the remainder of this discussion, we will assume that RNA

delivery is intravenous in humans and orally ingested in insects. Introduction of the

delivery vehicle directly to the bloodstream, through which the vector usually must arrive

regardless of origin, in humans bypasses a number of other obstacles, whereas an

effective crop pesticide requires oral ingestion of the vector within or on the crop foliage.

Additionally, while RNAi has been investigated in a wide variety of insect species, the

barriers discussed herein are most applicable to lepidopterans (i.e. moths and butterflies)

as they are typically refractory towards RNAi6 and are thus the subject of the research

described in Chapter 4.2.

Regardless of the target species or the specific RNA to be delivered (i.e. siRNA or

dsRNA), encapsulation and protection of the cargo is crucial. RNA is inherently

hydrolytically unstable as a result of the 2’ hydroxyl group that can readily attack the

neighboring phosphodiester groups, resulting in greater hydrolytic instability relative to

DNA.7 RNA hydrolysis is further compounded under alkaline conditions, and while

human plasma tends to remain relatively neutral, lepidopteran gut tracts can approach pH

values > 10-11.8 Furthermore, because much viral activity is heavily RNA-dependent,

most living organisms produce and excrete high levels of non-specific RNases that attack

4

and degrade the delivered RNA.9 Indeed, unmodified siRNAs in human plasma

experience up to 75% degradation in under 5 minutes.10 Similarly, lepidopteran

hemolymphs have been shown to contain extraordinary concentrations of RNases that

can result in dsRNA half-life of 27 min (ex vivo).11 Thus, a vector that is able to

hydrolytically stabilize RNA while protecting it from enzymatic degradation is required.

In human blood circulation, this issue is further compounded by the immune system,

which can potentially attack the vector itself,12 requiring delivery vehicles to have

“stealth” characteristics.13 Furthermore, the vector should be large enough in size to

prevent renal clearance and ensure extended circulation times required to reach the

targeted cells.9 Similarly to circulation requirements in humans, it has been hypothesized

that effective RNAi in insects requires extended gut retention times;11 however, this

theory has not been extensively investigated.

Once the RNA payload has been sufficiently protected and delivered to the

desired location, it must be internalized by the targeted cells to activate the RNAi

machinery. Cellular uptake of free RNA is typically hindered by the inability of the

anionic nucleic acids to transverse the hydrophobic interior of the cellular membrane.9

Most nucleic acid delivery strategies instead rely upon a vector that can induce

transfection, either by facilitating membrane transport, as in the case of cationic

surfactants and viral vectors,14 or by inducing endocytosis, in which the cell membrane

invaginates and separates from the membrane with the payload enclosed within (Figure

1.2).15 Most often this is achieved by targeting a membrane protein responsible for

triggering endocytosis16 – termed receptor-mediated endocytosis. Conveniently, this same

mechanism can also be used to impart cell-specificity to the vector, such as in the case of

5

folic acid to target tumor cells.17,18 In insects, however, most RNA delivery strategies rely

upon direct injection into the cells or passive uptake.5,19 Many insect cells will

autonomously internalize dsRNA to elicit RNAi by several proposed mechanisms.19 In

fact, most in vitro studies rely upon simply incubating the cells with dsRNA added to the

medium.5,19 However, lepidopterans, while able to internalize free dsRNA via

endocytosis, do not exhibit an RNAi response as a result of particularly efficient

endosomal entrapment,20 a problem common to human treatment as well.

Figure 1.2 Endocytosis and endosomal escape of siRNA.

Internalization via endocytosis results in the formation of a lipid vesicle known as

an endosome, in which the cargo is trapped (Figure 1.2). Endosomes rapidly either re-

fuse with the cell membrane, expelling their contents, or are trafficked to fuse with

lysosomes within the cell, resulting in the enzymatic destruction of their contents.15 Thus,

in order to initiate an RNAi response, the RNAi cargo must be able to effectively escape

endosomal entrapment and be released into the cytoplasm prior to its expulsion or

destruction. While there seem to be some mechanisms for naturally occurring endosomal

6

release, as evidenced by RNAi from naked RNAs in some cases, most delivery vectors

require an active approach to endosomal membrane disruption. In humans, endosomal

development is accompanied by a drop in pH from physiological 7.4 to 5-7, and thus

many delivery vehicles exploit this phenomenon to induce a pH-responsive behavior that

can disrupt the endosomal membrane.15 Endosomal entrapment is particularly effective in

lepidopterans, in which it is thought to be the predominant factor in their RNAi

insensitivity.20

Finally, once endosomal escape has been achieved, the RNA payload must be

released from its vehicle into the cytoplasm where it can be incorporated into and

processed by Dicer. The mechanism of RNA release is heavily dependent upon the vector

employed, but release rate has been hypothesized to have a drastic effect on gene

knockdown: slow release results in inefficient activation of the RNAi machinery while

too rapid release can increase susceptibility to RNase-mediated degradation within the

cytoplasm.21 Thus, an effective RNA delivery vehicle must protect its cargo while

ensuring delivery to the cytoplasm at an efficient rate.

In response to the above barriers to RNAi, a tremendous amount of research has

focused on the design and implementation of nucleic acid delivery vehicles, the most

popular of which fall into three categories: viral, lipid-based, and polymeric.

Because the primary function of viral activity is the injection of its own genetic

material into a host cell for reproduction, viral vectors are inherently very efficient RNA

delivery vehicles. As a result, many of the earliest efforts focused on their use for

delivery of exogenous RNA.22,23 However, while highly efficient in trafficking nucleic

acids to the cytoplasm, viral vectors suffer from lack of cell specificity and can elicit a

7

strong immune response.24 Lipid based vectors have also seen great success, particularly

as commercial transfections agents.25 They too, however, lack cell specificity, and their

toxicity poses significant concern.15 Conversely, synthetic polymeric vectors have

attracted the most attention as a result of the broad range of chemistries available to their

syntheses. While non-ionic polymer-based delivery vehicles like polymerosomes26 and

polymer-RNA conjugates have seen success,27 the vast majority of research has focused

on the development of polycationic systems that can electrostatically complex with the

negatively charged RNA backbone to form interpolyelectrolyte complexes (IPECs).9,21

1.3 IPECs: Physicochemical Considerations

The detailed kinetic and thermodynamic considerations for IPEC formation have

been extensively reviewed by Kabanov.28–33 The following is a condensed summary.

Polyelectrolytes result from the polymerization of charged or ionizable monomers

and, analogously to acids and bases, are categorized as either strong or weak. Strong

polyelectrolytes, such as those consisting of quaternary amines, phosphates, and

sulfonates, remain fully charged across non-extreme solution pH ranges. Conversely,

weak polyelectrolytes exhibit pH-dependent ionization across moderate pH ranges and

typically consist of non-quaternary amines and carboxylic acids. IPECs can be formed

most simply via the mixing of two aqueous solutions containing oppositely charge

polyelectrolytes (Figure 1.3), resulting in the following equilibrium:

(pA−b+)n + (pB+a−)m ⇌ [(pA−pB+)x ∙ (pA−b+)n−x ∙ (pB+a−)m−x] + xa− + xb+ (1)

where pA− and pB+ indicate repeat units of anionic and cationic polyelectrolytes

respectively, and a− and b+ indicate the respective small molecule counterions. From

8

equation 1, it becomes apparent that, in addition to Coulombic interaction, IPEC

formation is heavily driven by the entropically favorable release of a large number of

small molecule counterions. Indeed, small molecule salt concentrations have a significant

impact on IPEC dissociation and exchange (vide infra).

Figure 1.3 Schematic representation of IPEC formation

IPECs consisting of linear homopolymers fall into two categories as depicted in

Figure 1.4: stoichiometric and nonstoichiometric as defined by Z = m/n (i.e. the ratio of

cationic functionalities to anionic functionalities). For stoichiometric IPECs where Z = 1,

all complex charges are neutralized, resulting in insolubility and precipitation of the

resulting IPEC. For nonstoichiometric IPECs, if Z > 1 the resulting complex will be

cationic overall and anionic overall if Z < 1. In such systems, the polyelectrolyte in

excess is referred to as the host molecule, with the other referred to as the guest.

Figure 1.4 Schematic representation of stoichiometric and nonstoichiometric IPECs

9

IPECs are further characterized by the extent of the complex-forming reaction, ϴ,

defined as the ratio of salt bonds formed to total possible salt bonds. In terms of equation

1, ϴ = x/m when n ≥ m (i.e. the polyanion is the host), or ϴ = x/n when m > n (i.e. the

polycation is the host). For IPECs consisting of two strong polyelectrolytes, ϴ typically

can be considered to = 1. Exceptions generally are limited to circumstances where salt

bond formations are inhibited, such as at high ionic strength or IPEC precipitation prior

to full complexation when Z = 1.

On the other hand, for IPECs consisting of at least one weak polyelectrolyte, ϴ

has a pH-dependency and thus can be controlled by adjusting the solution pH. Both ϴ

and the degree of ionization of the weak polyelectrolyte in the absence of another, α, can

be calculated from their respective potentiometric titration curves. When α and ϴ are

plotted as a function of pH, there is a shift (ΔpH(α,ϴ)) between the α vs. pH curve and

the ϴ vs. pH curve due to the cooperativity of complex formation (Figure 1.5). If the

weak polyelectrolyte being titrated is the guest molecule, all of its charged units at a

given pH can be assumed to form a salt bond; i.e. α = ϴ for any given pH. This is related

to the free energy of complex formation, ΔGtotal, as a function of α, where α = α1 (= ϴ1):

∆𝐺𝑡𝑜𝑡𝑎𝑙 = −2.303𝑅𝑇 ∫ ∆pH(𝛼)𝑑𝛼

𝛼1

0

(2)

Thus, the greater ΔpH(α,ϴ), the more stable the complex formed.

10




shift in pH between α-pH and ϴ-pH curves is shown by a horizontal arrow.13

IPEC formation is remarkably fast: with a rate constant on the order of 109 M-1s-1,

complexation occurs within < 5 ms. Although IPECs technically exist in equilibrium

according to equation 1, the high entropic penalty for complex dissociation renders the

reverse rate virtually non-existent. As a result, complexes formed at low ionic strength

are most often kinetically trapped in a non-equilibrium state with random ionic pairings.

However, at moderate ionic strength (typically < 400 mM for NaCl), where the entropic

penalty is mitigated, there is a second slow step that sees the rearrangement of salt bonds

to form the equilibrium product. Further increasing ionic strength eventually results in

complex dissociation. Alternatively, when a weak polyelectrolyte is involved, the

equilibrium product can formed by slowly ionizing the weak polyelectrolyte via gradual

titration with the appropriate strong acid or base.

11

Ionic strength also plays an important role in IPEC exchange and substitution

reactions. Consider the following transfer of guest polyanion 𝑝𝐴− from host polycation

𝑝𝐵+ to 𝑝𝐶+:

[𝑝𝐴− ∙ 𝑝𝐵+] + 𝑝𝐶+ ⇌ [𝑝𝐴− ∙ 𝑝𝐶+] + 𝑝𝐵+

(3)

If 𝑝𝐵+ and 𝑝𝐶+ are identical, the reaction is termed exchange; if they are

different, it is termed substitution. Rather than occurring via a dissociation mechanism,

exchange occurs in a first fast step as 𝑝𝐶+diffuses to the IPEC to form triple complex

[𝑝𝐵+ ∙ 𝑝𝐴− ∙ 𝑝𝐶+]. As with IPEC formation, at low ionic strength the triple complex is

kinetically trapped in a random, non-equilibrium state. Moderate ionic strength allows for

a second slow step involving the redistribution of chains to the equilibrium product. The

rate of transfer is inversely proportional to the guest polyelectrolyte length.

1.4 Block Ionomer Complexes

The above discussion focuses on IPECs formed from linear polyelectrolyte

homopolymers, which are water insoluble at Z = 1 because all hydrophilic charges are

neutralized. However, if one or both of the polyelectrolytes in question is a block

copolymer with a neutrally charged, hydrophilic block, water soluble IPECs can be

formed at Z = 1.34 The result is a micelle-like structure known as a block ionomer

complex (BIC, Figure 1.6), whose morphology depends on the lengths and structures of

the hydrophilic and ionic blocks (Figure 1.7).34–36 Because the hydrophilic block(s) forms

a solubilizing corona around the hydrophobic IPEC, BICs can be formed with neutral

overall charges. Furthermore, the BIC corona stabilizes it across a wider pH range

compared to the IPEC from corresponding homopolymers.13 Like traditional IPECs, BICs

are sensitive to ionic strength and can undergo exchange/substitution reactions.13 With

12

the exception of solubility at Z = 1, the thermodynamic and kinetic considerations for

BICs are essentially the same as those for IPECs.

Figure 1.6 BIC formed from polyelectrolyte homo- and block copolymers. Red and green

indicate oppositely charged polyelectrolyte segments. Blue indicates neutrally charged

hydrophilic segments.

Figure 1.7 Schematic representation of the BIC formed between poly(ethylene oxide)-

block-poly(sodium methacrylate) and poly[4-vinylpyridine-stat-N-ethyl-4-

vinylpyridinium bromide]. (A) The contour length of the polycation chain is shorter or

equals that of the sodium methacrylate segment; (B) the polycation chain is much longer

than the sodium methacrylate segment.13

One important distinction, however, is that mixing two oppositely charged block

copolymers with charged blocks of equal length results in 1:1 BIC formation to form

what in appearance is an ABA triblock copolymer with a hydrophobic B block that can

self-assemble into higher order structures (Figure 1.8).37 Based on the charged block

lengths, the self-assemblies can range from micelles37 to vesicles.38 Furthermore,

changing one of the hydrophilic blocks to form mixed coronas can lead to even more

13

complicated nanostructures, such as Janus micelles.39 Although such assemblies are

outside the scope of this document, the reader is encouraged to read a recent review.40

Figure 1.8 Schematic model for charged block length-dependent recognition to form

BICs and BIC self-assembly.37

1.5 IPECs and BICs for Nucleic Acid Delivery

In addition to synthetic polyanions, polycations can also form IPECs and BICs

with nucleic acids by electrostatically complexing with the negatively charged

phosphodiester backbone. This relatively simple nucleic acid packaging strategy makes it

a rather obvious choice for investigation. Indeed as early as 1965, tertiary amine-

functionalized dextran was being used to deliver viral RNA to cells.41 By 1975, a range

of cationic polymers was being used to transfect mammalian DNA.42 Even with the

success of these rudimentary “off the shelf” polymers, researchers quickly began to

recognize the aforementioned barriers to delivery, and thus the focus shifted to more

specialized polymers to counter them. At this point, it is worth noting that while many of

14

the following studies focus on the delivery of DNA, the majority of the conclusions are

equally applicable to siRNA and dsRNA delivery.

RNA Protection

Perhaps the most obvious benefit of polycation complexation with a nucleic acid

is that it affords a significant degree of protection. Complexation greatly stabilizes

nucleic acids43,44 and sterically hinders nuclease access to them, preventing its catalyzed

degradation (Figure 1.9).44–47 Such protection is likely enhanced in BICs, where the

neutrally charged corona provides even greater steric hindrance. Furthermore, commonly

used hydrophilic blocks, like poly(N-(2-hydroxypropyl)methacrylamide) (PHPMA)48 and

poly(ethylene glycol) (PEG),49,50 often go unrecognized by the immune system,

improving the likelihood of the vehicle reaching its target destination. Additionally,

complexation resolves the issue of circulation time in humans: the threshold for renal

clearance is 6 nm,51 but nucleic acid complexation results in particle sizes ranging from

~11 nm for siRNA46 to hundreds of nm for DNA.52 Consequently, such IPECs and BICs

have prolonged circulation times and tend to accumulate in tumor tissue due to the

enhanced permeation and retention (EPR) effect.53,54

15

Figure 1.9 Enzymatic degradation of small interfering ribonucleic acid with RNase A in

the presence and absence of (HPMA258-b-DMAPMA13)46

Cell Transfection

IPEC and BIC formation with nucleic acids can facilitate cell transfection.

Although the mechanism is not fully understood, cationic IPECs (Z > 1) can readily enter

cells by interacting with either cellular membrane proteins55,56 or the negatively charged

membrane surface.57 While this results in cationic IPECs having a higher transfection

efficiency than their BIC counterparts,58,59 their disruption of the cellular membrane leads

to increase cytotoxicity.60,61 Thus, neutrally charged BICs (Z = 1) have been favored for

gene delivery more recently. However, their minimal cellular uptake requires a more

active approach to transfection.

One popular means of improving BIC transfection is to incorporate into the

polymer a moiety that targets membrane receptors to trigger endocytosis.16 Additionally,

if the targeted receptor is carefully chosen, the targeting moiety can also provide cell

specificity. For example, York et al.18 utilized folic acid to trigger receptor-mediated

endocytosis. Hydrophilic-block-cationic copolymers were synthesized using HPMA as

the hydrophilic block and N-[3-(Dimethylamino)propyl]methacrylamide (DMAPMA) as

16

the cationic block. N-(3-Aminopropyl)methacrylamide (APMA) was statistically

incorporated into the HPMA block to yield (HPMA-s-APMA)-b-DMAPMA copolymers

with primary amine handles for the covalent attachment of folic acid. After

functionalization with folic acid, the copolymers were used to form neutrally charged

BICs with siRNA (Figure 1.10). To demonstrate cell specificity, the BICs were incubated

with cells both overexpressing the folic acid receptor (KB) and minimally-expressing it

(A549). Whereas no siRNA could be detected in the A549 cells, siRNA presence was

readily apparent in KB cells (Figure 1.11), resulting in approximately 60% knockdown of

the targeted gene.

Figure 1.10 BIC formation from siRNA and (HPMA-s-APMA)-b-DMAPMA copolymers

functionalized with folic acid.18

17

Figure 1.11 Fluorescent microscope images of small interfering RNA (siRNA; cyanine-3

and fluorescein (FAM) labeled) delivery to KB cells (A, C, E) and A549 cells (B, D, F).

Lipofectamine (A, B; + control), unconjugated (N-(2-hydroxypropyl)methacrylamide315-

stat-N-(3-aminopropyl)methacrylamide13)-b-N-(3-

dimethylaminopropyl)methacrylamide23, (HPMA315-stat-APMA13)-b-DMAPMA23 (C, D;

− control), and folic acid conjugated (HPMA315-stat-APMA13)-b-DMAPMA23 (E, F).

Nuclei were stained with 4′,6-diamidino-2-phenylindole (blue). For clarity, FAM

fluorescence is not shown. Scale bars = 50 μm.18

To gain even more specificity in cell targeting, antibodies targeting specific

antigens expressed in cell membranes can be incorporated into polymers to produce

antibody-polymer conjugates (APCs). Lu et al.62 conjugated antibodies targeting HER2,

an epidermal growth factor receptor commonly overexpressed in breast cancers, to block

copolymers consisting of PEG and N-(3-guanidinopropyl)methacrylamide (GPMA)

(Figure 1.12). After complexing the APC block copolymers with siRNA, they

demonstrated cell-specific delivery (Figure 1.13) and up to 88% gene knockdown.

18

Figure 1.12 (A) Structure of pAcF. (B) Schematic illustration of pAcF- mutated anti-

HER2 antibodies, S202-pAcF Fab, Q389-pAcF IgG, and A121-pAcF IgG. (C) Synthetic

scheme for the antibody−polymer conjugates.62

Figure 1.13 Confocal microscopy of internalization of siRNA mediated by S202-Fab-P1.

HeLa (A,C,E) and SKBR-3 (B,D,F) cells were treated with buffer (A,B), 200 nM S202-

pAcF Fab + 200 nM P1 +50 nM siRNA-FITC (C,D), or 200 nM S202-Fab-P1 + 50 nM

19

siRNA- FITC (E,F) for 4 h. Cells were then stained with Hoechst 33342 (blue) and ER-

Tracker (red) and imaged with a Leica 710 confocal microscope. Bar = 10 μm.62

Another strategy to induce cellular uptake is to employ cell-penetrating peptide

(CPP) moieties or their mimics. CPPs are small cationic peptides that are able to pass

through cell membranes, often nonendocytotically.63 Inspired by guanidinium-rich CPPs,

Treat et al.64 synthesized FITC-labeled HPMA-b-GPMA copolymers and demonstrated

cellular transfection by both endocytotic and nonendocytotic pathways (Figure 1.14).

While they did not attempt siRNA delivery, several other groups have demonstrated

successful gene knockdown using guanidinium-rich IPECs and BICs to deliver siRNA.65–

68 CPPs and their mimics may also aid in endosomal escape in a similar fashion or by

bypassing the endosome altogether.64

Figure 1.14 Cellular uptake of FITC-labeled HPMA-b-GPMA copolymers.64

Endosomal Escape

IPECs and BICs are unique among nucleic acid carriers in that many tend to

innately promote endosomal escape.69 A popular theory for this ability is the so-called

“proton sponge” effect, in which polycations with protonatable groups act as buffers

during endosomal acidification. As a result of the polymers’ absorption of protons,

additional protons and chloride ions are pumped into the endosome, resulting in osmotic

swelling and eventual membrane rupture.70 While this theory is appealing, it does not

20

account for endosomal escape by IPECs and BICs lacking protonatable groups or that are

already fully protonated at physiological pH. It is far more likely that the negatively

charged endosomal membrane acts similar to a polyanion and undergoes a substitution

reaction with the IPEC or BIC carrier. The polycation, now “complexed” with the

endosomal membrane, disrupts the flow of the fluid-like liquid bilayer, causing it to leak

and/or fall apart. Conveniently, this should also facilitate release of the nucleic acid

payload (vide infra). Similarly, the polycations could also exchange with endosomal

transmembrane proteins to disrupt the membrane (Figure 1.15). While neither mechanism

has been extensively investigated, there is evidence for both types of interaction at the

cell membrane.12,55 Thus, it is not unreasonable that they should also occur in the

narrower confines of the endosome.

Figure 1.15 Scheme of polyelectrolyte-membrane protein interaction.12

Alternatively, pH-responsive moieties can be incorporated into the polymers for a

more active approach to endosomal escape by exploiting the decrease in pH during

endosome formation. For example, Holley et al.71 synthesized block copolymers

21

consisting of HPMA and L-Glu that, under endosomal pH conditions, formed

hydrophobic α-helices that can embed themselves in the endosomal membrane (Figure

1.16). Using circular dichroism (CD) spectroscopy, they demonstrated α-helix formation

with decreasing pH (Figure1.17). Incubation of these polymers under acidic conditions

with both artificial lipid membranes as well as red blood cells (RBCs) resulted in

membrane disruption and leakage.

Figure 1.16 HPMA-b-L-Glu copolymers form α-helices at low pH, allowing them to

embed themselves in cell membranes.71

Figure 1.17 Mean residue ellipticity as a function of pH for poly[HPMA220-b-(L-

Glu56)].71

Similarly, Convertine et al.72 used N-[3-(Dimethylamino)ethyl] methacrylate

(DMAEMA), propacrylic acid (PAA), and butyl methacrylate (BMA) to synthesize

22

DMAEMA-b-(DMAEMA-co-PAA-co-BMA) terpolymers (Figure 1.18) of varying

monomer content that formed pH-responsive IPECs with siRNA. Incubation of RBCs in

the presence of the IPECs at varying pH resulted in extensive hemolysis under endosomal

pH conditions and none at physiological pH. Additionally, delivery of siRNA to HeLa

cells by the terpolymers resulted in up to 88% gene knockdown for the polymers with the

highest hydrophobic content.

Figure 1.18 RAFT-mediated synthesis of diblock copolymers consisting of a cationic

poly(dimethylaminoethyl methacrylate) (DMAEMA, x=58) block and an endosomolytic

polyampholyte block incorporating DMAEMA and propylacrylic acid (PAA) in

equimolar ratios, and butyl methacrylate (BMA) (y~70)72

Payload Release

Once endosomal escape is achieved, the nucleic acid payload must be released

from the complex. Full IPEC dissociation occurs at high ionic strength,30 and although

the ionic strength of the cytosol tends to be higher than that of the extracellular fluid,

cytosolic ion concentrations rarely reach such levels.73 Rather, the most popular theory is

that nucleic acid release occurs via IPEC substitution reactions with biomacromolecular

polyelectrolytes within the cytosol, such as proteins, other nucleic acids, and anionic

23

polysaccharides.9,21,74 In fact, competitive binding assays with heparin are often used to

screen IPECs for their ability to release RNA or DNA efficiently.75,76 Because increased

polyelectrolyte length results in slower IPEC substitution/exchange reactions,30 which

should protect nucleic acids from enzymatic degradation at the expense of release rate,

there is thought to be an optimal balance between IPEC/BIC binding strength and release

rate.43,75,77–79 For example, Strand et al.75 demonstrated reduced plasmid DNA (pDNA)

transfection with increasing dextran length. Substitution of uncharged saccharide units

into higher molecular weight dextran restored its ability to transfect. Similarly, Sprouse et

al.79 demonstrated that some statistical copolymers formed from neutrally hydrophilic

and cationic monomers increase pDNA transfection over their block copolymer analogs,

yet others resulted in decreased plasmid expression, presumably due to premature

complex dissociation. Our group recently explored the effect of cationic block length in

siRNA-containing BICs: longer cationic blocks led to decreased knockdown rates but did

not affect total knockdown levels.43 The effect of cationic block charge density in siRNA-

containing BICs is the subject of Chapter 4.1.

Additionally, as with endosomal escape, a more active approach can be taken to

IPEC/BIC dissociation through the incorporation of degradable polymer backbones or

cationic group spacers.9 The endosomal pH change can be exploited using acid-labile

functional groups like carbonates, acetals, and hydrazones,80–85 but nucleic acid release in

the endosome could lead to early degradation. A more viable approach is to exploit the

difference in reductive environment between the extra- and intracellular fluids. The

cytosol contains relatively high levels of disulfide-reducing molecules like

glutathione,86,87 and thus disulfides within polymers can be cleaved selectively within the

24

cytosol.88 Lin et al.89 exploited this reductive difference for IPEC release by synthesizing

poly(amido amine) (PAAm) with repetitive disulfide linkages in the main chain (SS-

PAAm) that are cleaved under intracellular conditions (Figure 1.19). Because small

molecule electrolyte dissociation constants are non-zero,30 cleavage of the SS-PAAm

backbone while complexed with pDNA resulted IPEC dissociation and up to

approximately 9-fold higher transfection efficiency compared to a PDMAEMA standard.

Figure 1.19 (a) Formation of SS-PAAs/DNA polyplexes that are stable in the

extracellular environment. (b) Intracellular reduction of the disulfide linkages in the

polymer of the polyplex. (c) Dissociation of DNA from the degraded polymer.89

In an alternative approach, Truong et al.90 synthesized IPECs that would self-

catalyze the degradation of their cationic group spacer for a timed approach to release.

They synthesized varying length polymers of 2-dimethylaminoethyl acrylate (DMAEA),

which self-catalyzes its hydrolysis in water to acrylic acid, regardless of solution pH .91

Because hydrolysis of PDMAEA to PAA converts the polycation into a polyanion, IPECs

formed with oligo-DNA exhibited full dissociation ranging from 24-48 hr. depending on

PDMAEA length (Figure 1.20). Conversely, PDMAEA whose amines had been

quaternized exhibited no complex dissociation, even after a week in water.

25

Figure 1.20 IPECs formed from oligo-DNA and PDMAEA dissociate because of self-

catalyzed hydrolysis in water, whereas those formed from quaternized PDMAEA do

not.90

1.6 RAFT Polymerization

Optimization of IPECs and BICs to overcome the barriers to delivering genetic

material requires polymer syntheses that afford precise polymer compositions,

predetermined molecular weights with narrow polymer dispersities, and advanced

architectures comprised of monomers possessing a wide variety of functional groups.

Early efforts were hindered by lack of access to such chemistries, but the advent of

reversible-deactivation radical polymerization (RDRP) techniques, including nitroxide-

mediated polymerization (NMP),92,93 atom transfer radical polymerization (ATRP),94–97

and reversible addition-fragmentation chain transfer (RAFT) polymerization,98–103 has

enabled polymer synthesis with the above parameters. Of the RDRP techniques, RAFT is

arguably the most versatile, tolerating a wide variety of functional groups to polymerize

styrenics, (meth)acrylates, (meth)acrylamides, acrylonitriles, vinyl esters, and vinyl

amides. While RAFT polymerizations often take place in organic solvents, aqueous

RAFT (aRAFT) allows for the polymerization of monomers with biologically relevant

26

functional groups, including amines, carboxylic acids, phosphates, and sulfonates,

directly in water.64,104–112 Thus, it is greatly suited for the synthesis of polycationic

(co)polymers for complexation with RNA.

Scheme 1.1 Mechanism of RAFT polymerization

Unlike other RDRP techniques that rely on reversible termination, RAFT grants

polymerization control through rapid degenerative chain transfer to a thiocarbonylthio

chain transfer agent (CTA, RAFT agent) to keep the majority of propagating chains in a

27

dormant state. The currently accepted RAFT mechanism, depicted in Scheme 1.1, begins

with the decomposition of an initiator to yield primary radical I˙. In an ideal RAFT

polymerization, I˙ adds directly to CTA to form intermediate radical 4, which, although

reversible, should quickly fragment to yield 9 and R˙. R˙ then adds to monomer M with

rate RI with constant kI to yield 8, which would then quickly undergo chain transfer with

another CTA species. Alternatively, I˙ can add directly to M before undergoing chain

transfer to yield a similar process. This pre-equilibrium phase is considered complete

when all CTA has been converted to macroCTA 6. As with all RDRP techniques, it is

important that Ri > Rp (rate of propagation with kp) to ensure that all propagating chains

initiate at approximately the same time. For the same reason, the pre-equilibrium phase

should be relatively short. Once the steady-state radical concentration has been reached,

the polymerization proceeds according to the main equilibrium with pseudo-first order

kinetics defined by propagation rate Rp:

𝑅𝑝 = 𝑘𝑎𝑝𝑝[𝑀]

(4)

Under these conditions, the equilibrium heavily favors the dormant species, and

molecular weight increases linearly with monomer conversion. Although I˙ can add

directly to M to produce propagating chains, the vast majority of chains are assumed to

be derived from the fragmented CTA R-group. Thus, degree of polymerization (DP) is

calculated as

𝐷𝑃 =

𝜌[𝑀]0

[𝐶𝑇𝐴]

(5)

where ρ is monomer conversion and [M]0 is initial monomer concentration. Similarly,

number average molecular weight (Mn) is calculated as

28

𝑀𝑛 =𝜌[𝑀]

0

[𝐶𝑇𝐴]𝑀𝑀𝑊 + 𝐶𝑇𝐴𝑀𝑊

(6)

where MMW and CTAMW are the molecular weights of the monomer and CTA

respectively. Because in reality permanent CTA loss is inevitable, Equation 6 often

underestimates Mn.113 Successful RAFT polymerization will yield polymers whose α-

and ω-termini are functionalized with the R-group and thiocarbonylthio group

respectively. Although some chains are in fact initiated by I˙ and will thus have and I-

functionalized α-terminus, α-termini can all be made identical by selecting an initiator

such that I˙ and R˙ are identical.

The problem of initiator-derived chains can be avoided altogether by using an

irradiation source to fragment the CTA directly, making R˙ the primary radical source

without the need for an exogenous inititiator. γ,114 UV,115 and visible light116,117 radiation

are all capable of photolysing CTAs, especially trithiocarbonates (TTCs), and under such

initiating conditions, the RAFT agent behaves as an iniferter (INitiator-chain transFER

agent-TERminator), resulting in a hybrid RAFT/iniferter polymerization mechanism

(Scheme 1.2).

Scheme 1.2 Mechanism of light-mediated RAFT polymerization

29

Because R˙ is the primary radical source, light-mediated RAFT ensures that all

chains have CTA-derived α-termini. Additionally, the use of an irradiation sources

provides spatiotemporal control over the reaction and allows it to be performed at

ambient temperature. Visible light is of particular interest, not only because it

circumvents many of the side reactions that can occur under higher frequency radiation,

but also because it allows the polymerization to be performed under biologically-friendly

conditions if performed in water.

30

CHAPTER II – OBJECTIVES OF RESEARCH

RNA interference, with the power to suppress virtually any gene of interest, has

resulted in much research towards its implementation in pharmaceuticals and in crop

insecticides. However, a number of barriers inhibit its implementation in both fields,

namely RNA cargo protection, cell transfection, endosomal escape, and payload release

into the cytosol. A number of RNA delivery vehicles have been developed to combat

these barriers, and among them, cationic polymers for the formation of

interpolyelectrolyte complexes have been the most promising. Such IPECs are favored

both for their relative ease of preparation as well as inherent properties that often counter

the barriers to RNAi. Furthermore, the available chemistries for polycation synthesis

allow them to be tailored specifically to the desired application. Among polymerization

techniques, RAFT, and more specifically aRAFT, has emerged as one of the most

promising for its facile preparation of cationic (co)polymers with pre-determined

molecular weights, narrow dispersities, and access to a wide variety of functional groups

that allow for additional post-polymerization functionalization. A number of groups,

including the McCormick group, have developed polymeric cationic delivery vehicles

using RAFT and aRAFT to significantly enhance RNA delivery. While most of these

efforts have focused on the development of technologies to overcome the barriers to

RNAi, few have placed emphasis on the fundamental structure-behavioral relationships

that govern RNA delivery. Such emphasis has been the focus of recent work in the

McCormick group.

This dissertation seeks to continue to ascertain such structure-property

relationships affecting RNA delivery, as well as applying such insights toward enabling

31

RNAi in crop pest insects that remain highly resistant to such treatment. The work

addressed herein is divided into two main sections. The first section expands upon

previous McCormick group work on the structure-property relationships involved in

IPEC dissociation in the cellular environment. A series of hydrophilic-block-cationic

copolymers were synthesized via aRAFT polymerization in which the cationic block

statistically incorporates increasing amounts of neutral, hydrophilic monomer such that

the number of cationic groups remains unchanged but the cationic charge density is

diluted along the polymer backbone. These copolymers were intended to improve siRNA

release within cells, leading to greater gene suppression, but they instead revealed greater

insight into the barriers to siRNA delivery at the cellular level. The second section

applies the knowledge gained through previous binding and release studies to design a

polycation capable of navigating the hostile biology of the lepidopteran gut. Because the

aforementioned barriers to RNAi are substantially higher during caterpillar ingestion, a

cationic homopolymer was synthesized to address these specific conditions to protect and

deliver dsRNA through oral feeding. These experiments enabled extensive gene

knockdown in a lepidopteran species that has, until now, remained completely refractory

toward RNAi-based control.

The specific objectives of this research are the following:

1. Prepare a well-defined macroCTA comprised of HPMA and APMA and chain extend

it with varying ratios of HPMA and DMAPMA to afford a series of hydrophilic-

block-cationic copolymers in which the cationic block statistically incorporates

increasing amounts of neutral, hydrophilic monomer such that the number of cationic

32

groups remains unchanged but the cationic charge density is diluted along the

polymer backbone.

2. Functionalize said polymers with the cellular targeting group folic acid and prepare

BICs with siRNA and siRNA analogs.

3. Characterize BICs with respect to size and charge using dynamic light scattering, ζ-

potential, and gel electrophoresis.

4. Determine structure-property relationships governing binding strength, electrostatic

complex dissociation, and gene suppression utilizing solution differential scanning

calorimetry, potentiometric titration, confocal microscopy, cellular fractionation,

circular dichroism spectroscopy, and RT-qPCR.

5. Prepare a well-defined pGPMA homopolymer for the complexation and delivery of

dsRNA to the lepidopteran gut.

6. Prepare and characterize IPECs with dsRNA with respect to stoichiometry, size, and

charge using gel electrophoresis and dynamic light scattering.

7. Evaluate ability of pGPMA IPECs to transfect and elicit RNAi in lepidopteran cells

in vitro using confocal microscopy and RT-qPCR.

8. Establish pGPMA toxicity, IPEC-induced gene knockdown, and IPEC mortality in

live caterpillars through feeding experiments, dissection, and RT-qPCR.

33

CHAPTER III - EXPERIMENTAL



density on gene suppression

3.1.1 Materials

All reagents were purchased from Sigma-Alridch and used as received unless

otherwise noted. 4,4’-Azobiscyanovaleric acid (V-501) was purchased from Wako and

was recrystallized twice from methanol. Azobisisobutyronitrile (AIBN) was

recrystallized from methanol. N-(3-aminopropyl)methacrylamide hydrochloride (APMA)

was purchased from Polysciences. N,N-(3-dimethylaminopropyl)methacrylamide

(DMAPMA) and triethylamine (TEA) were distilled prior to use. 4-cyano-4-

[(ethylsulfanylthiocarbonyl)sulfanyl]pentanoic acid (CEP),72 di-N-hydroxysuccinimide-

activated folic acid (diNHS-FA),18 and N-(2-hydroxypropyl)methacrylamide (HPMA)118

were synthesized according to literature procedures. Sodium polystyrene sulfonate (PSS)

(Mn = 14.2 kDa, Ð = 1.13) was purchased from Scientific Polymer Products. HPLC

purified oligonucleotides (siRNA against human survivin; unlabelled and AlexaFluor594-

labeled, pre-diced siRNA; and oligomeric dsDNA) were purchased from Integrated DNA

Technologies, Inc. The siRNA sequences targeting human survivin are as follows: Sense

strand 5′-AGCCCUUUCUCAAGGACCACCGCAUCU-3′ and the antisense strand 3′-

UUUCGGGAAAGAGUUCCUGGUGGCGUAGAGGA-5′. The pre-diced siRNA

sequences are as follows: Sense strand 5'-GCUGGACUCCUUCAUCAACdTdT-3' and

the antisense strand 3'-dTdTCGACCUGAGGAAGUAGUUG-5' (“dT” indicates

deoxythiamine DNA base). The dsDNA sequences are as follows: Sense strand 5’-

34

AGATGTGCAATTTTGCTACCGCATCT-3’ and the antisense strand 5’-

AGGAGATGCGGTAGCAAAAGTTGCACATCTTT-3’. Oligonucelotides (siRNA and

dsDNA) were heated at 95 °C for 10 min and were allowed to slowly cool to room

temperature prior to use. Concentrations of oligonucleotide (siRNA and dsDNA) are

reported as duplex concentrations unless otherwise noted. Gibco® RPMI 1640 cell

culture media (with and without folic acid) and fetal bovine serum (FBS) were purchased

from Life Technologies Corporation. KB cells were purchased from ATCC. For reactions

requiring nitrogen, ultrahigh purity nitrogen (purity ≥ 99.998%) was used. Spectra/Por®

regenerated cellulose dialysis membranes (Spectrum Laboratories, Inc) with a molecular

weight cut-off of 12-14 kDa were used for dialysis.

3.1.2 Polymer Synthesis

3.1.2.1 Synthesis of poly(HPMA-stat-APMA) macroCTA

The macro chain transfer agent (macroCTA) was prepared employing V-501 as

the primary radical source and CEP as the chain transfer agent at 70 °C. HPMA (12.61 g,

95.1 mmol) and APMA (894 mg, 5.0 mmol) were added to a 250 mL round-bottomed

flask and dissolved in 1 M acetate buffer (pH = 4.5) with a final volume of 100 mL ([M]0

= 1 M). The initial feed composition was 95 mol % HPMA and 5 mol % APMA. The

round-bottomed flask was septum-sealed and purged with nitrogen for 1 hour prior to

polymerization. The macroCTA was prepared with a [M]0/[CTA] ratio = 400 while the

[CTA]/[I] ratio was kept at 5, and the reaction was allowed to proceed for 5 h. The

polymerization was quenched by rapid cooling in liquid nitrogen followed by exposure to

air. The macroCTA was isolated by dialysis (pH = 3-4) at 4 °C and recovered by

lyophilization.

35

3.1.2.2 Synthesis of poly[(HPMA-stat-APMA)-block-(HPMA-stat-DMAPMA)]

copolymers (P100, P75, P50, P25, and P0)

The poly(HPMA-stat-APMA) macroCTA was chain extended with HPMA and/or

DMAPMA using V-501 as the primary radical source at 70 °C. The macroCTA, HPMA,

and DMAPMA were dissolved in acetate buffer to give a total [M]0 = 1 M. The HPMA

and DMAPMA initial feed compositions were adjusted to 100 mol % DMAPMA (P100);

75 mol % DMAPMA and 25 mol % HPMA (P75); 50 mol % DMAPMA and 50 mol %

HPMA (P50); 25 mol % DMAPMA and 75 mol % HPMA (P25); and 100 mol % HPMA

(P0). The round-bottomed flask was septum-sealed and subsequently purged with

nitrogen for 1 h prior to polymerization. Block copolymers were prepared with

[M]0/[CTA] = 200 while [CTA]/[I] was kept at 5. Each polymerization was terminated at

predetermined time intervals by rapid cooling in liquid nitrogen and subsequent exposure

to air. The poly[(HPMA-stat-APMA)-block-(HPMA-stat-DMAPMA)] copolymers were

purified by dialysis (pH = 3-4) at 4 °C and recovered by lyophilization.

Block copolymer end-groups were removed via a standard literature procedure.119

A typical reaction is as follows: poly[(HPMA226-stat-APMA7)-block-DMAPMA14]

(P100) (575 mg, 14.3 μmol) was added to a 25 mL round-bottomed flask and dissolved

in 6 mL of DMF. AIBN (70.4 mg, 0.429 mmol) was then added to the flask resulting in

an AIBN/copolymer ratio of 30:1. The solution was then septum-sealed, purged with

nitrogen for 1 h, and allowed to react at 70 °C for 4 h. The resulting copolymer was

precipitated from DMF into cold anhydrous diethyl ether three times.

36

3.1.2.3 Copolymer functionalization with folic acid

DiNHS-FA was prepared following a slightly modified literature prcedure.18

Briefly, folic acid (1.00 g, 2.3 mmol), NHS (1.30 g, 11.3 mmol), DCC (4.68 g, 22.7

mmol), and DMAP (277.5 mg, 2.3 mmol) were dissolved in 15 mL DMSO and stirred in

the dark at room temperature for 24 h. The dicyclohexylurea precipitate was filtered off

and the resulting solution was used without further purification.

The aforementioned diNHS-FA solution was then used to label the primary amine

moieties of the APMA units in the chain-terminated block copolymers. A typical reaction

is as follows: 49.5 mg (1.23 μmol) P100 was dissolved in 1 mL DMSO along with 5 μL

TEA to serve as a catalyst. 1.53 mL of the diNHS-FA solution was added dropwise and

the resulting solution was stirred in the dark at room temperature for 48 h. The reaction

was quenched by the addition of excess ammonium hydroxide (100% by volume), and

this reaction was carried out for 24 h. The resulting solution was then dialyzed against 0.6

M NaCl for 24 h, followed by dialysis against DI water for 3 days. The polymer was

recovered via lyophilization.

3.1.3 Formation of hydrophilic-block-cationic/oligonucleotide complexes

3.1.3.1 Preparation of copolymer-dsDNA complexes for solution differential

scanning calorimetry

Poly[(HPMA-stat-APMA)-block-(HPMA-stat-DMAPMA)]-dsDNA complexes

were prepared with N:P = 1 (i.e. neutral complexes). The dsDNA duplex concentration

was maintained at 75 μM for all complexes. A typical preparation is as follows: 177 μL

of a 1.785 mM poly[(HPMA226-stat-APMA7)-block-DMAPMA14] (P100) stock solution

was added to 375 μL of a 200 μM dsDNA stock. The solution was diluted with 448 μL

37

sodium cacadylate buffer, and the resulting dsDNA-copolymer complex solution was

vortexed and equilibrated for 30 min. After equilibration, the solution was degassed for

30 min prior to DSC measurements. The dsDNA and polymer stock solutions were

prepared in 10 mM sodium cacadylate buffer at pH 7.2.

3.1.3.2 Preparation of copolymer-siRNA complexes for gene suppression

Folic acid-labelled poly[(HPMA-stat-APMA)-block-(HPMA-stat-DMAPMA)]-

siRNA complexes were prepared with N:P = 1, and the siRNA concentration was

maintained at 100 nM. A typical preparation is as follows: 2.8 μL of a 71.43 μM P100

stock solution was added to 3.3 μL of a 20 μM siRNA stock solution. The complex

solution was gently mixed and equilibrated for 20 minutes prior to dilution with 214 μL

folate- and serum-free RPMI, followed by gentile mixing. The siRNA and polymer stock

solutions were prepared in 10 mM phosphate buffer (pH = 7.4).

3.1.4 Cell Culture

KB cells were maintained and proliferated in RPMI 1640 (with folic acid)

supplemented with 10% FBS at 37 °C in 95% air humidified atmosphere and 5% CO2.

3.1.5 Gene Suppression of Human Survivin

24 hours prior to treatment, the KB cell medium was replaced with folic acid-free

RPMI 1640 supplemented with 10% FBS. Cells (200,000 cells/mL, 500 μL) were seeded

in a 48 well plate (Corning Inc.). Cells were treated with 50 μL of a polymer-siRNA

complex solution. Lipofectamine 2000 (Invitrogen) was used as the positive control, and

the Lipofectamine-siRNA complexes were prepared according to manufacturer protocol.

The final siRNA concentration delivered was maintained at 100 nM. After 24 hours, total

RNA was extracted with TriZol (Invitrogen) following manufacturer protocol. Survivin

38

transcript abundance was determined using RT-qPCR. First strand cDNA was

synthesized with the Reverse Transcription Kit (Fermentas). Amplification and

quantification was carried out with a 2X qPCR mix containing SYBR green (Fisher

Scientific) and a BioRad CFX 96. The primer pairs for detecting the survivin gene were

5′-AGCCCTTTCTCAAGGACCAC and 5′-TCCTCTATGGGGTCGTCATC. PCR

primers for β-Actin gene were 5′-CATGTACGTTGCTATCCAGGC and 5′-

CTCCTTAATGTCACGCACGAT.

3.1.6 Fluorescence Microscopy

24 hours prior to treatment, the KB cell medium was replaced with folic acid-free

RPMI 1640 supplemented with 10% FBS. Cells (200,000 cells/mL, 2 mL) were seeded

on cover glasses in a 6 well plate (Corning Inc.). Cells were treated with 500 μL of a

polymer-siRNA (siRNA tagged with AlexaFluor594) complex solution. Lipofectamine

2000 (Invitrogen) was used as the positive control, and the Lipofectamine-siRNA

complexes were prepared according to manufacturer protocol. The final siRNA

concentration delivered was maintained at 100 nM. After 24 hours, the cells were fixed

with 4% formaldehyde and washed with PBS prior to imaging. The cells were then

stained with 12 μL of 4’,6-diamidino-2-phenylindole (DAPI) mounting medium. The

cover glasses were then placed on precleaned microscope slides for analysis.

Fluorescence cell images were taken using a Zeiss LSM 510 scanning confocal

microscope and processed with manufacturer software. Multiple fields were examined

for each sample to ensure uniform distribution of complexes throughout. Representative

areas were selected in quadruplicate, the fluorescence intensities were determined in

39

ImageJ, and the corrected total fluorescence (CTF) of each area was calculated according

to the relation:

CTF = Integrated Density − (area × background mean fluroescence).

Statistical variance between samples was calculated via a one-way ANOVA with Tukey

analysis in Minitab (version 17.1.0).

3.1.7 Cell Fractionation

Prior to cell treatment, the siRNA 5’-phosphate was substituted with 32P-

containing phosphate using polynucleotide Kinase (Fisher) and γ-32P ATP (6000

Ci/mmol) as the source of isotope. The KB cell medium was replaced with folic acid-free

RPMI 1640 supplemented with 10% FBS. Cells (200,000 cells/mL, 2 mL) were seeded in

a 6 well plate (Corning Inc.). After 24 hours, cells were treated with 500 μL of a radio-

labelled polymer-siRNA complex solution. Lipofectamine 2000 (Invitrogen) was used as

the positive control, and the Lipofectamine-siRNA complexes were prepared according

to manufacturer protocol. The final siRNA concentration delivered was maintained at 100

nM. After 24 hours, the cell media was removed, and the cells were lysed with 1 mL

lysing buffer (150 mM HEPES, pH = 8.0; 0.25% Triton X; 10% glycerol).

Linear sucrose gradients (10%-50% w/w in 25 mM Tris-HCl (pH = 7.5), 25 mM

NaCl, 5 mM MgCl2) were prepared by carefully layering 400 μL of each sucrose solution

in a Beckman 13 x 51 mm thickwall polycarbonate tube at 0 °C. Total cell lysates were

carefully overlaid onto the gradients and centrifuged at 36,000 rpm for 2 hrs at 4 °C in a

SW 55 Ti rotor. Gradient fractions were then collected in 300 μL increments, and total

RNA was precipitated into 1 mL of isopropanol, employing 1 μL glycogen solution as a

co-precipitant. After centrifugation, the supernatant was removed, and the precipitants

40

were suspended in 2X RNA loading buffer from Ambion. RNA was separated on a 12%

acrylamide gel containing 8 M urea, and visualized with ethidium bromide staining on a

BioRad ChemiDoc MP. The gel was then electroblotted and crosslinked. The radio-

labelled siRNA was imaged using a GE Healthcare Life Sciences Typhoon FLA-7000.

The relative amount of siRNA was quantified in bands corresponding to both free siRNA

and that loaded in protein complexes using densitometry software ImageQuant.

3.1.8 Copolymer Cytotoxicity

The anti-proliferative activities of poly[(HPMA-stat-APMA)-block-(HPMA-stat-

DMAPMA)] copolymers were determined following a standard literature procedure. 24

hours prior to treatment, the KB cell medium was replaced with folic acid-free RPMI

1640 supplemented with 10% FBS. Cells (200,000 cells/mL, 100 μL) were seeded in a 96

well plate (Corning Inc.). Cells were treated with 50 μL of a polymer stock solution at a

polymer concentration equivalent to that used in the gene suppression studies. Cell

proliferation was determined via a standard MTT assay (Vybrant MTT Cell Proliferation

Assay Kit; Invitrogen). Cells were incubated for 48 h and 72 h before adding 10 µL of a

12 mM MTT reagent to each well. The cells were further incubated for an additional 4 h,

followed by adding 100 µL of a SDS (10%)/HCl (0.01 M) solution to each well. The

absorbance was then determined utilizing a Biotek Synergy2 MultiMode Microplate

Reader. All studies were performed in triplicate.

3.1.9 Characterization

All polymers were characterized by aqueous size exclusion chromatography

(ASEC) with an eluent of 1 wt % acetic acid and 0.1 M Na2SO4 (aq) at a flow rate of 0.25

mL/min at 25 °C, Eprogen Inc. CATSEC columns (100, 300, and 1000 Å), a Wyatt

41

Optilab DSP interferometric refractometer (λ = 690 nm), and a Wyatt DAWN-DSP

multi-angle laser light scattering (MALLS) detector (λ = 633 nm). Absolute molecular

weights and molecular weight distributions were calculated using Wyatt Astra (version 4)

software. dn/dc measurements for all (co)polymers were performed utilizing a Wyatt

Optilab DSP interferometric refractometer (λ = 690 nm) at 25 °C and Wyatt DNDC

(version 5.90.03) software. Polymer monomer conversions were calculated by comparing

the area of the monomeric refractive index signal at t0 to the area at tf.

Copolymer compositions were determined using a Varian MercuryPLUS 300

MHz NMR spectrometer in D2O utilizing a delay time of 5 s. 1H NMR was used to

determine copolymer compositions by integration of the relative intensities of the

methyne proton resonances of HPMA at 3.75 ppm and the dimethyl proton resonances of

DMAPMA at 2.75 ppm. The number of monomer units were calculated as n = (mol% ×

Mn, Exp)/MWmonomer. Conjugation of folic acid to the block copolymers was verified via

UV-Vis spectroscopy using a PerkinElmer Lambda 35 spectrophotometer utilizing an

average extinction coefficient of 8000 M-1cm-1 for free folic acid in phosphate buffter (10

mM Pi, 100 mM NaCl, pH = 7.4). 1H NMR was performed using a Varian MercuryPLUS

300 MHz spectrometer in DMSO-d6 with a delay time of 5 s. The amount of conjugated

folic acid was estimated by integration of the methine proton resonance of HPMA at 3.75

ppm and the proton resonance of folic acid at 8.64 ppm (s, PtC7H, 1 1H). These values

were estimated by employing a Lorentzian/Gaussian line fit using MestReNova (version

6.0.2-5475).

Variable-angle dynamic light scattering (DLS) measurements of copolymer-

siRNA complexes under aqueous conditions were performed using an incident light of

42

633 nm from a Research Electro-Optics Model 31425 He-Ne laser operating at 35 mW.

The angular dependence (60°-120° in 10° increments) of the autocorrelation function was

determined with a Brookhaven Instruments BI-200SM goniometer with an Avalanche

photodiode detector and TurboCorr autocorrelator. DLS measurements were carried out

at a complex concentration (siRNA + block copolymer) of 1.0 mg/mL in phosphate

buffer (10 mM Pi, pH = 7.4) at 25 °C. The mutual diffusion coefficients (Dm) were

determined from the relation

Γ = Dmq2

in which Γ and q2 represent the decay rate of the autocorrelation function and the square

of the scalar magnitude of the scattering vector respectively. The hydrodynamic radius

(Rh) was then calculated from the Stokes-Einstein equation:

Dm ≈ D0 = (kBT)/(6πηRh)

in which η is the solution viscosity, kB is Boltzmann’s constant, and T is the temperature

in K. Samples were vortexed to ensure homogeneity and equilibrated for 30 min at 25 °C

prior to measurement. To remove dust, samples were passed through a 0.45 μm Millipore

filter (PVDF) directly into the scattering cells. Measurements were performed in

triplicate.

Zeta-potential measurements were carried out at a complex concentration of 1.0

mg/mL in phosphate buffer (10 mM Pi, pH = 7.4) using a Malvern Zetasizer Nano

ZEN3600. Samples were vortexed to ensure homogeneity and equilibrated for 30 min at

25 °C prior to measurement. To remove dust, samples were centrifuged at 14,000 RPM

for 10 min. Measurements were performed in triplicate.

43

Quantification of polymer-complexed siRNA was achieved using the Agilent

2100 Bioanalyzer platform with the Small RNA kit following manufacturer protocol.

Samples were prepared with [siRNA] = 100 nM, N:P = 1 in RNase-free water. Samples

were vortexed to ensure homogeneity and equilibrated for 30 min at 25 °C prior to

measurement. The free siRNA concentration was determined from the area of the peak at

~39 s using the companion software. Percent complexed siRNA was calculated as 1 –

[siRNA39s, complex]/[siRNA39s, control].

All calorimetric experiments were carried out using a Calorimetric Sciences

Corporation Nano DSC-II solution differential scanning calorimeter (DSC). Sodium

cacadylate buffer (10 mM, pH = 7.2) was used as the running buffer. dsDNA (analog for

siRNA) concentration was maintained at 75 μM while copolymer concentrations were

adjusted to maintain N:P = 1. CpCalc (Version 2.1, Calorimetric Sciences Corp.) was

used to subtract buffer-buffer scans from buffer-sample scans.

Potentiometric titration experiments were carried out using a Metrohm 848

Titrino Plus autotitrator. Polymer samples were prepared in 5.0 mL of 18.2 MΩ diH2O

and concentrations were adjusted to maintain a total amine concentration (i.e. DMAPMA

unit concentration) of 1 mM. The pH of the solution was adjusted to 2.0 via the addition

of 1 N HCl, followed by autotitration to pH = 12.0 with 0.05 N NaOH at 25 °C. For

polymer-polystyrene sulfonate (PSS) complex solutions, polymer stock solutions were

adjusted to pH = 2.0 with 1 N HCl before addition to PSS stock solutions to afford

neutral complexes (i.e. [DMAPMA] = [SS]) followed by dilution to 5.0 mL (final

DMAPMA unit concentration = 1 mM). The complex solutions were then autotitrated to

pH = 12 with 0.05 N NaOH. The degree of protonation (α) and degree of complexation

44

(θ) as a function of pH for each polymer or polymer-PSS complex solution was

determined from the titration curves according to literature procedure.120

The kinetics of degradation of free and complexed siRNA with Riboshredder

RNase blend (Epicentre) were obtained by monitoring time-dependent ellipticity at λ =

212 nm utilizing a Jasco J-815 circular dichroism spectropolarimeter. Samples (V = 200

μL) were prepared in phosphate buffer (10 mM Pi, pH = 7.4) with [siRNA] = 5.0 μM. For

complex solutions, the copolymer concentrations were adjusted to maintain N:P = 1.

Samples were placed in a 400 μL quartz cuvette (path length = 1 mm), and the initial

spectra from λ = 200-320 nm were recorded with a scan rate of 50 nm/min, a 0.5 nm

bandwidth, and a time constant of 2 s. The signal-to-noise was doubled for all spectra by

averaging four scans. After establishing a baseline, 0.63 μL of Riboshredder stock

solution (0.25 unit/μL diluted in 10 mM Pi, pH = 7.4) was added followed by inversion of

the cuvette to promote mixing. The ellipticities at λ = 212 nm were then recorded over 20

min. with a 0.5 nm bandwidth and a time constant of 2 s.


Resistant Insect Pests

3.2.1 Materials

All reagents were purchased from Sigma-Aldrich at the highest available purity

and used as received unless otherwise noted. 4-Cyano-4-

[(ethylsulfanylthiocarbonyl)sulfanyl]pentanoic acid (CEP)72 and N-(3-

guanidinopropyl)methacrylamide (GPMA)121 were synthesized as previously reported.

Gibco Sf-900 II serum free media was purchased from Fisher. Sf9 (Spodoptera frugipera,

ovarian) cells were purchased from Millipore. Fall armyworm (S. frugiperda) larvae were

45

obtained from Benzon Research through USDA permit P526P-17-00512. For reactions

requiring nitrogen, ultrahigh purity nitrogen (purity ≥ 99.998%) was used. Spectra/Por

regenerated cellulose dialysis membranes (Spectrum Laboratories, Inc) with a molecular

weight cut-off of 12-14 kDa were used for dialysis.

3.2.2 Synthesis of pGPMA

Poly[N-(3-guanidinopropyl)methacrylamide] (pGPMA) was prepared employing

4,4’-azobiscyanovaleric acid as the primary radical source and CEP as the chain transfer

agent at 70 °C. GPMA (1.46 g, 6.6 mmol), CEP (15.6 mg, 59.2 x 10-6 mol), and 4,4’-

azobiscyanovaleric acid (3.3 mg, 11.8 x 10-6 mol) were added to a 25 mL round-bottomed

flask and dissolved in 1 M acetate buffer (pH = 4.5) with 1 mL MeOH (to improve CTA

and initiator solubility) with a final volume of 10 mL ([M]0 = 0.65 M). 50 μL dioxane

was added as an internal standard for 1H NMR analysis. The round-bottomed flask was

septum-sealed and purged with nitrogen for 1 h prior to polymerization. The polymer was

prepared with [M]0/[CTA] = 110 while [CTA]/[I] was kept at 5, and the reaction was

allowed to proceed for 19 h. Aliquots were taken via degassed syringe to monitor

monomer conversion. The polymerization was quenched by rapid cooling in liquid

nitrogen followed by exposure to air. The product was isolated by dialysis (pH = 3-4) at 4

°C and recovered by lyophilization.

3.2.3 In vitro Transcription of dsRNA

Using Taq DNA polymerase, ~500 nucleotide (nt) of exonic sequence was

amplified by polymerase chain reaction (PCR) for GFP, and the Spodoptera frugiperda

genes: sfV-ATPase, sfKIF and sfCDC27 genes (See Appendix A.2.1). Fragments were

ligated into pGEM®-T Easy plasmid (Promega), and sequence verified. PCR products

46

were generated from these constructs to add T7 promoter sequences to create templates

for bidirectional transcription. Using MEGAscript T7 Transcription Kit (Thermo

Scientific), in vitro transcription reactions were carried out, followed by LiCl

precipitation of products. RNAs were resuspended in nuclease free water and denatured

at 95 °C. After 2 minutes, the heat block was turned off to allow gradual reduction of

temperature to anneal RNAs. Annealing was carried out for 1 h, after which purity,

concentration, and quality was determined via UV spectroscopy with a nanoDrop-1000

and gel electrophoresis. dsRNAs were stored at -80 °C.

3.2.4 Polymer Characterization

pGPMA was characterized by aqueous size exclusion chromatography (ASEC)

with an eluent of 1 wt % acetic acid and 0.1 M LiBr (aq) at a flow rate of 0.25 mL/min at

25 °C, Eprogen Inc. CATSEC columns (100, 300, and 1000 Å), a Wyatt Optilab DSP

interferometric refractometer (λ = 690 nm), and a Wyatt DAWN-DSP multi-angle laser

light scattering (MALLS) detector (λ = 633 nm). Absolute molecular weight and

molecular weight distribution were calculated using Wyatt Astra (version 4) software;

dn/dc measurement for polymer was performed utilizing a Wyatt Optilab DSP

interferometric refractometer (λ = 690 nm) at 25 °C and Wyatt DNDC (version 5.90.03)

software. 1H NMR spectroscopy was performed using a Varian MercuryPLUS 300 MHz

NMR spectrometer in D2O utilizing a delay time of 5 s. Monomer conversion was

calculated from the 1H NMR spectra by monitoring the disappearance of the GPMA vinyl

peaks (5.27 ppm and 5.51 ppm) relative to the dioxane internal standard (3.58 ppm)

(Figure A.7).

47

3.2.5 Light Scattering

Variable-angle dynamic light scattering (DLS) measurements of copolymer-

dsRNA complexes under aqueous conditions were performed using an incident light of

633 nm from a Research Electro-Optics Model 31425 He-Ne laser operating at 35 mW.

The angular dependence (60°-120° in 10° increments) of the autocorrelation function was

determined with a Brookhaven Instruments BI-200SM goniometer with an Avalanche

photodiode detector and TurboCorr autocorrelator. DLS measurements were carried out

at a complex concentration (dsRNA + polymer) of 0.1 mg/mL in phosphate buffer (10

mM Pi, pH = 7.4 or 10) at 25 °C. To remove dust, polymer and dsRNA solutions were

individually passed through a 0.45 μm Millipore filter (PVDF) directly into the scattering

cell. The solution was gently mixed and allowed to equilibrate for 30 min. prior to

analysis. The mutual diffusion coefficient (Dm) was determined from the relation

Γ = Dmq2

in which Γ and q2 represent the decay rate of the autocorrelation function and the square

of the scalar magnitude of the scattering vector respectively. The hydrodynamic radius

(Rh) was then calculated from the Stokes-Einstein equation:

Dm ≈ D0 = (kBT)/(6πηRh)

in which η is the solution viscosity, kB is Boltzmann’s constant, and T is temperature in

K.

Static light scattering (SLS) measurements were performed using the same

instrumentation and samples as described above. The angular dependence of the inverse

excess scattering intensity (Iex) was analyzed via Berry analysis by plotting Iex-1/2 vs. q2,

yielding the radius of gyration (Rg) from the slope.

48

Zeta-potential measurements were carried out a complex concentration (dsRNA +

polymer) of 0.1 mg/mL in phosphate buffer (10 mM Pi, pH = 7.4 or 10) at 25 °C using a

Malvern Zetasizer Nano ZEN3600. To remove dust, polymer and dsRNA solutions were

individually passed through a 0.45 μm Millipore filter (PVDF) directly into the folded

capillary cell. The solution was gently mixed and allowed to equilibrate for 30 min. prior

to analysis. Measurements were performed in triplicate.

3.2.6 Polymer-dsRNA Binding Assay

pGPMA-dsRNA solutions were prepared to complex 1 μg dsRNA at varying

polymer-dsRNA weight ratios (0.25-100 μg polymer/μg dsRNA, ± = 0.5-180). Briefly, an

appropriate volume of a 1 μg/μL or 10 μg/μL pGPMA stock solution in 10 mM PBS was

added to 2 μL of a 0.5 μg/μL dsRNA solution in nuclease-free diH2O. The solutions were

gently mixed and allowed to equilibrate for 30 min. before being diluted with 15 μL of 2x

RNA loading buffer (Ambion). Gel electrophoresis was then performed on a 1% agarose

gel in 1X TAE buffer stained with ethidium bromide. The gel was soaked in diH2O for 30

min to remove excess ethidium bromide before being imaged.

3.2.7 Gene Suppression in Sf9 Cell Culture

Sf9 cells were grown in Sf-900 II SFM at 28 °C. Sf9 cells (1 million cells/mL, 2

mL) were seeded in a 6 well plate (Corning Inc.). pGPMA-dsRNA complexes were

formed to deliver a total of 5 μg dsRNA complexed with 20, 30, or 40 μg pGPMA per

well. Briefly, 20, 30, or 40 μL of a 1 μg/μL pGPMA stock solution in 10 mM PBS was

added to 10 μL of a 0.5 μg/μL stock solution of dsRNA targeting CDC27 in nuclease-free

diH2O. The solution was gently mixed and allowed to equilibrate for 30 min before being

added to the cell media, resulting in [dsRNA] = 7.4 nM. Identical complex solutions

49

using dsRNA targeting KIF were used as controls. After 24 h, total RNA was extracted

with TRI Reagent following manufacturer protocol. CDC27 transcript abundance was

determined via RT-qPCR. First strand cDNA was synthesized with the Reverse

Transcription Kit (Fermentas). Amplification and quantification were carried out with

qPCR mix containing SYBR green (Fisher Scientific) and a BioRad CFX 96. All

amplifications were performed in quadruplicate (primers listed in Appendix A.2.1).

Time-dependent gene suppression followed a similar procedure. Cells were

seeded as described above, and pGPMA-dsRNA complexes targeting CDC27 were

formed to deliver a total of 5 μg dsRNA complexed with 40 μg pGPMA. Lipofectamine

3000 (Invitrogen) was used as a positive control, and the Lipofectamine-dsRNA

complexes were prepared according to manufacturer protocol. Untreated cells were used

as a negative control. After 24, 48, or 72 h, total RNA was extracted, and RT-qPCR was

performed as described above.

3.2.8 Cell Viability Assay

Cells (1M cells/mL, 100 μL) were seeded in a 96 well plate (Corning Inc.). Cells

were treated with 1, 1.5, or 2 μL of a 1 mg/mL pGPMA stock solution to yield polymer

concentrations equivalent to those used in the gene suppression studies. Cell proliferation

was determined via a standard MTT assay (Vybrant MTT Cell Proliferation Assay Kit;

Invitrogen). Cells were incubated for 48 h before adding 10 µL of a 12 mM MTT reagent

to each well. The cells were further incubated for an additional 4 h, followed by adding

100 µL of a SDS (10%)/HCl (0.01 M) solution to each well. The absorbance was then

determined utilizing a Biotek Synergy2 MultiMode Microplate Reader. All studies were

performed in triplicate.

50

3.2.9 Confocal Microscopy

Sf9 cells (200,000 cells/mL, 500 μL) were seeded in a 48 well plate (Corning

Inc.). pGPMA-dsRNA complexes were formed to deliver a total of 25 ng Cy5-labeled

dsRNA complexed with 150 ng pGPMA per well. Briefly, 1.5 μL of a 0.1 μg/μL pGPMA

stock solution in 10 mM PBS was added to 1.02 μL of a 24.5 μg/μL dsRNA solution in

nuclease-free diH2O. The solution was diluted to 25 μL with 10 mM PBS, gently mixed,

and allowed to equilibrate for 30 min. before being added to cell media. A 25 μL solution

containing 25 ng Cy5-labeled dsRNA was also prepared and added to cells as a control.

After 24 h, the cells were collected and spun down at 4.5k RPM. The supernatant was

removed, and the cells were washed with 500 μL PBS. After spinning down again, the

cells were resuspended in 40 μL PBS and placed on pre-cleaned microscope slides. The

cells were then fixed with 4% formaldehyde, washed with PBS, and stained with 12 μL

4’,6-diamidino-2-phenylindole (DAPI) mounting medium before adding cover slips.

Fluorescence cell images were taken using a Zeiss LSM 510 scanning confocal

microscope and processed with manufacturer software. Multiple fields were imaged for

each sample to document uniform cytoplasmic distribution of complexes.

3.2.10 Larval Feeding Experiments

pGPMA-dsRNA complexes targeting V-ATPase or GFP (control) were formed in

8:1 weight ratio as previously described. Fall armyworm larvae were immobilized, and

either pGPMA alone or pGPMA-dsRNA complex solution (~100 ng/μL dsRNA) was put

directly on larval mouth parts, and ingestion verified by observation under a

stereomicroscope. Animals were then kept in a 26 °C incubator on larval food. Insect

midguts were dissected and homogenized in TRI reagent for total RNA extraction

51

following manufacturer protocol. V-ATPase transcript abundance was determined via

RT-qPCR as described above. For survival assay, the number of larvae/pupae was

counted in regular intervals (days) for mortality.

52

CHAPTER IV – RESULTS AND DISCUSSION




4.1.1 Overview

RNA interference (RNAi) triggers post-transcriptional gene suppression via

sequence-specific recognition and destruction of cellular transcripts.2 “Gene knockdown”

is achieved through delivery of synthetic small interfering RNA (siRNA), which can be

designed to target the gene of interest,14,122–124 making RNAi appealing for gene

therapeutics. However, RNA delivery vehicles must overcome a number of barriers,

including target specificity and vehicle cytotoxocity.124

Polymeric vectors can provide both enhanced stability and decreased

immunogenic response relative to more traditional vectors (e.g. viral and lipid-based).14

Of particular interest are polycationic polymers that electrostatically complex the

negatively-charged RNA phosphodiester backbone to form interpolyelectrolyte

complexes (IPECs).13,60 Such IPECs are often characterized by the molar ratio of

cationic functionalities (e.g. amines) to phosphodiester units, termed the nitrogen-to-

phosphate ratio (N:P). Non-stoichiometric IPECs from cationic homopolymers have been

extensively studied and provide enhanced protection from enzymatic degradation while

maintaining complex hydrophilicity.14,125 However, the excess charges required to

maintain solubility result in adverse effects: negatively-charged complexes (N:P < 1)

suffer from decreased transfection due to electrostatic repulsion at the negatively-charged

cellular membrane, and positively-charged complexes (N:P > 1) result in increased

53

cytotoxity and opsonization within the blood stream, leading to higher immune

response.60,61,126,127 Block copolymers consisting of a cationic block and a non-ionic,

hydrophilic block can form stoichiometric, neutrally charged IPECs with RNAs while

maintaining complex hydrophilicity. These so-called block ionomer complexes (BICs)

exhibit both decreased cytotoxicity and enhanced stability,13,128 and incorporation of

cellular targeting moieties within their hydrophilic, corona-forming blocks results in cell-

specific siRNA delivery.18,125

Our research group has maintained a strong interest in the rational design and

synthesis of drug delivery systems utilizing aqueous RAFT (aRAFT) polymerization

targeting controlled, tailored (co)polymers for stimuli-responsive micelles,129–131

theranostics,47 peptide mimics,64 modular copolymers,132,133 and vehicles for endosomal

escape.71 Our most recent efforts have focused on the development of siRNA-containing

BICs for cell-specific delivery as well as determining the effect of aRAFT copolymer

architecture on siRNA delivery efficacy. Previously, we demonstrated targeted cellular

delivery and subsequent gene knockdown using BICs formed between siRNA and

hydrophilic-block-cationic copolymers.18 Furthermore, we observed a correlation

between cationic block length and siRNA stabilization as well as gene knockdown

efficacy: longer cationic block lengths resulted in increasingly enhanced siRNA stability

as well as longer time periods required to achieve maximum gene knockdown in vitro.43

We attributed delayed gene suppression to slow release of the siRNA from the

complexes, presumably via macromolecular exchange. This correlates well with other

groups’ findings that enhanced complexation and stability in plasmid DNA (pDNA)

delivery result in inefficient DNA release, indicating that intermediate binding and

54

stability is desireable.134–136 Such intermediacy is likely achievable via alteration of the

cationic charge density. Indeed, IPECs formed from polymers with varying degrees of

cationic quaternization yield higher pDNA transfection efficiency and expression at

moderate charge densities as compared to linear polycations.135,136 However, variable

charge density has not been studied in BICs, specifically those containing siRNA.

In this study, we report the synthesis of a series of hydrophilic-block-cationic

copolymers via aRAFT polymerization in which the cationic block statistically

incorporates increasing amounts of neutral, hydrophilic monomer such that the number of

cationic groups remains unchanged but the cationic charge density is diluted along the

polymer backbone. These polymers were subsequently complexed with siRNA and

siRNA analogs. To our knowledge, this is the first study directed toward elucidating the

effect of cationic block charge density on BIC binding strength/stability and siRNA

delivery. siRNA stability and BIC binding strength were evaluated utilizing solution

differential scanning calorimetry (DSC) and potentiometric titration respectively, and

cellular siRNA delivery experiments were performed to correlate those results with gene

knockdown efficacy. Herein, we demonstrate reduced siRNA stability, binding strength,

and gene knockdown with decreasing cationic block charge density. We correlate these

trends to reduced siRNA delivery and uptake within the RNAi pathway, which suggests

greater siRNA vulnerability to enzymatic degradation. Indeed, we confirm higher rates of

enzymatic hydrolysis with reduced cationic charge density by establishing RNase

degradation kinetic profiles. We conclude that while reduced binding strength results in

more rapid siRNA release via macromolecular exchange, such facile exchange increases

the likelihood of degradation prior to activation of the RNAi pathway.

55

4.1.2 Synthesis of hydrophilic-block-cationic copolymers with varying cationic block

charge density

Based upon our previous observation that decreasing cationic block length

reduces the time required to achieve maximum gene knockdown,43 we reasoned that

reduced cationic block charge density should decrease BIC binding strength, facilitating

the release of siRNA from the complexes via more rapid macromolecular exchange.

Scheme 4.1 Synthetic pathway for the preparation of poly[(HPMA-stat-APMA)-block-

(HPMA-stat-DMAPMA)] copolymers and subsequent complexation with siRNA.

We therefore synthesized hydrophilic-block-cationic copolymers with varying

cationic block charge density (Scheme 4.1). The first step was accomplished using

aRAFT to prepare a statistical macroCTA consisting of an initial monomer feed ratio of

95 mol % N-(2-hydroxypropyl)methacrylamide (HPMA) and 5 mol % N-(3-

aminopropyl)methacrylamide (APMA) in 1 M acetate buffer (pH = 4.5) at 70 °C using 4-

cyano-4-[(ethylsulfanylthiocarbonyl)sulfanyl]pentanoic acid (CEP) as the CTA and 4,4’-

asobiscyanovaleric acid (V-501) as the initiator. HPMA contributes non-ionic

56

hydrophilicity to the copolymer, and is known to be non-immunogenic,48 promoting

greater biocompatibility. Incorporation of the primary amine functionality of APMA

provides a convenient handle for the conjugation of the cellular-targeting moiety folic

acid. 1H NMR analysis revealed a final copolymer composition of 97 mol % HPMA and

3 mol % APMA, which closely matches the monomer feed ratio.

The resulting poly(HPMA226-stat-APMA7) macroCTA was subsequently

subjected to a series of chain extensions with both HPMA and N,N-(3-

dimethylaminopropyl)methacrylamide (DMAPMA), targeting DMAPMA monomer

molar feeds, and thus charge densities, of 100% (P100), 75% (P75), 50% (P50), 25%

(P25), and 0% (P0, Appendix A.1.1). The tertiary amines of DMAPMA provide cationic

sites under physiological conditions (pH = ~7.4) for complexation with the negatively-

charged siRNA backbone. Additionally, the statistical incorporation of HPMA within the

cationic block allows for increased spacing of the cationic groups, and thus lower charge

density, along the polymer backbone while minimizing inter- and intramolecular

hydrophobic interactions of the copolymers. ASEC-MALLS chromatograms for the

macroCTA and the chain extensions are shown in Figure 4.1, and the relevant polymer

characterization data are summarized in Table 4.1. Shifts to lower elution volume while

maintaining low dispersities (Ð < 1.2) indicate successful chain extension, and 1H NMR

analysis (Figure A.1) revealed block compositions (block A: HPMA-stat-APMA; block

B: HPMA-stat-DMAPMA) closely matching the monomer feed ratios. Cationic block

charge densities are reported as molar percentages of DMAPMA within block B.

57

Table 4.1

Molecular weight (number average), dispersity (Ð), composition, and dn/dc values for

macroCTA and chain-extended polymers

Sample Mn, Th

(kDa)a

Mn, Exp

(kDa)b Ð

Block A

Comp

(mol %)c

Block B

Comp

(mol %)c

ρd Charge

Density (%)

macroCTA 30.1 33.8 1.10 97:3 -- 0.51 --

P100 34.9 37.5 1.07 97:3 0:100 0.03 100

P75 35.5 38.7 1.12 97:3 27:73 0.05 73

P50 38.2 41.3 1.15 97:3 61:39 0.14 39

P25 44.0 46.3 1.16 97:3 78:22 0.34 22

P0 42.7 45.5 1.16 97:3 100:0 0.31 0 aTheoretical Mn, (Mn,Th), calculated from conversion (ρ) using Mn,Th = ([M]o/[CTA] × Mw,monomer × ρ) + Mw,CTA. bExperimental Mn

(Mn,Exp) was determined by aqueous SEC-MALLS. cAs determined by 1H NMR. d Conversions were determined by comparing the

area of the monomeric refractive index signal at t0 to the area at tf..

Figure 4.1 ASEC-MALLS of poly(HPMA-stat-APMA) macroCTA and subsequent chain

extensions with DMAPMA and HPMA (P100-P0)

The post-polymerization modification of APMA units with folic acid, which our

group has previously demonstrated to function well as a cell-specific targeting moiety,18

was monitored via UV-Vis spectroscopy (Figure A.2). Based on an average extinction

coefficient for free folic acid at pH = 7.4, approximately 4 of 7 possible APMA units per

58

polymer were successfully labelled. This extent of folic acid conjugation, combined with

low APMA molar content, resulted in hydrophilic-block-cationic copolymers capable of

electrostatically complexing with oligonucleotides through the DMAPMA tertiary amines

of cationic block B, while the hydrophilic, cellular-targeting block A maintains BIC

solubility. Thus, oligonucleotide complexation with these well-defined hydrophilic-

block-cationic copolymers with varying charge density allows for correlation of cationic

block charge density to BIC complexation strength and in vitro gene knockdown.

4.1.3 Hydrophilic-block-cationic copolymers with varying charge density form

stable, neutrally-charged complexes

Having successfully synthesized a series of hydrophilic-block-cationic

copolymers with varying cationic block charge density, we used dynamic light scattering

(DLS) and ζ-potential measurements to confirm their ability to complex siRNA while

maintaining charge neutrality at N:P = 1. Table 4.2 presents the hydrodynamic radius

(Rh) and ζ-potential of each copolymer-siRNA complex. The siRNA-containing BICs

exhibited an average Rh of 9.6 nm, a value consistent with previously reported complexes

of similar cationic content.18 Copolymer solutions free of siRNA did not exhibit any

particles visible by DLS (data not shown), indicating that the observed hydrodynamic

radii indeed result from complex formation rather than copolymer aggregation. The near-

zero ζ-potential values confirm complex charge neutrality, targeted for preventing

cytotoxicity. The amount of polymer-complexed siRNA was quantified using the Agilent

Bioanalyzer platform (electropherograms in Figure A.3), and copolymers P25-P100

complexed approximately 76% of available siRNA, which is comparable to our previous

report.46

59

Table 4.2

The hydrodynamic radii (Rh), ζ-potential, and percent complexed siRNA for siRNA and

copolymer-siRNA complexes

Sample Rh (nm) ζ-potential (mV) Complexed siRNA

P100 8.3 -1.02 75.0 %

P75 11.8 -0.44 75.9 %

P50 10.7 -1.82 79.3 %

P25 7.8 -0.91 75.7 %

P0 N/Aa -7.55 34.7 %

siRNA N/Aa -9.99 -- aThe excess scattering compared to solvent was too low for accurate determination.

4.1.4 Reduced cationic block charge density decreases oligonucleotide stabilization

Relative oligonucleotide stability can be determined by elucidating the melting

temperature (Tm), i.e. the temperature at which the double-stranded duplex separates into

its single-stranded components. An increase in Tm, which is manifested as an endotherm

maximum in the DSC thermogram, is indicative of increased duplex stability.137

Previous work in our laboratories used dsDNA as an analog to siRNA in order to

ascertain the effect of cationic block length of the copolymer on oligonucleotide

stability.43 In the present study, we used samples P0-P100 to prepare BICs with dsDNA,

and the respective DSC thermograms are shown in Figure 4.2. Complexation results in

increased Tm over that of free dsDNA (Tm = 54.4 °C).43 Generally, Tm decreases with

decreasing cationic block charge density (P75, 83.3 °C > P50, 81.3 °C > P25, 75.0 °C).

Although P100 has a continuous (i.e. no neutral comonomer) cationic block, its actual

number of charges (14 DMAPMA units) is lower than for polymers P25-P75 (~20

DMAPMA units), resulting in a Tm = 81.0 °C. However, BICs formed with previously

reported (HPMA171-stat-APMA13)-block-DMAPMA27 (P2), which has a longer

continuous cationic block, exhibit a dsDNA Tm value of 88.4 °C.43 Therefore, we may

60

conclude that the enhanced oligonucleotide stability afforded by complexation decreases

as cationic block charge density decreases.

Figure 4.2 Differential power thermograms for copolymer-dsDNA complexes. Samples

shifted along Y-axis for clarity.

4.1.5 Reduced cationic block charge density reduces complex binding strength

Having confirmed that reduced charge density diminishes the oligonucleotide-

stabilizing effect of complexation, we next sought to demonstrate that it similarly reduces

BIC binding strength as characterized by the free energy of complex formation. The

cationic nature of weak polyelectrolytes, such as those containing DMAPMA, is due to

the pH-dependent protonation of the amine functionalities and thus can be monitored via

potentiometric acid-base titration. From the potentiometric titrations of a polyelectrolyte

and its corresponding IPEC, one can obtain the degrees of protonation (α, fraction of

protonated amines) and complexation (θ, fraction of ionic complex pairs out of total

possible pairs) respectively. Kabanov and co-workers13 have demonstrated that for IPECs

consisting of a weak polyelectrolyte (e.g. PDMAPMA) and a strong polyelectrolyte (e.g.

61

siRNA), all of the protonated units of the weak polycation will form ionic pairs with a

strong polyanion functionality, i.e. α = θ. Due to the cooperativity of IPEC formation, a

shift (ΔpH(α)) occurs in the θ vs. pH curve of an IPEC relative to the α vs. pH curve of

the corresponding free polycation (Figure 4.3A). The free energy of complex formation

(ΔGtotal) as a function of α, where α = α1 (= θ1), is given by the following:13

∆𝐺𝑡𝑜𝑡𝑎𝑙 = −2.303𝑅𝑇 ∫ ∆pH(𝛼)𝑑𝛼

𝛼1

0

Evaluation of the binding strength of oligonucleotide-containing BICs by

potentiometric titration is complicated by the pH-dependent protonation of DNA and

RNA bases. Thus, in this study we have adopted an approach similar to that of Lee et

al.120 who used polystyrene sulfonate (PSS) as a strong polyanion analog that does not

affect the titration curve over the pH range investigated. We selected PSS with low

molecular weight (Mn = 16 kDa, DP ≈ 69) such that the number of anionic charges is

similar to that of duplex siRNA (59 nucleotides).

Figure 4.3 depicts the α- and θ vs. pH curves for P0-P100 and lists the free

energies of complexation for copolymer-PSS BICs. In general, the magnitude of free

energy decreases with decreasing cationic block charge density (P75, -3.28 kJ/mol; P50,

-2.42 kJ/mol; P25, -1.35 kJ/mol). Consistent with the DSC experiments in the previous

section, despite being a continuous cationic block, the fewer number of charges in P100

relative to P25-P75 results in a lower binding strength with a ΔGtotal value of -2.61

kJ/mol. However, titration of (HPMA171-stat-APMA13)-block-DMAPMA27 (P2)43 with a

longer continuous cationic block length yields a ΔGtotal value of -4.4 kJ/mol. Thus, we

may conclude that binding strength decreases with reduced cationic charge density.

62

Figure 4.3 3. α- and θ vs. pH curves for (A) P100, (B) P75, (C) P50, (D) P25, (E) P0,

and (F) P2 and their respective complexes with PSS

4.1.6 Reduced charge density diminishes gene knockdown efficacy

Based on the trends of decreasing oligonucleotide stabilization and BIC binding

strength with decreasing cationic block charge density, one would expect that decreased

charge density would lead to greater bioavailability of the siRNA within cells via more

rapid release and, therefore, enhanced gene knockdown. However, experimental results

63

were opposite of this expectation. Figure 4.4 depicts the relative survivin mRNA levels

24 hours after treatment with copolymer-siRNA BICs. P100 and P75 copolymers

resulted in 2- and 3-fold mRNA expression, respectively, relative to the Lipofectamine

positive control. However, polymers with charge density less than 75% exhibited no

decrease in survivin mRNA levels relative to the untreated negative control. Although

diminished gene knockdown with reduced charge density is the opposite of the expected

trend, these results are likely the result of more rapid macromolecular exchange due to

reduced binding strength: rapid exchange with extra- and intracellular proteins results in

reduced cellular delivery of siRNA and increased susceptibility to degradation by RNases

(vide infra).

Figure 4.4 RT-qPCR analysis of down-regulation of human survivin mRNA by

copolymer-siRNA complexes. mRNA expression normalized to Lipofectamine.

64

4.1.7 Cellular delivery and RNAi pathway activation decrease with reduced

cationic block charge density

To determine the relative cellular loading of siRNA by each polymer, cells were

treated with copolymer-(fluorescently-labelled siRNA) BICs and were subsequently

imaged via confocal fluorescence microscopy (images in Figure A.5). The corrected total

fluorescence (CTF) of representative areas for each treated cell culture is depicted in

Figure 4.5, and decreasing cellular siRNA content was observed with decreasing cationic

block charge density. Furthermore, statistical analysis of CTF revealed a significant

decrease in siRNA content between P75 and P50, which corresponds well to lack of gene

knockdown for copolymers with charge density < 75%. Because siRNA release must

result from a macromolecular exchange reaction rather than spontaneous dissociation,13

the decreased cellular delivery must be the result of exchange reactions with

biomacromolecules in the extracellular media. However, P50 and P25 successfully

delivered moderate amounts of siRNA, yet no gene knockdown was observed, suggesting

reduced siRNA participation in the RNAi pathway.

65

Figure 4.5 Corrected total fluorescence of siRNA labelled with AlexaFluor594 delivered

via copolymer complexes. Samples not belonging to same letter grouping were found to

have statistically significant variance via Tukey analysis.

Quantification of protein-bound siRNA within the cells serves as an indication of

the level of RNAi activity. Because the threshold in cationic block charge density

required for successful gene knockdown lies between P75 and P50, these copolymers

were used to deliver radio-labelled siRNA, and the treated cells were subjected to cellular

fractionation via sucrose density gradients. The fractions were then subjected to PAGE,

followed by electroblotting to quantify the relative amounts of radio-labelled siRNA in

each, depicted in Figure 4.6. Higher fraction numbers correspond to increased gradient

density; therefore, farther migration of siRNA into the heavier fractions is indicative of

siRNA-protein complexes (i.e. siRNA entry into the RNAi pathway). siRNA levels have

been normalized to fraction 1 (i.e. protein-free siRNA) for each cell culture. P75

complexes resulted in a greater amount of protein-complexed siRNA relative to its

protein-free siRNA than did P50, indicating that, in addition to increased cellular siRNA

concentration, P75 complexes resulted in a greater percentage of that siRNA participating

66

in RNAi. Therefore, as cationic block charge density decreases, less siRNA is trafficked

into the cells, and even less RNAi activation is achieved.

Figure 4.6 Relative radio-labelled siRNA content after copolymer complex delivery and

cell fractionation. Higher gradient numbers correspond to heavier fractions. siRNA

content normalized to fraction 1 for each complex.

When taken in conjunction, the fluorescence microscopy and cell fractionation

results suggest that instead of increasing siRNA bioavailability, decreasing cationic block

charge density leaves the siRNA more vulnerable to enzymatic degradation by RNases

within the cell culture media and within the cells themselves. Reineke and coworkers79

reported similar results utilizing hydrophilic-stat-cationic and hydrophilic-block-cationic

copolymers to deliver luciferase-expressing plasmid DNA (pDNA): statistical

copolymerization of their tertiary amine-containing monomer resulted in decreased

luciferase expression relative to the block copolymer. The authors suggested that

statistical copolymerization may have resulted in more rapid complex dissociation and

thus inefficient trafficking of the pDNA to the nucleus. Our demonstration of decreased

67

BIC binding strength with lower charge density, along with diminishing siRNA delivery

and RNAi activation, corroborates their conclusion: weaker binding likely results in more

rapid macromolecular exchange with cellular proteins like RNases.

4.1.8 Enzymatic degradation rates increase as cationic block charge density

decreases

Although siRNA degradation within cells cannot be directly observed, circular

dichroism (CD) spectroscopy can be used to monitor the degradation kinetics of siRNA

by RNases in vitro. The characteristic CD spectrum peaks of siRNA result from its

secondary structure,138 and thus we monitored the molar ellipticity at 212 nm ([θ]212) as

the siRNA was hydrolyzed along its phosphodiester backbone by Riboshredder RNase

blend (Figure 4.7A). Figure 4.7B depicts normalized [θ]212 of siRNA and copolymer-

siRNA complexes as a function of time. Based upon the gene knockdown, cellular

loading, and cell fractionation experiments, we expected to see an increase in the rate of

degradation with decreasing cationic block charge density. Indeed the decay rate of [θ]212

increases from P100 to P0, indicating that decreased charge density does result in

decreased protection from enzymatic degradation. This notion is in good agreement with

the decreased oligonucleotide stabilization and binding strength, determined via solution

DSC and potentiometric titration respectively, as a function of decreasing charge density.

68

Figure 4.7 (A) Molar ellipticity of siRNA before and 20 minutes after addition of

Riboshredder RNase blend. (B) Enzymatic degradation of free and copolymer-complexed

siRNA with Riboshredder RNase blend as monitored by the normalized disappearance of

the CD band at 212 nm.

69



4.2.1 Overview

Insect crop pests are a major global concern that exacerbate increasing pressures

on food supplies from overpopulation and global warming. Unfortunately, use of

chemical pesticides cause collateral environmental damage and kill non-target insects.139

Transgenic strategies such as Bt toxin can alleviate these concerns,140 however resistance

can emerge which limits their effectiveness.141–143 An increasingly exciting option for

control of plant pests is the use of RNA interference- (RNAi-) based technologies.5,6

RNAi is a process in which small, 19-30 nucleotide RNA molecules trigger the

destruction or decay of complementary transcripts.1 RNAi-based pest control improves

upon traditional small molecule pesticides by providing high specificity to the target

species.144

RNAi in insects can be induced through introduction of double stranded RNA

(dsRNA), which is processed into small interfering RNA (siRNA) effectors.145–147

Feeding of dsRNA to crop pests is effective in some species.4 Indeed, transgenic corn

expressing dsRNA is currently being used to control western corn rootworm (WCR) by

targeting vacuolar ATPase (V-ATPase).148 dsRNAs can also be applied as crop sprays,149

which enables use of synthetics to increase efficiency. Use of dsRNA in sprays is a very

attractive mode of delivery as it eliminates the need for transgenics, which are not

feasible to generate for some crops.

Unfortunately, while attempts at RNAi-based pest control have been successful in

some species, many insect orders seem refractory to ingested RNAi.150–153 Although

70

feeding is ineffective in these insects, dsRNA injection is often capable of eliciting

RNAi,150–155 indicating that barriers to dsRNA uptake primarily exist in the digestive

tract. Indeed, high nuclease activity in the migratory locust gut renders dsRNA feeding

ineffective.150 Furthermore, additional barriers may exist, such as the endosomal

entrapment of dsRNA found in lepidopterans (i.e. moths and butterflies).156 To address

this problem we sought to develop a polymeric dsRNA vector that can circumvent

barriers to uptake via ingestion and facilitate the use of RNAi in crop sprays.

Polycations have gained interest for their ability to electrostatically complex the

negatively-charged RNA phosphodiester backbone to form interpolyelectrolyte

complexes (IPECs).13,60 Recently, we demonstrated that polymers synthesized from N-(3-

guanidinopropyl)methacrylamide (GPMA) are able to enter cells readily via both

endocytotic and nonendocytotic routes,64 and Tabujew et al. subsequently demonstrated

these polymers can bind and protect siRNAs.157 pGPMA guanidinium groups provide

moieties similar to arginine-rich cell penetrating peptides (CPPs), which are observed to

accumulate in endomembrane vesicles, where they can cross membranes.158–160 CPPs

have also been found to enter cells through nonendocytotic routes.63 In Sf9 cells, an

RNAi-insensitive cell line derived from fall armyworms (Spodoptera frugiperda), naked

dsRNAs are eliminated in endosomal compartments.156 In this study, we test the ability of

pGPMA to enable RNAi in fall armyworms, through cytoplasmic delivery of dsRNAs.

We find that pGPMA-dsRNA IPECs can elicit RNAi in fall armyworm cells and

larvae that are otherwise insensitive to ingested RNAi.156 Confocal microscopy revealed

successful dsRNA delivery to Sf9 cells, and RT-qPCR analysis showed potent gene

suppression. These IPECs, when fed to fall armyworm larvae, resulted in a similar degree

71

of knockdown. Through targeting a gene known to have a role in digestive physiology,

IPECs induced larval mortality and significant gut hypertrophy. To our knowledge, this is

the first study demonstrating successful gene suppression via orally ingested RNAi in an

otherwise insensitive lepidopteran species.

4.2.2 pGPMA synthesis and IPEC characterization

Employing aqueous reversible addition-fragmentation chain transfer (aRAFT)

polymerization, poly[N-(3-guanidinopropyl)methacrylamide] (pGPMA, Figure 4.8a,b)

was synthesized to serve as a dsRNA delivery vehicle. This cationic polymer shares

features with other synthetic carriers of nucleic acids that can electrostatically bind to

negatively-charged RNA phosphodiester groups to form interpolyelectrolyte complexes

(IPECs).13,60 Formation of IPECs confers enhanced RNA stability, providing protection

from RNase-mediated degradation.43,44,46 pGPMA is chemically similar to CPPs, which

can traverse biological membranes and enter cells through both endocytotic and

nonendocytotic pathways,63 and we have previously shown similar behavior for

pGPMA.64 It has been demonstrated that pGPMA has a modest capacity to deliver

plasmid DNAs to nuclei, though toxicity was observe in one cell line.161 However,

studies in other cell culture systems have demonstrated negligible toxicity for similar

polymers,65,68,162,163 suggesting that any toxicity may be confined to certain cell types or

configuration of IPECs. Use of pGPMA-dsRNA IPECs also addresses the alkalinity and

high RNase activity in insect guts: lumen pH ranges from 10-11,8 where commonly used

tertiary amine-containing nucleic acid carriers become deprotonated, leading to IPEC

dissociation. However, the guanidinium functionalities of pGPMA, with pKa = 12.5,

should retain cationic charges in the same pH range. Additionally, GPMA-based

72

polymers can form multiple hydrogen bonds with the dsRNA phosphodiester moieties to

provide greatly enhanced binding to siRNAs,157 which subsequently should increase

protection from enzymatic degradation of dsRNA within the insect gut.

In order to form IPECs with a several hundred nucleotide dsRNA, a pGPMA

homopolymer with a degree of polymerization (DP) = ~100 repeat units was synthesized

using a protocol previously developed in our laboratories.64 Because the GPMA

guanidinium moieties serve as sites for both dsRNA complexation and cell penetration,

polymer-dsRNA IPECS must be formed at IPEC charge ratio (±) > 1 (i.e. net cationic

charge) to ensure solubility as well as uncomplexed GPMA units to interact with cell

membranes. Gel electrophoresis was performed to determine the polymer/dsRNA weight

ratio(s) at which ± > 1 (Figure 1c). On the basis of moderate IPEC gel migration toward

the anode, subsequent experiments used weight ratios of 4×, 6×, or 8× (± = 7, 11, 15,

respectively). At these ratios, the IPECs possess the desired net cationic charges while

not being so cationic that they encourage excessive protein opsonization and IPEC

exchange in vivo.

73

Figure 4.8 (a) Structure, number average molecular weight (Mn), and dispersity (Ð) of

pGPMA. (b) SEC trace of pGPMA (c) Gel electrophoresis of pGPMA-dsRNA IPECs.

Numbers indicate polymer/dsRNA weight ratio. (d) Proposed morphological changes in

IPEC structure between pH = 7.4 and pH = 10.

74

Table 4.3

ζ-potential measurements of pGPMA-dsRNA complexes at varying weight ratios and pH

Weight

ratio

ζ-potential (mV)

pH = 7.4

ζ-potential (mV)

pH = 10

4× 16.0 12.0

6× 18.4 12.8

8× 19.9 13.8

dsRNA -11.8 * *ζ-potential was not measured due to dsRNA hydrolytic instability at pH = 10.

Table 4.4

Dynamic and static light scattering measurements of pGPMA-dsRNA complexes at

varying pH (weight ratio = 8)

Sample Rh (nm) Rg (nm) Rg/Rh

IPEC, pH = 7.4 318.1 341.7 1.07

IPEC, pH = 10 239.2 436.5 1.82

dsRNA 35.7 71.8 2.01

ζ-potential measurements were also performed using these ratios to confirm net

cationic charges, both at neutral pH and under the alkaline conditions found in the

lepidopteran gut. As demonstrated in Table 4.3, ζ-potential values increase with

increasing polymer-dsRNA weight ratio at both pH conditions. As one might expect for a

system approaching the GPMA repeat unit pKa, some deprotonation likely occurs at pH =

10, leading to overall lower ζ-potential values relative to pH = 7.4 However, the positive

ζ-potential values for each IPEC indicate that a net cationic charge is maintained, even

under alkaline conditions.

A similar trend was revealed in dynamic light scattering (DLS) analysis of IPECs

formed at a weight ratio of 8 (Table 4.4). IPECs formed at pH = 7.4 resulted in a uniform

population (see Figure A.8 for histograms) with hydrodynamic radius Rh = 318.1 nm, but

75

at pH = 10, some pGPMA deprotonation leads to a partial collapse of the IPEC corona,

resulting in Rh = 239.2 nm. Additionally, the large increases in Rh for the IPEC vs. naked

dsRNA may suggest multiple dsRNAs per complex. Static light scattering (SLS) analysis

was performed to determine IPEC and dsRNA radii of gyration (Rg) and thus Rg/Rh,

which serves as an indicator of morphology. dsRNA alone exhibits Rg/Rh = 2.01,

indicating the rigid rod-like morphology expected of long dsRNA. The Rg/Rh values for

the IPECs at both high and low pH also indicate high aspect ratio morphologies as one

would expect from pGPMA chains binding to and forming a corona around a rigid rod.

The increase in Rg/Rh at pH = 10 further supports this notion: as the pGPMA corona

slightly collapses upon partial deprotonation, the IPEC increasingly adopts the

morphology of the dsRNA (Figure 4.8d).

4.2.3 pGPMA-dsRNA IPEC transfection and gene suppression in lepidopteran cell

culture

The IPECs were tested for their ability to enter Sf9 cells and affect gene

expression. This cell line is derived from embryonic fall armyworms and, unlike some

insect lines (e.g., Drosophila S2), is insensitive to dsRNA added to growth media.156 To

verify the ability of pGPMA to facilitate uptake of dsRNA, Cy5-labeled dsRNA was

complexed with pGPMA (8×) and added to Sf9 cell culture media. Cells were imaged

following incubation with the complex for 24 and 48 h. Significant accumulation of the

Cy5 signal could be observed in the pGPMA-dsRNA complex-treated cells after both 24

(Figure 4.9a) and 48 h. (Figure 4.9b). Conversely, cells treated with Cy5-dsRNA alone

(Figure A.9) exhibited no Cy5 signal. Accumulation appears constant, likely due to

continued uptake from media. Primarily the dsRNA localized to cellular bodies that are

76

likely endosomal, consistent with observations that guanidinium-functionalized

oligomers facilitate uptake of nucleic acids through an endocytosis-dependent

mechanism.162 Significantly, treatment with the polymers resulted in negligible

cytotoxicity (Figure 4.9c).

Figure 4.9 Sf9 cells treated with Cy5-labeled dsRNA (red) complexed with pGPMA after

(a, top row) 24 h or (b, bottom row) 48 h. Nuclei were stained with DAPI (blue). Scale

bars = 5 μm. (c) Cell viability assay of pGPMA after 48 h employing polymer

concentrations identical to the indicated weight ratios used in IPECs. Cell viability was

determined relative to the untreated control. Error bars represent the standard deviation

from triplicate experiments.

The CDC27 gene, which was targeted by RNAi in Sf9 cells in a previous study164

that relied on Caenorhabditis elegans SID-1 to transport dsRNA into the cytoplasm, was

used to test the ability of pGPMA to enable gene knockdown. pGPMA was complexed

either with CDC27-dsRNA or control dsRNA and added to Sf9 media. After a 48-h

incubation, expression levels were quantitated by RT-qPCR (Figure 4.10a). We observed

extensive knockdown of CDC27 (> 90%) that was sequence dependent. Time-dependent

gene suppression at an 8× weight ratio was then evaluated relative to untreated cells and

those transfected using Lipofectamine 3000 (Figure 4.10b). pGPMA-dsRNA IPECs

77

induced knockdown comparable to Lipofectamine and showed better performance after

72 h. To ensure that changes in gene expression were not induced by the polymer itself,

CDC27 expression was evaluated after treatment with uncomplexed pGPMA equivalent

to that of 8× weight ratio. No gene suppression from the polymer alone was observed

(Figure A.10).

The amount of dsRNA delivered at an 8× weight ratio was quantified via RT-

qPCR employing primers specific to the dsRNA, rather than the targeted mRNA (Figure

4.10c). After 24 h, pGPMA transfected similar amounts of dsRNA to Lipofectamine.

However, at 48 and 72 h, cells treated with IPECs maintained significantly higher levels

of transfected dsRNA than did those treated with Lipofectamine. The relatively high

levels of dsRNA transfected by pGPMA resulted in consistent levels of gene suppression

over 3 days. Lipofectamine, on the other hand, yielded decreasing levels of transfected

dsRNA over the observed time period that correspond to a trend of decreasing

knockdown. These results suggest that the IPEC does result in greater dsRNA protection

and retention within the cells, traits that would be advantageous when delivering dsRNA

through feeding.

78

Figure 4.10 (a) Expression of CDC27 determined by RT-qPCR in Sf9 cells following

incubation with pGPMA complexed with either CDC27- or control-dsRNA. Numbers

indicate polymer/dsRNA weight ratios. Values are normalized to CDC27 expression in

respective control (KIF-dsRNA-treated) samples. Errors bars represent SEM. (b)

Expression of CDC27 determined by RT-qPCR in Sf9 cells following incubation with

CDC27 dsRNA complexed with either pGPMA (8x) or Lipofectamine 3000. Values are

normalized relative to respective untreated controls. Error bars represent SEM. (c) RT-

qPCR quantification of CDC27-dsRNA transfected by pGPMA, Lipofectamine 3000, or

untreated control. Values are relative to zero. Error bars represent SEM. For plots (a-c),

groupings indicated with asterisks (*) were found to be significantly different after Tukey

analysis.

4.2.4 pGPMA-dsRNA IPEC gene suppression in lepidopteran larvae after oral

ingestion

Having demonstrated that pGPMA-dsRNA IPECs successfully elicit gene

knockdown in an otherwise refractory cell line, we evaluated their ability to trigger RNAi

in live caterpillars through feeding. RNAi has been used to target WCR worm V-ATPase

through feeding.4 Thus, we sought to similarly target a fall armyworm V-ATPase

ortholog (sfV-ATPase) using pGPMA. Larvae were fed pGPMA-dsRNA IPECs targeting

either sfV-ATPase or Green Fluorescent Protein (GFP, control dsRNA). 100 ng of

dsRNAs were fed to 2nd or 3rd instar larvae in complex with 8× pGPMA (w/w). Seven

days after feeding, total RNA was extracted from mid-guts, and RT-qPCR was performed

to determine changes in sfV-ATPase expression (Figure 4.11a). As in the cell culture

79

experiments, dsRNA delivered by pGPMA resulted in > 80% knockdown of the target

gene, indicating that pGPMA-dsRNA IPECs can successfully navigate the hostile

environment of lepidopteran guts, resulting in gene suppression after feeding.

Because suppression of sfV-ATPase leads to decreased nutrient uptake,4 such

extensive knockdown was expected to result in large increases in larval mortality.

However, only moderate larval death (Figure 4.11b), was observed after 29 days. Such

low mortality suggests that the inhibition of gene expression by RNAi is transient, or that

knockdown of a different gene may prove more effective. This could be addressed with

multiple doses of the IPEC, similar to what would be ingested through continuous

feeding on sprayed foliage. In any case, larval mortality was associated with the

significant gut hypertrophy expected from decreased nutrient uptake (Figure 4.11d), as

would be expected from sfV-ATPase knockdown. Additionally, when larvae were fed

pGPMA alone, no death was observed, even when fed 100x the amount used in the IPEC

feeding experiments (Figure A.11). These results, along with those of the Sf9 viability

assay, suggest low pGPMA toxicity, a necessary requirement for full implementation into

crop sprays.

80

Figure 4.11 (a) Expression of V-ATPase mRNA in midgut tissue from 2nd instar fall

armyworm larvae fed with pGPMA complexed with either V-ATPase dsRNA or GFP

dsRNA determined by RT-qPCR. Letters indicate individual animals. Days between

feeding and harvesting are indicated in parentheses. Values are normalized to V-ATPase

expression in control sample. Errors bars represent SEM. (b) Percent survival of 2nd and

3rd fall armyworm larvae fed pGPMA complexed with dsRNA targeting V-ATPase (N =

25) or control dsRNA (N = 31). (c) Image of fall armyworm larval gut after feeding with

pGPMA complexed with dsRNA targeting GFP or (d) sfV-ATPase. Scale bars = 2 mm.

81

CHAPTER V – CONCLUSIONS




The aRAFT polymerization of hydrophilic-block-cationic copolymers with

varying cationic block charge densities and their subsequent complexation with siRNA

and siRNA analogs has been demonstrated. Reduced charge density in these BICs

resulted in lower oligonucleotide stabilization and binding strength, characteristics that

predict more rapid siRNA release and thus enhanced gene suppression. However,

decreased cellular transfection and RNAi activation, which resulted in decreased gene

knockdown, indicate that decreased binding strength afforded by reduced charge density

promotes greater susceptibility to enzymatic degradation. Indeed the higher rate of in

vitro RNase degradation with decreasing charge density supports this notion.

Components of the RNAi pathway likely have a higher affinity for siRNA than do other

RNA-binding proteins (e.g. RNases). Thus, they likely are able to extricate siRNAs from

higher charge density copolymers, whereas less specific RNases cannot.

These results indicate that for hydrophilic-block-cationic copolymers with

relatively few charges (i.e. ~20 DMAPMA units), siRNA delivery is most effective

utilizing a fully charged cationic block without non-ionic comonomer. However, it is

worth noting that decreasing cationic block charge density diminishes copolymer

cytotoxicity (Figure A.6). Thus, application of variable charge density to block

copolymers with a greater number of DMAPMA units should improve polymer

biocompatibility while providing sufficient number of cations to maintain siRNA

82

protection. The effect of cationic block charge density in copolymers with greater

cationic content is the subject of ongoing investigation.



aRAFT polymerization of pGPMA and complexation with dsRNA has been

described. pGPMA successfully delivered dsRNA, targeting genes in a sequence specific

manner in otherwise refractory in Sf9 cells. Feeding pGPMA-dsRNA IPECs to fall

armyworm larvae likewise caused suppression of target mRNA accumulation, resulting

in moderate animal mortality. Furthermore, pGPMA alone seems to be relatively non-

toxic to the larvae and exhibited no significant toxicity in Sf9 culture. As previously

mentioned, pGPMA has exhibited cytotoxicity towards one cell line,161 but similar

guanidinium-functionalized polymers have exhibited negligible cytotoxicity in a myriad

other cell lines.65,68,162,163 To account for this variance, extensive toxicology studies

across multiple cell lines will be necessary before implementation into a commercial

product.

This is the first time to our knowledge that pGPMA-based polymers have been

shown to elicit RNAi in lepidopterans after oral ingestion, a strategy that has heretofore

been unsuccessful. The species specificity of RNAi makes this approach attractive from

an environmental perspective, and insect inability to develop resistance points to long-

term efficiency of this strategy. Furthermore, aRAFT polymerization provides access to

higher order polymer architectures with tailorable functionalities while maintaining

precise control. Thus, RNAi-based pesticides built on this IPEC platform could be

candidates for commercial development into crop sprays. Dosing optimization, toxicity

83

studies in animal models, and alterations to the polymer architecture for spray

formulation will be necessary to progress this technology and are the subjects of ongoing

investigation in our laboratories.

84

APPENDIX A – SUPPLEMENTARY INFORMATION

A.1 Block ionomer complexes consisting of siRNA and aRAFT-synthesized



A.1.1 Discussion of experiments pertaining to control copolymer PO

Mixing siRNA with non-ionic control polymer P0, which has no DMAPMA

content, resulted in solutions exhibiting insufficient excess scattering intensity during

DLS measurement to determine Rh (Table 2), indicating BICs were not formed. This

phenomenon, combined with the highly negative ζ-potential near that of free siRNA,

demonstrates the requirement of DMAPMA for BIC formation with these polymers.

Furthermore, these results confirm that the remaining folic acid-free APMA units,

protonated under physiological conditions, are unable to form stable electrostatic

complexes with siRNA. These conclusions are corroborated by the DSC (Figure 2) and

potentiometric titration experiments (Figure 3): P0-dsDNA exhibited a Tm (57.8 °C) near

that of free dsDNA (Tm = 54.4 °C), and P0-PSS near-zero ΔGtotal (-0.36 kJ/mol) as

expected from a non-complexing copolymer. Interestingly, Bioanalyzer quantification

revealed only 65.3 % free siRNA relative to the control. However, this apparent decrease

in uncomplexed siRNA is likely the result of hydrogen bonding or other non-ionic

copolymer-siRNA interaction.

85

A.1.2 Supplementary Figures

Figure A.1 1H NMR spectra of P100, P75, P50, P25, P0, and macroCTA

86

Figure A.2 UV-Vis spectroscopy of conjugated folic acid P100, P75, P50, P25, and P0

copolymers.

87

Figure A.3 Agilent 2100 Bioanalyzer electropherograms for copolymer-siRNA

complexes, free siRNA, and ladder. Uncomplexed siRNA concentration determined from

area of peak at ~39 s. Peak at ~35 s corresponds to 4-nucleotide marker.

88

Figure A.4 . Potentiometric titration curves for (A) P100, (B) P75, (C) P50, (D) P25, (E)

P0, and (F) P2 and their respective block ionomer complexes with polystyrene sulfonate

(PSS).

89

Figure A.5 Confocal fluorescence microscopy images of KB cells treated with

AlexaFluor594-tagged siRNA (red) delivered via (A) P100, (B) P75, (C) P50, (D) P25,

and (E) P0. Nuclei were stained with DAPI (blue). Scale bars = 10 μm.

90

Figure A.6 Cell viability assays of P100, P75, P50, P25, and P0 after 48 and 72 hours.

The cell viability was determined relative to KB cells. Error bars represent the standard

deviation from triplicate experiments.

91

A.2 Guanidinium-functionalized Polymer Carriers Enable RNAi in Insensitive Crop

Pest Arthropods

A.2.1 Primers Used in this Study

dsRNA Synthesis

SfV-ATPase_dsRNA F GAGGCTCTTCGTGAGATCTCAGG

SfV-ATPase_dsRNA R GAAACGATCGTATGACGAGTAGCTG

SfCDC27_dsRNA F ATTGTTCAAGAACCTATACAGGTTATCGTTTG

SfCDC27_dsRNA R CAGGAGCTTGAGTCTCTGGTGTGATGCTGG

M13 F TGTAAAACGACGGCCAGT

Sp6-T7 R TAATACGACTCACTATAGGGAGCTCTCCCATATGGTCGAC

RT-qPCR

SfV-ATPase_qPCR F TGTCCGTTCTACAAGACCGTGG

SfV-ATPase_qPCR R TCACGGATGACGTTCCAGGTG

dsCDC27 F CCACCAAGATGATTGTTCAAG

dsCDC27 R GAGTCTCTGGTGTGATGCTGG

SfKIF23 F AAGGAACTGATGGCACATTTGGAAATGAGG

SfKIF23 R AGTGGCGGTCAAGCGTTCTTCCAGAGCTCT

SfActin_qPCR F AGATGACACAGATCATGTTCG

SfActin_qPCR R GAGATCCACATCTGTTGGAAG

GFP_qPCR F TGAAGTTCATCTGCACCACCGG

GFP_qPCR R TTGAAGAAGTCGTGCTGGCG

92

A.2.2 Supplementary Figures

Figure A.7 1H NMR spectra of pGPMA polymerization aliquots taken at (a) t0 and (b) 19

hr.

93

Figure A.8 Hydrodynamic size histograms for pGPMA-dsRNA IPEC (8x weight ratio) at

pH = 7.4 (black) and pH = 10 (red) and free dsRNA (pH = 7.4, green) as determined by

dynamic light scattering (DLS) at 90° scattering angle.

94

Figure A.9 Sf9 cells treated with free Cy5-labeled dsRNA (red) after (a, top row) 24 hrs

and (b, bottom row) 48 hrs. Nuclei were stained with DAPI (blue). Scale bars = 5 μm.

Figure A.10 Expression of CDC27 determined by RT-qPCR in Sf9 cells following

incubation with free pGPMA at identical concentration as used for 8x IPEC. Values are

normalized to respective untreated controls. Error bars represent SEM.

95

Figure A.11 Survival of fall armyworm larvae after ingestion of pGPMA. Animals were

directly fed masses indicated on the left y-axis (black line). The percentage of animals

viable after feeding on right y-axis (grey line). (N = 4)

96

APPENDIX B – INFLUENCE OF Z-GROUP HYDROPHOBICITY IN VISIBLE LIGHT-

MEDIATED AQUEOUS RAFT POLYMERIZATION

B.1 Overview

In recent years, considerable efforts have been made into the development of

external control over reversible deactivation radical polymerization (RDRP) techniques,

most notably toward the use of light for its spatiotemporal control, ease of use, and

inexpensive sources.117 Of the available RDRP techniques, reversible addition-

fragmentation chain transfer (RAFT) polymerization has grown into one of the most

popular as a result of its excellent polymerization control while tolerating a wide variety

of monomers and functional groups under relatively mild reaction conditions. Successful

control of a RAFT polymerization is maintained through the use of a thiocarbonylthio

chain transfer agent (CTA) which undergoes rapid degenerative chain transfer with the

propagating radical species.

Although the majority of RAFT polymerizations have relied upon the thermal

decomposition of an exogenous initiator to generate the steady-state radical

concentration, it has been established that UV and γ irradiation can excite the C=S

moiety of the CTA, leading to the fragmentation of the adjacent C—S bond to generate a

thiyl radical and a carbon-centered radical.114,165 In addition to recombination with the

thiyl radical to regenerate the CTA, the carbon-centered radical can undergo addition to

monomer or can participate in degenerative chain transfer with a non-fragmented CTA to

participate in the usual RAFT process (Scheme B1). This initiator- and catalyst-free

iniferter/RAFT process under UV light has been successfully used by a number of

research groups in the synthesis of a number of well-defined polymers and polymer

97

architectures.165–171 However, tendency of many organic compounds to absorb UV light

increases the probability of unwanted side reactions. Furthermore, CTAs have been

shown to irreversibly degrade under UV light, especially at higher monomer

conversions.165,167,169,171,172

Scheme B.1 Mechanism of light-mediated RAFT polymerization

More recently, however, both Qiao117 and Boyer116 have demonstrated that a

number of CTAs, most notably trithiocarbonates (TTCs), will fragment also under visible

light, resulting in iniferter/RAFT-type polymerization. Their groups, among others, have

demonstrated the successful visible light-mediated RAFT polymerization of a number of

monomers, predominantly acrylate-/acrylamide- and methacrylate-type, in a variety of

solvents under extremely mild conditions, including room temperature, while maintaining

excellent chain-end fidelity. Indeed, given the appropriate polymerization conditions,

high monomer conversions can be rapidly achieved while maintaining temporal, “on-off”

control of the reaction via switching the light source on and off. Furthermore, this

technique has also been demonstrated to be amenable to “one-pot” type chain extensions,

allowing for facile synthesis of higher order polymer architectures, including multi-block

copolymers.

98

Our research group has maintained a long-standing interest the development of

aqueous RAFT (aRAFT) polymerization to synthesize well-defined, advanced polymer

architectures. aRAFT is highly advantageous not only for its environmental friendliness

as a result of polymerization directly in water, but also because aqueous conditions make

it highly suitable for biomedical applications. The application of visible light mediation

to aRAFT could augment these advantages by imparting low-temperature reaction

conditions under visible light that is more amenable to biological conditions.

Furthermore, the exclusion of exogenous thermal initiators from the reaction mixture

decreases the requirement of extensive product purification. Although catalyst- and

initiator-free photo-mediated aRAFT using UV irradiation as the light source has been

documented,166 such polymerization in the presence of visible light has only been

investigated for water-solvent mixtures.117

In considering an appropriate CTA for visible light-mediated aRAFT

polymerization, we reasoned that, in addition to the usual considerations in selecting R-

and Z-groups with regard to reactivity toward propagating radical species and subsequent

radical fragmentation, the hydrophilicity of both the R- and Z-groups must also be taken

into consideration. A wide variety of water-soluble TTCs are available commercially and

synthetically, and while some of them incorporate hydrophilic Z-groups, the most

popular throughout the literature consist of an S-dodecyl or -ethyl Z group, relying solely

on hydrophilic moieties in the R-group to maintain water solubility. In thermally-initiated

aRAFT polymerization using a water-soluble initiator, such hydrophobic Z-groups do not

significantly affect the water solubility of the TTC because the degenerative chain

transfer mechanism results in a hydrophilic R-group always being present on the RAFT

99

agent. The significance of this hydrophobicity is further diminished as hydrophilic

monomer is incorporated into the propagating polymer chain. However, in visible light-

mediated aRAFT polymerization, photolysis of the initial TTC results in the generation

of a thiyl radical that should be minimally soluble with an alkyl Z-group. Furthermore, it

has been established that under such conditions, two TTC thyil radicals can couple to

form a bis-TTC disulfide.116 Such a byproduct of TTCs with alkyl Z-groups would

certainly be hydrophobic and should precipitate from an aqueous solution, causing loss of

polymerization control. Herein, we investigate the conditions under which such

byproduct formation and precipitation occurs and how it affects visible light-mediated

aRAFT polymerization.

B.2 Experimental

B.2.1 Materials

All reagents were used as received unless otherwise noted. 4-((((2-

carboxyethyl)thio)carbonothioyl)thio)-4-cyanopentanoic acid (CETPA) was purchased

from Boron Molecular. Sodium ethyltrithiocarbonate, bis(ethylsulfanylthiocarbonyl)

disulfide, and 2-(ethylthiocarbonothiolthio)-2-(2-imidazolin-2-yl)propane hydrochloride

(ImET) were synthesized as previously reported.173 3-

(methacryloylamino)propyltrimethylammonium chloride (MAPTAC) was purchased

from Wako. N-(2-hydroxypropyl)methacrylamide (HPMA) was synthesized as

previously reported.118

B.2.2 Light Source

All polymerizations were carried out in a home-built photo-reactor consisting of a

multicolor LED flexible tape strip (Commercial Electric, Model #16508) wrapped

100

around the inside of a large recrystallization dish that sat on a magnetic stir plate. The

outside was wrapped in aluminum foil to ensure maximum reflectivity, and a cardboard

top was constructed with a hole in the center to ensure that all reaction vials were placed

in the exact center of the reactor. All polymerizations were carried out with the LED strip

set to blue (λ = 450 nm) at maximum brightness.

Figure B.1 Home-built photo-reactor used for visible light-mediated aRAFT

polymerizations

B.2.3 General Procedure for Blue Light-Mediated aRAFT Polymerizations

A typical reaction is as follows. MAPTAC (1.10g, 5.0 mmol) and ImET (19.1 mg,

6.7 x 10-5 mol) were added to an 8 mL vial with pierceable cap (Kimble Chase) equipped

with a stir bar. 2.0 mL 18.2 MΩ diH2O was added, and the solution was vortexed until all

solids were fully dissolved. The vial was placed in the home-built photo-reactor and was

purged with Ar for 40 min. in the dark, after which the mixture was then stirred in the

presence of blue light. The reaction was terminated by removal from the light source and

exposure to air. The final product was isolated by precipitation into acetone, followed by

redissolution in ethanol and precipitation into ether. To monitor polymerization kinetics,

aliquots were removed at regular time intervals via degassed syringe. Aliquots were

analyzed by 1H NMR (D2O) to determine monomer conversion and ASEC-MALLS to

101

determine molecular weights and dispersities. Monomer conversion was determined via

1H NMR by monitoring the disappearance of the MAPTAC vinyl peak (s, 1H, 5.54 ppm)

relative to the MAPTAC trimethyammonium and methylene peaks (13H: t, 4H, 3.20 ppm

and s, 9H, 2.95 ppm). For polymerizations of HPMA, monomer conversion was

determined via 1H NMR by monitoring the disappearance of the HPMA vinyl peak (s,

1H, 5.69 ppm) relative to the methine peak (m, 1H, 3.94 ppm).

B.2.4 Isolation of TTC Byproduct

ImET (32.3 mg, 1.13 x 10-4 mol) was added to an 8 mL vial with pierceable cap

(Kimble Chase) equipped with a stir bar and dissolved in 2.0 mL 18.2 MΩ H2O. The vial

was placed in the home-built photo-reactor and was purged with Ar for 40 min. in the

dark, after which the mixture was then stirred in the presence of blue light for 24 hr to

yield a cloudy yellow solution. The precipitate was extracted into diethyl ether (1 mL),

separated from the aqueous layer, and was isolated by rotary evaporation followed by

drying in vacuo to yield a yellow oil (3.0 mg).

B.2.5 Characterization

NMR spectra for structural analysis and monomer conversions were obtained

using a Varian INOVA 300 MHz NMR spectrometer. Polymer molecular weights and

dispersities (Ð) were determined by aqueous size exclusion chromatography (ASEC)

with an eluent of 1% (v/v) acetic acid and 0.2 M NaCl (aq) at a flow rate of 0.25 mL/min,

Eprogen Inc. CATSEC columns (100, 300, and 1000 Å) connected in series with a Wyatt

Optilab DSP interferometric refractometer (λ = 690 nm) and Wyatt DAWN DSP multi-

angle laser light scattering (MALLS) detector (λ = 633 nm). Absolute molecular weights

and Ð were calculated using a Wyatt ASTRA SEC/LS software package. Values of dn/dc

102

were determined offline utilizing a Wyatt Optilab DSP interferometric refractometer (λ =

690 nm) at 25 °C and Wyatt ASTRA DNDC software. FT-IR measurements were

performed with a Nicolet 6700 FTIR equipped with Smart iTR ATR accessory with

diamond crystal, a KBr beamsplitter, and a DTGS KBr detector. Samples in CHCl3 were

deposited directly onto the crystal and allowed to dry before measurement. FT-IR spectra

were recorded from 4000-650 cm-1 by averaging 32 scans with 4 cm-1 resolution.

B.3 Results and Discussion

B.3.1 Trithiocarbonate Byproduct

Figure B.2 Structures of monomers and RAFT agents used in this study.

To confirm that visible light-mediated aRAFT using a TTC with a hydrophobic

Z-group indeed results in byproduct precipitation, a preliminary polymerization of 3-

(methacryloylamino)propyltrimethylammonium chloride (MAPTAC) was performed

using 2-(ethylthiocarbonothiolthio)-2-(2-imidazolin-2-yl)propane hydrochloride (ImET)

as the CTA. [M]0:[CTA] = 75, and the reaction was allowed to proceed under blue light

irradiation overnight, resulting in a viscous, cloudy solution, indicating the formation of

precipitate. The most likely mechanism for precipitate formation is depicted in Scheme

B2. Upon irradiation, ImET photolyses to the TTC thyil radical and the 2-(2-imidazolin-

2-yl)propane hydrochloride R-group radical (R˙). In addition to the reverse reaction, R˙

103

behaves as would the initiating species in a thermally-initiated RAFT polymerization by

either undergoing degenerative chain transfer with another CTA or adding directly to

monomer before doing the same. Additional iniferter-type reactions can occur throughout

these processes. However, the ImET TTC thyil radical should be relative hydrophobic

and, after either coupling with another to form the bis-TTC or abstracting a hydrogen

from somewhere, should precipitate.

Scheme B.1 Proposed mechanism for byproduct precipitation during visible light-

mediated aRAFT polymerization of MAPTAC using ImET as CTA.

To verify the identity of the precipitate, ImET was allowed to react under blue

light in the absence of monomer. After 24 hr, the yellow precipitate was collected and

analyzed by 1H NMR and FT-IR spectroscopy. For comparative purposes, bis-TTC and

the trithiocarbonic acid were synthesized separately and also analyzed. The 1H NMR

spectra are depicted in Figure B3. Clear overlap of the resonances at 3.31 ppm and 1.36

ppm for the byproduct and bis-TTC suggest that bis-TTC is indeed the major byproduct.

104

The quartet appearing at 2.97 ppm is likely the result of disulfide formation between the

TTC and ethanethiol, which can spontaneously form along with CS2 from the

spontaneous decomposition of TTC. The FT-IR spectra are depicted in Figure B4.

Overlaying the spectra for both bis-TTC and ethyltrithiocarbonic acid on top of the

byproduct reveals overlap of every peak for the byproduct. Thus, at least from a

qualitative standpoint, the byproduct precipitate formed during polymerization is a

mixture of the bis-TTC and the trithiocarbonic acid, with the bis-TTC being the

predominant product.

Figure B.3 1H NMR spectra of ethyltrithiocarbonic acid, bisethyltrithiocarbonate, and

TTC byproduct.

105

Figure B.4 FT-IR spectra of ethyltrithiocarbonic acid, bisethyltrithiocarbonate, and TTC

byproduct.

B.3.2 Effect of Byproduct Precipitation on Polymerization

Bis-TTC precipitation indicates polymer end-group loss, and thus several possible

negative impacts are to be expected throughout the polymerization. If TTC loss occurs

prior to initiation, [M]0:[CTA] would increase, resulting than higher-than-expected

molecular weights. End-group loss throughout polymerization would reduce the number

of chain transfer and reversible end-capping species, lowering polymerization control and

increasing polymer dispersity. Finally, because photolysis of the TTC is the only radical

source during the polymerization, TTC loss should result in a decrease in the radical

concentration throughout the polymerization.

To establish which negative effects were taking place, the preliminary

polymerization was repeated, and aliquots were taken at regular intervals to monitor the

evolution of molecular weight and dispersity as a function of monomer conversion

(Figure B5a). As a control, a tandem polymerization was performed using 4-((((2-

W a v e n u m b e r (c m-1

)

Ab

so

rb

an

ce

1 0 0 02 0 0 03 0 0 0

0

5 0

1 0 0

B is e th y ltr ith io c a rb o n a te

E th y ltr ith io c a rb o n ic a c id

P re c ip ita te

106

carboxyethyl)thio)carbonothioyl)thio)-4-cyanopentanoic acid (CETPA), which contains a

hydrophilic Z-group, as the RAFT agent (Figure B5b). As expected, the polymerization

with ImET resulted in a cloudy solution, whereas CETPA resulted in a clear solution.

However, both polymerizations apparently exhibited none of the aforementioned negative

effects on polymerization: molecular weight evolved linearly as a function of monomer

conversion and closely matched the theoretical. While ImET did result in higher

dispersity than did CETPA, both reactions maintained Ð < 1.2, indicating good control.

Furthermore, the pseudo-first order kinetic plots were linear and exhibited no downward

curvature, which would be expected if radical concentration decreased.

Figure B.5 Mn vs. conversion plots, pseudo-first order kinetic plots, and photos of final

polymerization solutions for visible light-mediated aRAFT polymerization of MAPTAC

using (a) ImET and (b) CETPA as CTA. [M]0:[CTA] =75 for both reactions.

In addition to the aforementioned issues, TTC loss should also drastically affect

chain extension. Figure B6 depicts the ASEC traces for MAPTAC macroCTAs made

107

from ImET and CETPA as well as their respective chain extensions to form MAPTAC-b-

MAPTAC. Both macroCTAs result in a shift to lower elution volume, as would be

expected. However, whereas the CETPA resulted in minimal low molecular weight

tailing with Ð < 1.2, the ImET chain extension has a clear low molecular weight shoulder

that corresponds to the original macroCTA, and Ð = 1.36. Both of these are consistent

with extensive end-group loss and are what would be expected after TTC precipitation.

Figure B.6 ASEC traces for MAPTAC macroCTAs and their chain extensions to form

MAPTAC-b-MAPTAC using (a) ImET and (b) CETPA as CTA.

Although polymerizations employing ImET always resulted in precipitate

forming, to the naked eye it seemed that precipitation only occurred at > 90% monomer

conversion, which would be consistent with the apparent lack of effect on polymerization

kinetics. To quantify when precipitation occurrs, photoreactions of ImET and MAPTAC

as well as ImET alone were performed in a quartz cuvette, and the solution transmittance

at λ = 550 nm was monitored as a function of time (Figure B7). Whereas ImET alone

resulted in immediate precipitation, the transmittance for the polymerization of

MAPTAC did not decrease until > 300 min, corresponding to a monomer conversion of >

95%. Clearly polymerization prevents precipitation. A high rate of propagating chain

108

transfer to the byproduct or recombination with the TTC radical likely prevents

byproduct precipitation. However, such reactions require collision of a chain end with a

small molecule. As polymer length, and accordingly solution viscosity, increases, such

collision frequencies should decrease, reducing the rates of their respective reactions.

Because TTC-TTC coupling requires only the collision of two small molecules, its rate

should not be decreased at high DP and thus could be higher than reactions with the

polymer chain end, resulting in byproduct precipitation. If such a theory is true, then

deleterious polymerization effects should manifest once a given DP has been reached.

T im e (m in )

% T

ra

ns

mit

tan

ce %

Co

nv

ers

ion

0 2 0 0 4 0 0 6 0 0

2 0

4 0

6 0

8 0

1 0 0

0

2 0

4 0

6 0

8 0

1 0 0

Im E T + M A P T A C

Im E T

% C o n v e rs io n

Figure B.7 Percent transmittance and monomer conversion as a function of time for

reaction of ImET with MAPTAC and ImET alone in the presence of blue light.

In the aforementioned polymerization, the onset of byproduct precipitation

coincided with monomer conversion = 95%, corresponding to DP = 71. Therefore, we

hypothesized that upon targeting a higher theoretical DP, the polymerization would be

negatively affected after DP > 71. Unfortunately, unforeseen issues obtaining additional

monomer from the manufacturer prevented further investigation with MAPTAC. To

109

explore the effect of DP on polymerization kinetics, additional visible light-mediated

aRAFT polymerizations were performed using neutrally-charged N-(2-

hydroxypropyl)methacrylamide (HPMA). A series of polymerizations were conducted

using HPMA and ImET for which [M]0:[CTA] = 200, 400, 500, and 1000. The Mn vs.

conversion and pseudo-first order kinetic plots for [M]0:[CTA] = 200, 400, 500, and 1000

are depicted in Figure B8. In all cases, Mn closely matches the theoretical and Ð remains

< 1.2. Additionally, no downward curvature is present in any of the pseudo-first order

plots. These results suggest that the above hypothesis is incorrect: even for extremely

high DP, where the viscosity is substantially increased, the polymerization remains

unaffected. Thus, likely no TTC precipitation occurs throughout the polymerization.

110

Figure B.8 Mn vs. conversion plots and pseudo-first order kinetic plots for visible light-

mediated aRAFT polymerization of HPMA using ImET as CTA. [M]0:[CTA] = (a) 200,

(b) 400, (c) 500, (d) 1000.

111

Although extremely high DP does not appear to decrease the rate of propagating

chain transfer to the byproduct or recombination with the TTC radical, high rates of these

events must be what precludes byproduct precipitation. The only remaining conclusion is

that extended exposure to the light source after monomer depletion or loss of propagating

chain radicals via end-to-end coupling at high monomer conversion eventually allows for

byproduct formation to occur. Which of these events occurs likely depends upon the

monomer in question. There is no indication of end-to-end coupling in GPC traces of the

MAPTAC polymerizations (Figure B6), likely due to increased repulsion of the highly

cationic chains. Therefore, byproduct precipitation most likely occurs after monomer

exhaustion for this system. Alternatively, the GPC traces of some HPMA

polymerizations exhibit high molecular weight shoulders and low molecular weight

tailing at high conversion (Figure B9), indicative of end-to-end coupling and end-group

loss. Therefore, in such systems, loss of propagating chain radicals at high conversion

most likely results in byproduct formation and precipitation.

112

Figure B.9 ASEC-MALLS traces of visible light-mediated aRAFT polymerization of

HPMA using ImET as the RAFT agent targeting [M]0:[CTA] = 1000.

B.4 Conclusions

Although use of a TTC with a hydrophobic Z-group in visible light-mediated

aRAFT can result in CTA byproduct formation and precipitation at extremely high

monomer conversion, there appears to be no evidence that such an event occurs during

the earlier stages of the polymerization. The kinetic plots for a myriad of polymerization

conditions exhibited none of the deviations expected from TTC loss. Only chain

extensions after full monomer conversion seem to be negatively impacted when using a

hydrophobic Z-group. However, this can be mitigated through the use of a hydrophilic Z-

group or stopping macroCTA polymerization at lower conversions. These results speak to

the robustness of visible light-mediated aRAFT polymerization as a whole. As evidenced

in Figure B8d, molecular weights over 100 kDa can be achieved with excellent

polymerization control. It seems that one need not be concerned with Z-group

hydrophobicity as long as extreme monomer conversions are avoided. Furthermore, we

113

suspect that careful selection of the appropriate monomer, CTA, and polymerization

conditions may open an avenue to aRAFT polymerization without the need for product

purification. If full monomer conversion could be reached in pure water without TTC

byproduct formation, there would be none of the residual monomer, initiator, or buffer

typical of conventional aRAFT polymerization. All that would remain is the final product

in water, which could be lyophilized or used as is. Such an achievement would be greatly

desirable from both a “green” chemistry perspective as well as reduction of time required

for polymer synthesis.

114

APPENDIX C RIGHTS & PERMISSIONS

Material in Chapters 3.1, 4.1, and 5.1 reproduced from Ref.44 with permission from the

Royal Society of Chemistry.

Material in Chapters 3.2, 4,2, and 5,2 reprinted with permission from Parsons, K.H.;

Mondal, M.H.; McCormick, C.L.; Flynt, A.S. Biomacromolecules. 2018. DOI:

10.1021/acs.biomac.7b01717. Copyright 2018 American Chemical Society.

Figures 1.5 and 1.7 reprinted from Advanced Drug Delivery Reviews, Vol. 30, Kabanov,

A.V. and Kabanov, V. A., “Interpolyelectrolyte and block ionomer complexes for gene

delivery: physicochemical aspects,” 49-60, Copyright 1998, with permission from

Elsevier.

Figure 1.8 from Harada, A.; Kataoka, K. Science. 1999, 283 (5398), 65–67. Reprinted

with permission from AAAs.

Figure 1.9 reprinted with permission from Scales, C. W.; Huang, F.; Li, N.; Vasilieva, Y.

A.; Ray, J.; Convertine, A. J.; McCormick, C. L. Macromolecules 2006, 39 (20), 6871–

6881. Copyright 2006 American Chemical Society.

Figures 1.10 and 1.11 reprinted with permission from York, A. W.; Zhang, Y.; Holley, A.

C.; Guo, Y.; Huang, F.; McCormick, C. L. Biomacromolecules 2009, 10 (4), 936–943.

Copyright 2009 American Chemical Society.

Figure 1.12 and 1.13 reprinted with permission from Lu, H.; Wang, D.; Kazane, S.;

Javahishvili, T.; Tian, F.; Song, F.; Sellers, A.; Barnett, B.; Schultz, P. G. J. Am. Chem.

Soc. 2013, 135 (37), 13885–13891. Copyright 2013 American Chemical Society.

115

Figure 1.13 reprinted with permission from Treat, N. J.; Smith, D.; Teng, C.; Flores, J.

D.; Abel, B. A.; York, A. W.; Huang, F.; McCormick, C. L. ACS Macro Lett. 2012, 1 (1),

100–104. Copyright 2012 American Chemical Society.

Figure 1.15 reprinted with permission from Wiley. Copyright 1986 Hüthig & Wepf

Verlag.

Figures 1.16 and 1.17 reprinted with permission from Holley, A. C.; Ray, J. G.; Wan, W.;

Savin, D. A.; McCormick, C. L. Biomacromolecules 2013, 14, 3793–3799. Copyright

2013 American Chemical Society.

Figure 1.18 reprinted from J. Control. Release, Vol 133, Convertine, A. J., Benoit,

D.S.W., Duvall, C.L., Hoffman, A.S., Stayton, P.S., “Development of a novel

endosomolytic diblock copolymer for siRNA delivery,” 221-229, Copyright 2009, with

permission from Elsevier.

Figure 1.19 reprinted with permission from Lin, C.; Zhong, Z.; Lok, M. C.; Jiang, X.;

Hennink, W. E.; Feijen, J.; Engbersen, J. F. J. Bioconjug. Chem. 2007, 18 (1), 138–145.


Figure 1.20 reprinted with permission from Truong, N. P.; Jia, Z.; Burgess, M.; Payne,

L.; McMillan, N. A. J.; Monteiro, M. J. Biomacromolecules 2011, 12 (10), 3540–3548.


116

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