UNIVERSITY of CALIFORNIA Los Angeles Trehalose Glycopolymers for Protein and Cell Stabilization a Dissertation Submitted in Part

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UNIVERSITY OF CALIFORNIA

Los Angeles

Trehalose Glycopolymers for Protein and Cell Stabilization

A dissertation submitted in partial satisfaction of the

requirements for the degree Doctor of Philosophy

in Bioengineering

Uland Y Lau

2016

Uland Y Lau

2016

ABSTRACT OF THE DISSERTATION

Trehalose Glycopolymers for Protein and Cell Stabilization

Uland Y Lau

Doctor of Philosophy in Bioengineering

University of California, Los Angeles, 2016

Professor Heather D. Maynard, Chair

Proteins, biomolecules, and cells play an important role for their use as therapeutics and reagents for research applications. However, one main limitation is their inherent instability.

Proteins can easily denature and lose activity under a variety of stresses, and cells can easily lose viability due to cytotoxic compounds and conditions. This has led to the investigation of excipients that can aid in preserving activity, including polymers, osmolytes, proteins, and sugars. Among the sugars, trehalose has been shown to have remarkable properties for general protein and cell stabilization. This dissertation presents the utilization of trehalose glycopolymers for the stabilization of cells and proteins in three different applications. The polymers were used for protein stabilization during protein micro- and nano-patterning, stabilization of a therapeutic- relevant protein, and for cell preservation.

Direct write patterning of multiple biological molecules on surfaces has tremendous potential in applications of tissue engineering, diagnostics, proteomics and biosensors. Precise

ii control over the position and arrangement of proteins, especially on the micro- and nanoscale level is a particular challenge towards the improvement of miniaturized devices. A trehalose glycopolymer was utilized as a resist material that protects proteins during electron beam lithography, and thus enables direct write multiplexed protein patterns with micro- and nanoscale features and alignment capabilities (Chapter 2). The trehalose glycopolymers behave as negative resists, crosslinking to the substrate upon electron beam irradiation, and provide stabilization to proteins against the harsh processing conditions. Patterns with user-defined shapes were generated with a variety of protein types, such as antibodies, enzymes, and growth factors, and were shown to retain their activity. To further demonstrate the technique, fabrication of antibody patterns for multiplexed cytokine detection from live cells was achieved (Chapter 3).

A sandwich immunoassay was developed for the detection of interleukin-6 (IL-6) and tumor necrosis factor alpha (TNFα) secreted directly from stimulated macrophage cells. Multiplexing with both IL-6 and TNFα on a single chip was demonstrated successfully with high specificity and in relevant cell culture conditions. The ability to monitor cytokine release over time following cell stimulation was also demonstrated. Simultaneous detection of these extracellular signaling markers demonstrates the potential application towards disease profiling and diagnostics.

Next, the trehalose glycopolymers were used for the stabilization and enhancement of the pharmacokinetic properties of a therapeutic protein, granulocyte colony-stimulating factor (G-

CSF) (Chapter 4). G-CSF was designed with a polyhistidine-tag, maltose binding protein for solubility, and an enzyme cleavage site, and was expressed in E. coli and purified for the studies.

Trehalose polymers with polystyrene or polymethacrylate backbones (P1, P2, and P3), as well as a degradable trehalose polymer (p(BMDO-co-trehalose)) were synthesized to investigate their

iii potential for therapeutic use. The trehalose polymers were shown to be non-cytotoxic in mouse and human cell lines up to a concentration of 8 mg/mL. However, the degraded products of p(BMDO-co-trehalose) exhibited cytotoxicity at a lower concentration of 1 mg/mL. Stability screening with the trehalose glycopolymers on G-CSF showed that styrenyl acetal linked trehalose polymer, P1, best stabilized G-CSF compared to the other polymers and was selected for G-CSF conjugation. Therefore, P1 with a benzaldehyde end group was synthesized and conjugated to G-CSF via reductive amination. The resulting G-CSF-P1 conjugate retained higher bioactivity compared to native G-CSF when subjected to heat and lyophilization stressors.

Finally, the trehalose glycopolymers were investigated for the stabilization and preservation of cells (Chapter 5). Three different approaches were evaluated. First, extracellular protection by the covalent attachment of trehalose polymers to cells using modified sugars was tested. To evaluate the effectiveness of extracellular trehalose on cell stabilization, the trehalose polymers were added as excipients. Jurkat cells were heated at 50 °C and the added trehalose polymer did not significantly stabilize the cells at longer stress times. Cryopreservation with a combination of the polymer and dimethylsulfoxide (DMSO) showed slight increase in cell viabilities (65.7% ± 3.1 with polymer and DMSO compared to 55.6% ± 2.1 with DMSO only), but the results suggest that trehalose polymers are not likely to fully replace DMSO as a cryopreservative agent. The final approach utilized a bacterial pore-forming toxin, streptolysin O

(SLO), to permeabilize human dermal fibroblast (HDF) cells for loading trehalose polymers.

Permeabilization and loading was characterized by fluorescence with a fluorophore-labeled trehalose polymer and by the anthrone assay. No significant improvements in cell preservation were observed through trehalose polymer loading by this approach. Other techniques for cytosolic uptake of trehalose polymers should be considered for future work.

The dissertation of Uland Y Lau is approved.

Yong Chen

Andrea M. Kasko

April Dawn Pyle

Heather D. Maynard, Committee Chair

University of California, Los Angeles

2016

TABLE OF CONTENTS

ABSTRACT OF THE DISSERTATION ...... ii

TABLE OF CONTENTS ...... vi

LIST OF FIGURES, SCHEMES & TABLES ...... viii

LIST OF ABBREVIATIONS ...... xii

ACKNOWLEDGEMENTS ...... xv

CHAPTER 1. Introduction and Literature Review ...... 1

1.1 Overview of Trehalose ...... 2

1.2 Protein Therapeutics and Challenges ...... 3

1.3 Protein Patterning...... 10

1.4 Cell Preservation ...... 15

1.5 References ...... 19

CHAPTER 2. Trehalose Glycopolymer Resists Allow Direct Writing of Protein Patterns by Electron Beam Lithography ...... 31

2.1 Introduction ...... 32

2.2 Results and Discussion ...... 35

2.3 Conclusions ...... 43

2.4 Materials and Methods ...... 44

2.5 References ...... 48

CHAPTER 3. Direct Write Protein Patterns for Multiplexed Cytokine Detection from Live Cells using Electron Beam Lithography ...... 52

3.1 Introduction ...... 53

3.2 Results and Discussion ...... 55

3.3 Conclusions ...... 66

3.4 Materials and Methods ...... 67

3.5 References ...... 73

CHAPTER 4. Enhanced Stability of Granulocyte Colony-Stimulating Factor using Trehalose Glycopolymers ...... 76

4.1 Introduction ...... 77

4.2 Results and Discussion ...... 79

4.3 Conclusions ...... 102

4.4 Materials and Methods ...... 103

4.5 References ...... 112

CHAPTER 5. Trehalose Glycopolymers for Cell Stabilization and Preservation...... 114

5.1 Introduction ...... 115

5.2 Results and Discussion ...... 117

5.3 Conclusions ...... 133

5.4 Materials and Methods ...... 134

5.5 References ...... 140

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LIST OF FIGURES

Figure 2-1. Electron beam lithography process for multiple protein patterns ...... 35

Figure 2-2. Dose test of PolyProtek on protein patterning ...... 36

Figure 2-3. Proteins patterned by direct electron beam using PolyProtek as a resist ...... 37

Figure 2-4. Multiple layers and nanopatterning ...... 39

Figure 2-5. Fibronectin, VEGF, and bFGF patterns ...... 40

Figure 2-6. Multiple IgG nanopatterns ...... 42

Figure 2-7. HRP and IgG nanopatterns...... 42

Figure 2-8. Growth factors nanopatterns ...... 43

Figure 3-1. Non-specific adsorption of neutravidin-conjugated 30 nm gold nanoparticles to treated silicon substrates ...... 56

Figure 3-2. Direct electron beam patterning process for generating antibody patterns ...... 57

Figure 3-3. Effect of ascorbic acid on anti-TNFα patterning and immunoassay signals...... 57

Figure 3-4. Dark field micrographs for the sandwich assay of human IgG patterns ...... 58

Figure 3-5. ELISA detection of IL-6 and TNFα from LPS-stimulated RAW 264.7 macrophages (5 × 105 cells) ...... 59

Figure 3-6. The effect of anti-TNFα spin-coating concentration on immunoassay signals ...... 61

Figure 3-7. Cytokine detection with a bound silver enhanced gold nanoparticles immunoassay on surface immobilized micro- and nanopatterns ...... 61

Figure 3-8. Detection sensitivity of anti-TNFα patterns to varying concentrations of TNFα in media ...... 62

Figure 3-9. Detection sensitivity of anti-IL-6 patterns to varying concentrations of IL-6 in media ...... 63

Figure 3-10. Repeated processing and exposure cycles of anti-TNFα patterned on the same substrate ...... 64

Figure 3-11. Dark field micrographs of multiple antibody patterning and cytokine detection of IL-6 and TNFα ...... 64

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Figure 3-12. Multiplexed detection of IL-6 and TNFα from stimulated RAW 264.7 macrophages over time ...... 65

Figure 4-1. Expression and purification of His6-MBP-GCSF fusion protein following a Ni-NTA column ...... 80

Figure 4-2. Enterokinase digestion of His6-MBP-GCSF fusion protein, SDS-PAGE analysis...... 81

Figure 4-3. SDS-PAGE analysis of the purification of G-CSF from the His6-MBP-GCSF fusion protein ...... 82

Figure 4-4. MALDI spectra of purified G-CSF protein ...... 83

Figure 4-5. Bioactivity of purified G-CSF in NFS-60 cell proliferation assay ...... 83

Figure 4-6. 1H NMR spectrum of P4 ...... 85

Figure 4-7. 1H NMR spectrum of P5 ...... 87

Figure 4-8. 1H NMR spectrum of p(BMDO-co-trehalose) ...... 87

Figure 4-9. GPC analysis of p(BMDO-co-trehalose) before and after degradation with 5% KOH ...... 88

Figure 4-10. Cytotoxicity assay of P1-P3, 20 kDa PEG, and trehalose with NIH 3T3, RAW 264.7 macrophages, HDF, and HUVEC cells ...... 90

Figure 4-11. Cytotoxicity assay of p(BMDO-co-trehalose) before and after degradation and benzyl trehalose in HDF cells by LIVE/DEAD assay, and of the non-degraded, degraded products, and P1 in NFS-60 cells based on cell proliferation and LIVE/DEAD assay ...... 92

1 Figure 4-13. H NMR spectrum of benzaldehyde CTA (CDCl3) ...... 95

13 Figure 4-14. C NMR spectrum of benzaldehyde CTA (CDCl3) ...... 96

Figure 4-15. 1H NMR spectrum of benzaldehyde end-functionalized trehalose polymer from RAFT (D6DMSO) ...... 97

Figure 4-16. Silver stained SDS-PAGE analysis of G-CSF and P1-benzaldehyde conjugation with various reaction conditions after 48 hours ...... 98

Figure 4-17. Western blot analysis of G-CSF-P1 conjugate ...... 99

Figure 4-18. Bioactivity in NFS-60 cell proliferation of G-CSF and G-CSF-P1 conjugate after 5 cycles of lyophilization ...... 100

Figure 4-19. Bioactivity of G-CSF and G-CSF-P1 conjugate after heat ramping from 35-70 °C, increasing 5 °C every 30 minutes ...... 101

Figure 5-1. Fluorescent micrographs of NIH 3T3 cells incubated with aminooxy-biotin and Alexa Fluor 488 streptavidin ...... 118

Figure 5-2. Confocal fluorescent images of NIH 3T3 cells showing DAPI stained nuclei (blue channel) and trehalose polymer (red channel) ...... 120

Figure 5-3. Cell proliferation of Jurkat cells with the addition of P1 and P3 at 0.01, 0.1, and 1 mg/mL following heat treatment at 50 °C ...... 121

Figure 5-4. Cell proliferation of Jurkat cells with the addition of P3 at 1, 10, and 100 µg/mL with and without DMSO ...... 122

Figure 5-5. LDH activity of porcine platelets with and without excipients, trehalose or P3, stored at 4 °C and 23 °C ...... 123

Figure 5-6. Growth curves of bacteria after heated with trehalose or P3 (0.1 to 10 mg/mL) at 50 °C for 30 minutes ...... 124

Figure 5-7. Phase contrast micrographs of cells stained with Trypan blue solution with or without activated SLO ...... 125

Figure 5-8. Percent permeabilization of NIH 3T3 and HDF cells by SLO ...... 126

Figure 5-9. 1H NMR spectrum of P3-Rho ...... 127

Figure 5-10. Uptake of P3-Rho in HDF cells with and without permeabilization by SLO...... 128

Figure 5-11. Anthrone assay standard curve generated using trehalose (red circles) and P3 (green squares) ...... 129

Figure 5-12. Intracellular concentration as determined by anthrone assay in HDF cells when loaded at a range of 0 to 10 mg/mL of trehalose and P3 ...... 130

Figure 5-13. Percent cell viability of cryopreserved HDF cells with or without intracellular P3 and in preservation media (cryo-SFM) ...... 131

Figure 5-14. Effects of SLO concentration ranging from 0 to 750 ng/mL on cell viability of cryopreserved HDF cells ...... 132

LIST OF SCHEMES

Scheme 4-1. Structures of trehalose polymers (P1, P2, P3, and P1-benzaldehyde) ...... 84

Scheme 4-2. RAFT polymerization of BMDO and bMA ...... 85

Scheme 4-3. Synthesis of p(BMDO-co-trehalose) ...... 86

Scheme 4-4. Synthesis of benzaldehyde end-functionalized CTA ...... 95

Scheme 4-5. RAFT polymerization of trehalose monomer with benzaldehyde CTA (P1-benzaldehyde) ...... 96

Scheme 4-6. Preparation of G-CSF-P1 conjugate by reductive amination ...... 97

Scheme 5-1. Synthesis of aminooxy-functionalized fluorophore-labeled P3 (Atto590-AO-P3) ...... 119

Scheme 5-2. Synthesis of fluorescent P3 by free radical polymerization (P3-Rho) ...... 126

LIST OF TABLES

Table 2-1. Spin-coated thickness of varying concentrations of HRP and PolyProtek ...... 38

LIST OF ABBREVIATIONS

AFM Atomic force microscopy

AIBN 2,2-azobis(2-methylpropionitrile)

β-gal β-galactosidase bMA But-3-enyl methacrylate

BMDO 5,6-benzo-2-methylene-1,3-dioxepane

BMP Bone morphogenic protein

BSA Bovine serum albumin bFGF Basic fibroblast growth factor

CTA Chain transfer agent

DPN Dip-pen nanolithography

DMEM Dulbecco’s Modified Eagle Medium

DMSO Dimethyl sulfoxide

DTT Dithiothreitol

EBL Electron beam lithography

EC50 Half maximal effective concentration

ECM Extracellular matrix

EK Enterokinase

ELISA Enzyme-linked immunosorbent assay

ELISPOT Enzyme-linked immunospot

FBS Fetal bovine serum

FCS Fetal calf serum

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FDA Food and Drug Administration

FPLC Fast protein liquid chromatography

G-CSF Granulocyte colony-stimulating factor

GOx Glucose oxidase

GPC Gel permeation chromatography

GRAS Generally regarded as safe

HDF Human dermal fibroblast

HES Hydroxyethyl starch

HRP Horseradish peroxidase

HUVEC Human umbilical vein endothelial cells

ICC Intracellular cytokine cytometry

IgG Immunoglobulin G

IL-6 Interleukin-6

IPTG Isopropyl β-D-1-thiogalactopyranoside

LDH Lactate dehydrogenase

LPS Lipopolysaccharide

LSPR Localized surface plasmon resonance

MALDI Matrix-assisted laser desorption/ionization

ManLev N-levulinoyl mannosamine

ManNAc N-acetyl-D-mannosamine

MBP Maltose binding protein

Mn Number average molecular weight

MWCO Molecular weight cut-off

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Ni-NTA Nickel(II)-nitrilotriacetic acid

NMR Nuclear magnetic resonance

OD600 Optical density at 600 nm

PBS Phosphate buffered saline pCB Poly(carboxybetaine)

PDMS Polydimethylsiloxane

PEG Poly(ethylene glycol) pI Isoelectric point

PPL Polymer pen lithography

PRP Plasma rich platelets

RAFT Reversible addition-fragmentation chain transfer

RNase A Ribonuclease A

SAv Streptavidin

SDS-PAGE Sodium dodecyl sulfate poly(acrylamide) gel electrophoresis

SLO Streptolysin O

S/N Signal-to-noise ratio

Tm Melting temperature

TNFα Tumor necrosis factor alpha

VEGF Vascular endothelial growth factor

WHO World Health Organization

xiv

ACKNOWLEDGEMENTS

I would first like to express my deepest gratitude to my advisor, Prof. Heather D.

Maynard, for her mentorship, guidance, and support throughout the course of my research and graduate studies.

Special thanks to Dr. Sina Saxer, Dr. Erhan Bat, Dr. Juneyoung Lee, Dr. En-wei Lin, Dr.

Yang Liu, Jeong Hoon Ko, and Emma Pelegri-O’Day. I would not have been able to finish my dissertation without their mentorship and significant contributions.

I would like to also extend my appreciation to my committee members, Prof. Yong Chen,

Prof. Andrea Kasko, and Prof. April Pyle.

Thank you to all past and present Maynard group members: Dr. Chris Kolodziej, Dr.

Rocky Mancini, Dr. Steevens Alconcel, Dr. Juneyoung Lee, Dr. En-wei Lin, Dr. Nicholas

Matsumoto, Dr. Kathy Nguyen, and Dr. Caitlin Decker, Samantha Paluck, Natalie Boehnke, Eric

Raftery, Emma Pelegri-O’Day, Jeong Hoon Ko, Marco Messina, Priera Panescu, Kyle Tamshen, and Kathryn Mansfield.

Thank you to my family and friends for their support and encouragement. And finally, thank you to Carol, for her motivation and support during all these years.

VITA

EDUCATION

B.S. in Bioengineering 9/2006-6/2010 University of Washington, Seattle, WA Cum Laude with distinction in Bioengineering

RESEARCH EXPERIENCE

Graduate Student Researcher 4/2011-1/2016 University of California, Los Angeles Advisor: Heather D. Maynard, Ph.D.

 Design and development of nanoscale polymer patterning for biomedical applications  Development of glycopolymers for enhancement of therapeutic protein properties

Internship 1/2014-3/2014 Seattle Genetics, Bothell, WA

 Development and evaluation of novel drug linkers for antibody-drug conjugates

Undergraduate Researcher 6/2009-6/2010 University of Washington, Seattle Advisor: Thomas Horbett, Ph.D.

 Design of improved hemocompatible polyurethanes and coating protocol

Clinical Research Experience for Engineers Program 6/2009-9/2009 University of Washington, Seattle Advisors: Thomas Horbett, Ph.D., David Castner, Ph.D., Steven Goldberg, M.D., F.A.C.C.

PUBLICATIONS

Lau, U. Y.; Saxer, S. S.; Lee, J.; Bat, E.; Maynard, H. D., “Direct Write Protein Patterns for Multiplexed Cytokine Detection from Live Cells Using Electron Beam Lithography,” ACS Nano, 2016, 10, 723-729.

Bat, E.; Lee, J.; Lau, U. Y.; Maynard, H. D., “Trehalose Glycopolymer Resists Allow Direct Writing of Protein Patterns by Electron-Beam Lithography,” Nature Communications, 2015, 6, 6654.

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Lee, J.; Lin, E.-W.; Lau, U. Y.; Hedrick, J. L.; Bat, E.; Maynard, H. D.; “Trehalose Glycopolymers as Excipients for Protein Stabilization,” Biomacromolecules, 2013, 14, 2561-2569.

MANUSCRIPTS IN PREPARATION

Matsumoto, N. M.; Raftery, E. D.; Lau, U. Y.; Grubbs, R. H.; Maynard, H. D., “Protein- Reactive Unsaturated PEG Analogs by ROMP,” to be submitted.

TEACHING EXPERIENCE

Teaching Assistant 9/2014-12/2015 University of California, Los Angeles Chemistry 153L: Biochemical Methods. Dr. Anne Hong-Hermesdorf, Fall 2014; Winter 2015; Spring 2015; Fall 2015.

Supervised Students Bridges Summer Research Program 7/2015-9/2015 Student: Jonathan Moran, Improving upon therapeutic protein drugs with protein- polymer conjugates

POSTERS/PRESENTATIONS

Glenn T. Seaborg Symposium. “Direct Write Protein Patterns using Electron Beam Lithography.” UCLA. Oct. 19, 2015.

Chemistry-Biology Interface (CBI) Day 2014. “Cytokine Detection by Antibody Micro/Nano-Patterns using Trehalose Glycopolymer and E-Beam Lithography”. UCLA. Aug. 27, 2014.

Chemistry-Biology Interface (CBI) Day 2013. “Nano/microscale Biosensor for Extracellular Cytokine Detection.” UCLA. Sept. 5, 2013.

Molecular Biology Institute Annual Retreat. “Plasmonic Biosensor Platform for Detection of Cell-signaling Molecules.” Lake Arrowhead, CA. Oct. 19-21, 2012.

AWARDS/MEMBERSHIPS

Center for Scalable and Integrated NanoManufacturing 2012-2016 NIH Chemistry-Biology Interface Training Fellow 6/2011-9/2014 University of Washington’s Annual Dean’s List 2006-2010

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CHAPTER 1

INTRODUCTION AND LITERATURE REVIEW

1.1 Overview of Trehalose

Trehalose is a disaccharide comprised of two glucose units in a α,α-1,1-glycosidic linkage. The linkage makes trehalose a non-reducing sugar that is not easily hydrolyzed by acid and not readily susceptible to cleavage by most α-glycosidases.1,2 In fact, trehalose is among the most chemically stable sugars and has low reactivity toward most proteins and other reactive molecules.3 Trehalose is naturally synthesized and accumulated in large amounts as a response to environmental stresses by insects, plants, fungi, and organisms, but not mammals, when they are desiccated (androhydrobiosis) or exposed to low temperatures (cryobiosis).4,5 Trehalose has been shown to function as an energy source or as a protective agent to dramatically increase resistance against extreme environmental stresses by protecting cellular membranes and proteins.6-8 This sugar has also been reported to protect cells and biomolecules against damage from radiation and radical damage derived from radiation.9,10 The exact mechanism of how trehalose protects these biological molecules is still not fully understood, though three credible hypotheses exist. In the water replacement hypothesis, the hydroxyl groups on trehalose molecules directly interact with the cellular components via hydrogen bonding, which helps maintain protein conformation and consequently its activity.11,12 The high glass transition temperature of trehalose (~117 °C) supports the vitrification hypothesis.13-15 Due to the high viscosity in the glassy state, molecules entrapped within the glassy matrix of the sugar experience reduced flexibility and are thus stabilized against unfolding. Finally, the water entrapment theory hypothesizes that trehalose helps maintain hydration by trapping a layer of water between the sugars and molecule.16 Each hypothesis is supported by substantial evidence and the protective capabilities of trehalose may be dependent on a combination of multiple mechanisms depending on the specific type of stress and environment.

The unique chemical and physical properties of trehalose have made this sugar an attractive material in commercial and industrial applications including cosmetics, food production, and pharmaceuticals, as well as in academic research.17,18 The U.S. Food and Drug

Administration (FDA) has classified trehalose as generally regarded as safe (GRAS) in 2000 and trehalose is present in many food products. In addition, trehalose has been employed as an excipient in four pharmaceutical products on the market, namely Avastin, Lucentis, Herceptin

(Genentech/Roche), and Advate (Baxter).18 The prevalence and safety of this disaccharide has generated interest and development towards additional uses in many applications.

1.2 Protein Therapeutics and Challenges

Proteins are widely used in the clinical and research settings as therapeutics and as laboratory reagents for research applications. Protein therapeutics have great advantages over small-molecule drugs due to their highly specific activities and functions, and fewer adverse effects.19 Over 130 different proteins or peptides have been approved for clinical use by the US

FDA with sales in 2010 reaching over 108 billion dollars.19,20 However, proteins are generally unstable, presenting challenges with their handling, transportation, and long-term storage.

Proteins can easily denature and lose activity under a variety of stresses, including desiccation, light, and shifts in temperature and pH.21-24 In addition, protein therapeutics present pharmacokinetic challenges for in vivo protein stability. When introduced into the bloodstream, protein drugs suffer from immunogenicity, rapid clearance and degradation leading to short half- lives.25 Formulation optimization of protein therapeutics aims to increase the protein stability and thus solve many of these challenges.

Protein instability can be classified into physical or chemical instability. Common physical instabilities include protein unfolding, aggregation, and adsorption. Protein unfolding

3 can be caused by heat, changes in pH, or by denaturing chemicals. Aggregation can result from protein unfolding, leading proteins to form dimers and oligomers. This can reduce protein solubility, inactivate proteins, and reduce bioactivity. Adsorption relates to the protein interaction with surfaces. The protein can unfold, exposing the hydrophobic residues to interact with the surface.26 Common chemical protein damage pathways include oxidation, hydrolysis, and deamidation. Especially, amino acids methionine and cysteine readily undergo oxidation.

Oxidation of cysteine yields a disulfide bond, which may lead to aggregation and loss of activity.27 Although peptide bonds are fairly stable to hydrolysis, the aspartic acid-proline peptide bond is the most susceptible to hydrolysis under acidic conditions.28 Deamidation can occur, where amide groups on asparagine and glutamine residues are hydrolyzed to free carboxylic acid groups, which can significantly alter protein activity.29,30

To address the instability of protein drugs, many excipients including amino acids, salts, sugars and polyols, surfactants, and polymers have been used as additives in protein formulations for storage.31 Amino acids, usually in addition with other excipients, most likely stabilize proteins by preferential exclusion, where the molecule is excluded from the protein surface, keeping the protein surface hydrated and correctly folded in its native state.32 Common amino acids used as excipients include arginine, histidine, and glycine. The addition of arginine has been reported to reduce aggregation of thermally inactivated lysozyme, as well as interleukin-6 and monoclonal antibodies.33,34 Effects of arginine on the therapeutic protein, fibroblast growth factor 20 (FGF20), had a maximal effect at pH 6.0 with a 1000-fold increase in solubility.35

Histidine and glycine have been used as excipients in approved protein pharmaceuticals for a humanized monoclonal antibody and recombinant antihemophilic factor.36 Surfactants are also common excipients, which reduce the surface tension of the protein solution, reducing

4 aggregation and mediating protein refolding. Polysorbate 20 and polysorbate 80 are surfactants widely used in protein pharmaceuticals, and have been shown to help stabilize proteins by reducing surface denaturation.37 Other surfactants used as protein stabilizers include sodium dodecyl sulfate and poloxamers. Hydrophilic polymers are also used and stabilize proteins by the excluded volume effect, which thermodynamically favors the compact native structure of the protein. Examples include polysaccharides (dextran), hydroxyethyl starch (HES), poly(ethylene glycol)s (PEGs), gelatin, polyvinyl alcohol, and polyvinylpyrrolidone. Sugars and polyols are widely used in protein formulations with sucrose, trehalose, sorbitol, and mannitol as among the most employed stabilizers. Sucrose has been reported to stabilize rhDNase and rhIFN-γ;38,39 however, sucrose can hydrolyze to fructose and glucose, which are reducing sugars and can react with amino groups in proteins.

The properties of trehalose have made the disaccharide extremely useful as a non-specific protein stabilizer among other applications. The addition of trehalose has been demonstrated to enhance thermal stability of various biological molecules including a wide range of proteins with different physico-chemical properties. Sola-Penna and Meyer-Fernandes showed that the addition of 1.5 M trehalose increased the thermal stability of yeast pyrophosphatase by increasing the time to inactivation at 50 C by 13-fold.40 A follow up study showed that trehalose also protected against conformational changes promoted by guanidinium chloride with the same enzyme.41 In addition, trehalose not only protected against thermal inactivation, but also was shown to increase the catalytic activity of phosphofructokinase.42 Lin and Timasheff studied the interaction of trehalose and the model protein, ribonuclease A (RNaseA), by thermal unfolding experiments.43 Increasing trehalose concentration led to an increase in the melting temperature

(Tm), where the Tm increased by 10 C in a 1 M trehalose solution compared to buffer only.

Thermal stabilization of RNaseA with trehalose was also reported at pH 2.8 and pH 5.5.44

Kaushik and Bhat investigated thermal denaturation of five well-characterized proteins, RNaseA, lysozyme, cytochrome c (cyt c), α-chymotrypsinogen (α-CTgen), and trypsin-inhibitor (Trp-Inh) over a range of trehalose concentrations (0 to 2 M) and pH values (2.5, 4.0, and 7.0).7 The chosen proteins represented a large range of differing properties, including the molecular size from 12.3 kDa to 25.7 kDa, and isoelectric point (pI) ranging from 4.1 to 10.7. Their results showed that all proteins exhibited increased transition temperature (Tm) with increasing trehalose concentrations. Maximal effects were observed with RNaseA with an increase in Tm of 18.2 C at 2 M trehalose and pH 2.5. Furthermore, RNaseA activity was retained in the presence of trehalose at elevated temperatures. Other examples of thermal stabilization with trehalose include alcohol dehydrogenase and acidic fibroblast growth factor.45,46

Often the stability of liquid protein formulations is still insufficient, and requires preparation in the dried state. However, the process of lyophilization, or freeze-drying, usually denatures proteins. The addition of trehalose has been shown to increase the stability of many different proteins during the steps of freeze-drying, desiccation, and storage. Carpenter and

Crowe reported storage stability of freeze-dried lysozyme and phosphofructosekinase.47 Fourier- transform infrared spectroscopy analysis of the dried lysozyme in the presence of trehalose indicated that trehalose preserved the protein conformation during freeze-drying. Miller and co- workers reported lactate dehydrogenase stabilization after freeze-thawing and centrifugal vacuum-drying/rehydration with a mixture of trehalose and sodium tetraborate.48 Activity of

80% was retained for freezing and 65% for drying, even after storage in these conditions for several weeks. Studies on freeze-dried β-galactosidase (β-gal) by Kawanishi and co-workers reported that the local mobility of the protein is more important towards stability than the glass

6 transition temperature. Increased amount of trehalose decreased local mobility, resulting in enhanced storage stability.49 Monoclonal immunoglobulin M (IgM) and immunoglobulin E (IgE) antibodies have been stabilized with trehalose during freeze-drying.50,51 Mixtures of trehalose and HES have shown to increase storage stability of recombinant human interleukin-11.52

However, the contribution from trehalose was more significant towards freeze-drying protection as HES alone did not provide stabilization. Moreover, trehalose has been shown to stabilize

RNaseA and biosynthetic Factor VIII to lyophilization, demonstrating its utility as a general protein stabilizer.53,54

Trehalose has been also extensively used for vaccine applications, stabilizing viruses to improve their shelf life and heat stability. Typically, live attenuated viruses are freeze-dried for prolonged storage. Truong-Le and co-workers reported spray-drying a measles vaccine that included trehalose, sucrose, L-arginine, human serum albumin, and glycerol and showed that the vaccine formulation was stable for 8 weeks at 37 C.55 The increased shelf life was well above the general requirements of 1 week storage at 37 C as set by the World Health Organization

(WHO). Other dried viruses shown to be stabilized with trehalose include 17D Yellow Fever virus,56 Newcastle disease virus,57 and respiratory syncytial virus.58 In addition, influenza virus- like particle vaccine was stabilized with trehalose for coating microneedles.59 Trehalose has successfully been shown to provide protection to various biologics and under many different stress conditions showcasing the tremendous potential of trehalose as a non-specific stabilizer.

The main limitations in protein therapeutics are their general inherent instability, insufficient pharmacokinetic properties, and immunogenicity. The covalent attachment of poly(ethylene glycol) (PEG) to proteins, a process referred to as PEGylation, has become a widely used technique for addressing these challenges. First reported by Abuchowski and co-

7 workers in 1977, PEG was conjugated to bovine serum albumin and bovine liver catalase.60,61

They reported that the PEGylated proteins exhibited lower immunogenicity and increased circulation times while retaining their bioactivities. Since the discovery and advent of recombinant protein technology, much research has been conducted in this field. The first

PEGylated protein drug approved by the FDA in 1990 was Adagen, or PEGylated adenosine deaminase, which is used for the treatment of severe combined immunodeficiency disease. Since then, there are now currently eleven FDA-approved PEGylated products in use in the clinic.

Some select examples include PEG-INTRON (PEG-interferon alfa-2b, approved 2001) for the treatment of chronic hepatitis C, and Neulasta (PEG-G-CSF, approved 2002) for treating febrile neutropenia. In all cases, the pharmacokinetic properties are significantly improved after

PEGylation. PEGylated interferon has an increased circulation half-life (48-72 hr) relative to the native protein (7-9 hr). Similar improvements is seen with PEG-G-CSF with an increased serum half-life (42 hr) relative to native G-CSF (3.5-3.8 hr). Longer serum half-lives mean less frequent dosing, which is advantageous for patient compliance. However, modifying proteins with PEG has its drawbacks, most notably the decrease in activity relative to the unmodified protein.

However, the benefits of longer circulation time and increased proteolytic stability from

PEGylation compensates for the loss of activity, resulting in conjugates that still perform better than their native counterparts.62

Decades of clinical experience and research in PEGylated products have been valuable, not only in learning about its benefits, but also in discovering many types of potential problems in using these products. These side effects include hypersensitivity and immunogenicity against

PEG, accumulation of PEG in various organs, toxicity of side products formed during the

8 synthesis of PEG, and their non-biodegradability.63-65 These disadvantages provide a strong motivation for the development of alternative polymers that can be used for protein therapeutics.

Many other biocompatible, but non-biodegradable, polymers have been reported for drug delivery applications and tested to enhance the pharmacokinetic properties of proteins. These examples include poly(glycerol)s,66 poly(oxazoline)s,67-69 poly(vinylpyrrolidone),70,71 and poly(N-(2-hydroxypropyl)methacrylamide)s.72-74 These polymers as PEG alternatives may eliminate the immunogenicity associated with PEG, but are susceptible to similar side effects due to non-biodegradability and accumulation in vivo.

One category of PEG alternatives are biodegradable polymers that alleviate the issue of polymer accumulation in the body. Poly(amino acid)s, such as poly(glutamic acid), poly(hydroxyethyl L-glutamine), and poly(hydroxyethyl L-asparagine) have been demonstrated to be enzymatically degradable, but still have issues with immunogenicity and complement activation.75,76 HES, a biodegradable semi-synthetic polymer, has been studied in vivo with HES conjugates of erythropoietin and interferon-alpha.77 Polysialic acid, degradable by neuraminidase, has been reported to extend the effects of insulin to over 6-9 hours compared to unmodified insulin (3 hours),78 and reduce immunogenicity of an single-chain Fv fragment.79

In recent years, the development of polymers with additional benefits other than improved pharmacokinetic properties has been of great interest. These next generation polymers hold tremendous potential as PEG alternatives. For example, polymers that stabilize proteins against environmental stressors can be used to eliminate the loss of protein activity associated with storage and handling. Maynard and co-workers reported the development of glycopolymers with trehalose side chains for enhancing stability to conjugated lysozyme.80 The lysozyme- glycopolymer conjugate retained activity against freeze-drying and heating at 90 °C. Later,

Maynard and co-workers demonstrated these trehalose glycopolymers can be used as excipients to stabilize horseradish peroxidase (HRP), glucose oxidase (GOx), and β-gal against lyophilization and heat stress.81 Further work regarding the trehalose glycopolymers has been aimed towards stabilizing more clinically relevant protein therapeutics, such as granulocyte colony-stimulating factor (G-CSF) as discussed in detail in Chapter 4. A designed heparin- mimicking polymer, poly(styrene sulfonate-co-PEGMA) (p(SS-co-PEGMA)), has been reported by the same group to stabilize basic fibroblast growth factor (bFGF).82 The bFGF-p(SS-co-

PEGMA) conjugate showed protection and retained activity to therapeutically relevant stresses including heat, acidic conditions, extended storage (over 40 days), and proteolytic degradation.

Keefe and Jiang reported the use of poly(zwitterionic) protein conjugates for protein stabilization.83 In this example, poly(carboxybetaine) (pCB) was conjugated to α-chymotrypsin and exhibited stability at elevated temperatures without any decrease in binding affinity or bioactivity. Recently, Jiang and co-workers investigated the pharmacokinetic properties of pCB.84 Uricase was encapsulated in a pCB gel network, which exhibited enhanced stability, increased in vivo circulation half-life, and no immunogenicity response. The pCB polymer network showed superior improvements over the PEGylated version.

1.3 Protein Patterning

The immobilization and positioning of biomolecules in well-defined patterns while retaining their bioactivity is important for the development of biomaterials and bioactive surfaces.85,86 There are a variety of patterning techniques that have been used for the immobilization of biomolecules.87-89 These include ink-jet printing, stamping techniques such as microcontact printing and nanoimprint lithography, scanning probe-based methods, and other maskless lithography techniques. However, generating these protein arrays often denatures the

10 biomolecules, which loses activity during the patterning process. Some limitations of various patterning techniques includes exposure and use of organic solvents, heating, drying steps, vacuum conditions, and surface interactions. This section focuses mainly on materials and methods that have been used in various techniques to help maintain protein activity during the direct write of biomolecules onto surfaces.

The majority of techniques for producing protein patterns and arrays involve indirect processes. Structures and features are generated with desired properties, then the proteins are directed onto the patterns by some type of specific interaction. In this manner, the biomolecules are left out of any harsh processing conditions encountered during patterning, and are used towards the end of the process to ensure optimal protein activity. High resolution streptavidin- patterned surfaces are commonly prepared by generating biotin-functionalized surfaces.90,91

Another example involves patterning 16-mercaptohexadeconic acid onto gold substrates, followed by subsequent processing and utilizing coupling chemistries to attach proteins such as streptavidin and rabbit IgG.92,93 Many other strategies of presenting functionalized surfaces or making protein-repellent and adhesive areas have been developed.

Direct protein patterning is more beneficial in that the processing methodologies become simpler with fewer number of steps, but has significant challenges in maintaining protein structure and activity. One main issue is the compatibility of the protein-surface interaction, such that the protein does not denature. Substrates with high surface hydrophobicity tend to denature proteins. Loss of protein activity also occurs typically when the solutions are dried out.94

Commonly used protective agents added to help alleviate these issues include glycerol, disaccharides, and low molecular weight PEGs.95 Glycerol is a known additive for protein stabilization and is also useful in circumventing evaporation problems. A solution of 40%

11 glycerol in phosphate buffered saline (PBS) has been used to slow down dehydration during the microarraying of antibodies.96 The addition of glycerol facilitated optimal immobilization of the antibodies on the surfaces. Chrisey and co-workers have also demonstrated that a 40% glycerol

PBS solution improved pattern uniformity and maintained activity using a laser evaporation direct write technique.97 Higher concentrations of glycerol (60% in PBS) has been employed in a liquid nanodispensing technique that utilizes a modified atomic force microscopy (AFM) probe to pattern nanoparticles and proteins.98

Protein-surface interactions are especially important in soft lithography techniques such as microcontacting printing. An elastomeric polymer, typically made of polydimethylsiloxane

(PDMS), is used to transfer patterns of a selected material onto a substrate through conformal contact, similar to that of stamp and ink. Several important factors need to be considered for direct protein stamping to maintain protein structure and activity. Hydrophilic stamps are used to prevent protein denaturation, and designed such that the affinity of the protein solution has to be greater towards the substrate than towards the stamp for proper transfer. The activity of microcontact printed anti-goat antibodies was compared against directly absorbed antibodies onto glass substrates. A loss of 30% binding efficiency to a fluorescent antibody was observed.99

The following examples highlight methods of preserving protein activity during microcontact printing. Williams and co-workers have used trehalose as a protective and fixing agent to preserve the structure and organization of proteins during subsequent processing in microcontact printing.100 In this process, after the target protein is deposited on the stamp substrate, a thin protective layer of trehalose is spin-coated on top of the protein layer, followed by the deposition of a polymer layer as support. Transfer onto the substrate was achieved using a cyanoacrylate- based adhesive and subsequent separation of the substrate from the adhesive. A polycationic

PEG-grafted copolymer, functionalized with RGD peptide and fibronectin, has been used in conjunction with microcontact printing for cell adhesion to patterned regions by Danuser and co- workers.101

The use of hydrogel “wet” stamps in microcontact printing have been developed to circumvent issues of protein denaturation on hard, dry substrates. Patterns of bovine serum albumin (BSA) and GOx immobilized on PEG hydrogel surfaces maintained 50% of their initial activity, while immobilization on a polystyrene substrate resulted in only 20% retention after 24 hours.102 Dittmer and co-workers synthesized a hydrogel stamp material consisting of 2- hydroethyl acrylate and PEG diacrylate and was able to retain the activity of antibodies against human parathyroid hormone following stamping.103 In addition to PEG hydrogels, Whitesides and co-workers have used agarose hydrogels as stamps to absorb proteins and minimize denaturation. These materials could be stamped up to twenty times before requiring re-inking.104

Protein gradients can be readily prepared due to diffusion of the protein throughout the hydrogel.

Hydrogel stamps have the benefit of entrapping proteins in an aqueous environment and not requiring a drying step after protein application.

Ink-jet printing is another material deposition technique in which proteins can be used as inks for patterning and therefore can be used to directly deposit proteins onto surfaces. The technique requires generation of a pressure pulse to propel the liquid out of a nozzle onto the substrate, which is commonly achieved thermally or by a piezoelectric actuator. The resulting stresses in the form of high temperatures or high shear rates can denature proteins. With a thermal ink-jet printer, β-gal printed patterns lost 15% of its activity from the thermal shock.105 In a piezoelectric ink-jet printer, loss of activity of GOx was caused from the printing conditions.106 Holloway and co-workers have shown that the addition of sugars (trehalose and glucose at a 10:1 w/w ratio) to

HRP resulted in higher retained bioactivities using a piezoelectric actuated ink-jet printer.107

Printed HRP without sugar lost about 60% of its activity due to the mechanical stress from ejection of the liquid, while the patterns printed with the sugars were able to retain 100% activity at lower shear rates. Polymers have also been used to aid in printing delivery and maintain activity of the printed proteins. Schuhmann and co-workers added vinyl acetate and ethylene copolymer with GOx, which helped with enzyme immobilization and uniformity.108 A systematic investigation of the effects of various other polymers as viscosity modifiers in piezoelectric ink-jet printing on the activity of HRP was conducted by Risio and Yan.109 PEG of different molecular weights, ethylene glycol, glycerol, poly(vinyl alcohol), and carboxymethyl cellulose at a range of concentrations with GOx were evaluated. Their results showed that carboxymethyl cellulose increased viscosity to the ideal range for printing without negatively affecting HRP activity.

Direct protein patterning at nanoscale resolutions encounter similar challenges. Mirkin and co-workers have developed dip-pen nanolithography (DPN), which uses a modified AFM tip, to deposit dry proteins onto surfaces.110-112 These methods involve chemical modification to the AFM tips as well as the substrate to maintain the biological activity of the printed proteins.

Gold-modified AFM cantilevers were coated with a thiolated PEG to make the tips hydrophilic and to prevent denaturing the adsorbed proteins. In this manner, lysozyme and IgG protein nanopatterns were created on gold surfaces.

Electron beam (e-beam) lithography (EBL) is another technique commonly used for nanofabrication. During e-beam processing, vacuum conditions and irradiation by high energy electrons damage and denature proteins, and therefore direct protein write methods with this technique are typically not considered. However, some limited methods have been shown to

14 protect biological molecules from this patterning process. Poly(acrylic acid) as a resist has been used for the direct e-beam patterning of bacteriorhodopsin.113 Omenetto and co-workers reported the use of silk as a positive and negative resist for EBL.114 Incorporation of silk with green fluorescent proteins and HRP during e-beam processing helped retain fluorescence and activity.

In another example, Maynard and co-workers have developed a trehalose glycopolymer resist that enabled the direct writing of proteins under EBL conditions.115 Covered in Chapters 2 and 3, the addition of the trehalose glycopolymer allowed the retention of activity of a wide range of proteins with high resolution features.

1.4 Cell Preservation

The ability for long-term preservation of living cells holds tremendous potential for human therapeutics and research applications.116,117 Preservation of cells is critical for establishing cell banks for future use, permits quality control testing prior to use, and facilitates shipping and transportation between research centers and medical facilities. Cryopreservation is the most widely applied method for long-term storage of cells utilizing dimethyl sulfoxide

(DMSO) as the most commonly used cryoprotectant agent. These cryoprotectant agents work by penetrating the cell membrane and replacing part of the water in the cell, reducing ice formation and thus reducing cell damage.118 However, there are significant drawbacks to these cryoprotectants, such as cytotoxicity and negative influence on cell differentiation. DMSO exposure to mouse embryonic stem cells has been shown to affect the epigenetic system through

DNA methylation and histone modification.119 Cells cryopreserved with DMSO have been clinically used for direct infusion into patients to avoid cell loss from post-thawing, but adverse health effects and even case fatalities have been attributed to the toxicity of DMSO.120 Many different cell types, such as adult and embryonic stem cells, islet cells, and lymphocytes are

15 being developed into new and emerging cell therapy products. Such therapies may require cell pooling and direct transplantation of cells to its targeted site. Therefore, there is a significant need to decrease cellular toxicity associated with DMSO by removal techniques or through the use of alternative preservation solutions and agents.

There are a number of approaches in developing compounds for partially or fully replacing DMSO. Ficoll and dextran polymers at concentrations from 43 to 49 weight percent in combination with 11 to 27 weight percent of ethylene glycol have been shown to preserve mouse embryos to greater than 96% viability in vitro and 62% viability in vivo.121 Reduction of DMSO using HES and albumin mixture have been evaluated in marrow cells.122 Polyvinylpyrrolidone has been shown to be an effective cryoprotectant for adipose tissue-derived adult stem cells, but some decrease in post-thaw cell viability was observed.123 Derivatives of e-poly-L-lysine were able to preserve mesenchymal stem cells with retained potential for proliferation and differentiation.124 Other common alternatives include glycerol and ethylene glycol.125,126 More recently, natural osmoprotectants, hydroxyectoine and ectoine, have been used to replace

DMSO.127

There is an active interest in sugars, particularly trehalose, as an alternative cryoprotectant and stabilizer. Accumulation of trehalose occurs in many organisms and serves as a protective agent when the organism is under dehydration stress. These organisms include many types of bacteria,128 resurrection plants,129 crustaceans, such as brine shrimp, Artemia, and dry baker’s yeast,130 but trehalose does not naturally occur in mammals. The ability to confer protection using trehalose in mammalian cells would then have huge implications for biotechnology. Many techniques have been used for the introduction of intracellular trehalose,

16 and research has suggested that trehalose must be present on both sides of the cell membrane for stabilization.131

One of the first examples where trehalose was used to preserve mammalian cells was reported by Hayek and co-workers in 1997.132 The native mammalian cell membrane mechanisms were utilized to introduce both DMSO and trehalose into pancreatic islet cells during cooling. The leakiness of the cell membranes during the cooling phase transition was exploited to load trehalose into the islet cells. Following cryopreservation, 92% of islet cells loaded with trehalose and DMSO were recovered compared to 58% with using DMSO alone.

The cells were also transplanted into mice and demonstrated to remain viable. Loading of trehalose by osmotic shock was achieved with red blood cells.133 Trehalose was loaded into red blood cells through a combination of osmotic imbalance and phospholipid phase transition at 37

C over a span of 7 hours with a final intracellular trehalose concentration of up to 40 mM.

Sukhorukov and co-workers used reversible electropermeabilization to open pores in mouse myeloma cells to load trehalose to a concentration of 100 mM.134 Delivery of trehalose via liposomes was reported by Acker and co-workers in 2009 into human red blood cells.135

Cryopreservation of trehalose-loaded red blood cells had a post-thaw recovery of 66% compared to the control of 29%, and had improved post-thaw membrane quality. Mechanical methods that have been used include microinjection, where the material is injected directly into the cells using a micropipette. Toth and co-workers utilized this technique to inject trehalose into human and mouse oocytes, and zygotes.136,137 The survival rate of the intracellular trehalose group (66%) were significantly better than the control group (13%) and as well as to cell stabilized with extracellular trehalose (22%), suggesting the importance of trehalose loading within the cells.

In a different approach to introduce trehalose into mammalian cells, Levine and co- workers transferred genes for trehalose synthesis from E. coli into human primary fibroblasts using a recombinant adenovirus vector.138 The fibroblasts expressing trehalose were able to remain viable in the dried state for up to five days. Okuda and co-workers have cloned a trehalose transporter gene from an anhydrobiotic insect and introduced the gene into fibroblasts, ovary cells, and hepatoma cells.139 Additionally, methods for introducing engineered pores into cell membranes for the uptake of trehalose has been developed. Toner and co-workers have used a pore forming bacterial toxin, α-hemolysin, that forms 2 nm homoheptameric transmembrane pores into lipid bilayers.140,141 Control over closed and open positions of the pore could be modulated with the addition or removal of zinc ions, and they showed that loading with 0.2 M intracellular trehalose protected 80% of mouse fibroblasts and 70% of human keratinocytes. As an alternative to bacterial pore forming agents, a pH-responsive biopolymer has been synthesized that increases permeability of the membrane to trehalose and showed uptake into human erythrocytes.142

The various methods of incorporating trehalose into cells have been shown to aid in preserving many different cell types. Our previous work with trehalose glycopolymers have demonstrated their ability to stabilize proteins. The next step of expanding the use of trehalose glycopolymers towards stabilizing cells was investigated and presented in Chapter 5.

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CHAPTER 2

TREHALOSE GLYCOPOLYMER RESISTS ALLOW DIRECT WRITING OF PROTEIN PATTERNS BY ELECTRON BEAM LITHOGRAPHY

This chapter contains portions of an edited version of a paper published as: Bat, E.; Lee, J.; Lau, U. Y.; Maynard, H. D., “Trehalose Glycopolymer Resists Allow Direct Writing of Protein Patterns by Electron-Beam Lithography,” Nat. Commun. 2015, 6, 1-8.

2.1 Introduction

The patterning of proteins at micro- and nano-scale resolutions is useful in many applications including biosensors, diagnostics, and tissue engineering.1-4 There are several examples of protein patterned arrays for a variety of purposes such as identifying protein-protein interactions for proteomics and drug delivery research,5 controlling cell adhesion for modulating cell behavior,6 and detecting and measuring disease biomarkers for diagnostic assays.7 Ongoing advances for fabricating miniaturized protein detection systems, especially at nanometer length scales, are highly desired but challenging to fabricate.8 The push towards further miniaturization drives the ability to generate ultra-high density protein patterns, resulting in the benefits of lower reagent volumes, better sensitivities, and high-throughput analyses.

There are a variety of patterning techniques that have been used for biomolecule immobilization with micro- and nanoscale features.9,10 Ink-jet printing is capable of depositing picoliter volumes with positional accuracy of 30 µm. This technique has been used to pattern growth factors, such as fibroblast growth factor 2 (FGF2) and bone morphogenic protein 2

(BMP2), to study their effects on stem cell differentiation.11 Stamping techniques include microcontact printing and nanoimprint lithography. Microcontact printing typically involves coating the target molecule on an elastomeric polymer then transferring it to the substrate by conformal contact. This has been used for creating patterns of antibodies for biosensor applications.12,13 Nanoimprint lithography relies on a fabricated template, which is pressed against a deformable polymer-coated substrate, and then cured by heat or light. Through this technique, high resolution streptavidin-patterned surfaces were generated utilizing the biotin- streptavidin interaction.14,15 Other techniques include scanning probe-based methods that utilize cantilever tips for depositing materials on surfaces and dip-pen nanolithography, which has been

32 used for patterning streptavidin and rabbit IgG.16,17 A more recently developed scanning probe- based method, polymer pen lithography, utilizes a soft elastomeric tip that combines the capabilities of dip-pen nanolithography and large-area contact printing.18 Another method, electron beam lithography (EBL), utilizes focused beams of electrons to generate patterns. Libera and co-workers found that PEG irradiated with electrons cross-links to silicon substrates forming hydrogels to adsorb or repel extracellular matrix proteins.19 PEG-based polymers with functionalized end groups were patterned by EBL and then incubated with proteins to specifically bind to patterned areas. These examples include protein nanopatterns formed by oxime bonds,20 and RGD peptide and basic fibroblast growth factor (bFGF) patterning for cell attachment.21

Direct write patterning of biomolecules at micrometer and nanometer length scales has the potential to enable pattern generation with fewer fabrication steps. In addition, direct write approaches facilitate the generation of geometrically complex multicomponent patterns at the micrometer and nanometer scale with the advantage of not having a limit in the number of different proteins that can be patterned, as opposed to indirect strategies. Yet despite these advantages, only a few examples have been reported to directly pattern biomolecules on surfaces at the submicron scale.18,22-25 The majority have employed physical deployment onto surfaces using an atomic force microscopy (AFM) tip such as dip-pen lithography or polymer pen lithography; these techniques involve pressing or flowing the material onto a surface and do not require a resist. This is because the approaches in which radiation is utilized, such as EBL, require the development of resists that are water soluble and protect the biomolecules from denaturation during irradiation and other processing steps. These requirements have significantly hindered the investigation of direct write biomolecule patterns by techniques such as EBL.

EBL is a maskless patterning technique that generates user-designed complex patterns at high resolution. Although a serial technique, EBL offers nanometer-scale alignment capability, which enables interfeature spacings that are so small that different protein features may be touching or arrayed one on top of the other allowing for complex, multiplexed patterns.26 The majority of protein patterning by EBL has been accomplished by indirect approaches,19,26-30 because the high vacuum and high-energy radiation inactivates proteins. Indeed, the harsh conditions required have been exploited to pattern by selective ablation of proteins on electron- beam exposure.27,28 Thus far, there have only been two resists that have been employed for direct protein patterning by EBL; proteins bacteriorhodopsin and green fluorescent protein were patterned using poly(acrylic acid) and silk as resists.22,24 In both studies, the authors noted that the proteins used had exceptionally stable structures, which enabled them to be stable under harsh conditions of EBL.

Our group has previously demonstrated a new resist material based on trehalose glycopolymers that stabilized a variety of proteins to repeated exposure to vacuum and to electron beams allowing for direct write by EBL, as well as multiplexing.31 An emphasis on patterning sensitive proteins, such as growth factors, as well as generating nanoscale features with multiplexing are presented in this chapter.

2.2 Results and Discussion

The trehalose glycopolymer utilized for EBL has a polystyrene backbone and trehalose side chains (PolyProtek, Figure 2-1). The glycopolymer was anticipated to crosslink under electron beam irradiation because polystyrene is known as a negative-tone electron beam lithography resist.30,32 Upon exposure to electron beams, the glycopolymer was crosslinked and covalently bound to the silicon substrates. Overexposure was observed at doses above 70

µC/cm2, while the patterns looked sharp at an area dose of 40 µC/cm2 (Figure 2-2). When irradiated by electrons, the trehalose polymer crosslinks to the silicon substrate by a mechanism similarly observed for other polymers, such as PEG.19,33

Trehalose is known to accumulate in large amounts in many types of plants and animals under desiccation stress, which helps with increasing the resistance of these organisms, as well as cells and biomolecules, against such environmental stresses.34-36 Recently, we have reported that polymers with trehalose side chains provide superior protection to proteins against heating and freeze drying compared to trehalose alone.37,38 The glycopolymers combine the advantageous properties of the osmolyte and non-ionic surfactant class of stabilizers into one material. The trehalose glycopolymer, PolyProtek, was utilized here to stabilize proteins to EBL processing conditions.

Figure 2-1. Electron beam lithography process for multiple protein patterns. Spin coating with PolyProtek and target protein solution, writing of the first layer, followed by rinsing of the unexposed PolyProtek with the protein. New layer spin coated with PolyProtek and a different protein, alignment to the first layer and writing of the second layer. Multiple protein patterns are obtained by repeated spin coating, alignment, writing and rinsing steps. All of the steps including electron beam lithography were performed outside the cleanroom. Protein structure from PDB#

1W4Y.

Figure 2-2. Dose test of PolyProtek on protein patterning. Fluorescence micrographs showing the effect of area dose on PolyProtek-HRP patterns. Doses range from 3-96 µC/cm2, therefore the patterns are visible only above a threshold dose. HRP concentration in spin-coating solutions was 10 µM. The patterns have been stained with Alexa Fluor 488 goat anti-HRP antibody.

The process of fabricating protein patterns by direct EBL is depicted in Figure 2-1. An aqueous solution containing the protein to be patterned and PolyProtek is first spin-coated onto silicon substrates, which is then processed by EBL. Horseradish peroxidase (HRP) was used as a model protein to investigate the optimal electron beam patterning properties for direct writing

(Figure 2-3). The spin-coated protein concentration and PolyProtek concentration was varied to

36 determine the highest signal-to-noise ratio (S/N) following patterning. The film thicknesses of spin-coated layers with varied concentration of PolyProtek or HRP concentration measured by ellipsometry are shown in Table 2-1. The effects of film thicknesses were more dependent on the

PolyProtek concentration rather than protein or HRP concentration. The optimal polymer concentration was 0.5 wt%. At lower concentration (0.3 wt%), a weak antibody signal was observed either because the polymer concentration was not high enough to stabilize the protein or because the starting film thickness was much smaller, suggesting much less protein present on the surface. Results also indicate that the protein was immobilized within the patterned features, and that PolyProtek helped preserve the antibody binding site of the proteins.

Figure 2-3. Proteins patterned by direct electron beam using PolyProtek as a resist. a)

Effect of HRP and PolyProtek concentration on S/N ratio of PolyProtek-HRP patterns. Top:

PolyProtek concentration was set to 0.5 wt% and bottom: HRP concentration was 10 µM; in each case S/N calculated for optimal dose is shown. b) Fluorescence micrographs of fluorescent

37 antibody-stained PolyProtek-IgG, PolyProtek-SAv, PolyProtek-GOx. Insets show that fluorescent antibodies do not bind non-specifically to PolyProtek patterns that did not contain respective proteins. Scale bars = 25 µm.

Table 2-1. Spin-coated thickness of varying concentrations of HRP and PolyProtek. Measured Thickness (Å) 0.3 wt% 0.5 wt% 1.0 wt%

PolyProtek PolyProtek PolyProtek 10 µM HRP 184.0 ± 1.8 238.8 ± 2.9 388.1 ± 8.7

0.1 µM HRP 1 µM HRP 10 µM HRP 100 µM HRP 0.5 wt% 230.0 ± 2.9 222.8 ± 2.3 218.1 ± 1.6 367.6 ± 12.7 PolyProtek

Glucose oxidase (GOx), immunoglobulin G (IgG) and streptavidin (SAv) were patterned to determine the ability of PolyProtek to stabilize a range of proteins (Figure 2-3b). Following fluorescent staining, the patterns exhibited high fluorescent signals (S/N > 120) for all of the patterned proteins. Non-specific binding of the fluorescent antibodies was not observed in patterns of PolyProtek only without protein (Figure 2-3b, insets). Therefore, the fluorescence signals seen in Figure 2-3b are due to specific antigen-antibody interactions. Successful staining of the protein patterns demonstrates that the recognition property was preserved after EBL processing, and that direct writing of proteins with PolyProtek is a promising method for generating protein patterns with high-resolution and alignment capabilities.

When four successive HRP patternings were applied on the same substrate, the fluorescence intensity of the first layer was comparable to the fourth layer (exposed to four electron beam lithography cycles, Figure 2-4a). This indicates that repeated exposure of the

PolyProtek-protein patterns to vacuum-vent and rinsing cycles did not affect protein integrity.

The protein patterns were also visualized by scanning electron microscopy (Figure 2-4b). Square

38 patterns with 10 µm and 500 nm side lengths as well as sub-100 nm dots and lines were achieved. Nano sized protein features were also observed by AFM as shown in Figure 2-4c. HRP was patterned with 110 nm line widths with 200 nm spacing before fluorescent antibody binding.

Because the nanometer features were arrayed close together, the overall fluorescence was observable (Figure 2-4c, left) demonstrating that the HRP in the nanometer features still bound antibody.

Figure 2-4. Multiple layers and nanopatterning. a) PolyProtek-HRP patterns (first and fourth layer, written on the same substrate are shown) and fluorescence intensity profiles drawn along second row of squares from the top. Inset shows that fluorescent antibodies do not bind non- specifically to PolyProtek patterns that did not contain respective proteins. Scale bars = 25 µm. b) Scanning electron micrographs showing micro- and nanopatterns of PolyProtek-HRP. Scale bars = 10 µm (top left), 2 µm (top right), 1 µm (bottom left and bottom right). c) HRP patterns with nanometer line spacing showing fluorescence, AFM and height profile. Scale bar = 5 µm and arrows indicate line width of 110 nm.

Patterning relevant proteins that are components in the extracellular matrix (ECM) are valuable for preparing bioactive surfaces in cell-material interaction applications and for understanding fundamental cell-ECM interactions. For example, Arnold and coworkers have shown that nanometer interfeature spacings of integrin-binding peptides are critical to cell adhesion.39 Thus, important ECM proteins, such as fibronectin, and signaling molecules such as bFGF and vascular endothelial growth factor (VEGF) were patterned with PolyProtek (Figure 2-

5). Particularly, bFGF is known for its remarkable instability.40,41 The ability to both stabilize as well as pattern and present bioactive signaling molecules as shown here has great potential.

Figure 2-5. Fibronectin, VEGF, and bFGF patterns. Fluorescent micrographs of generated patterns of fibronectin, VEGF, and bFGF with PolyProtek. The doses range from 5-35 µC/cm2.

Fibronectin patterns were stained with mouse anti-fibronectin, followed by Alexa Fluor 488 anti- mouse. VEGF stained with mouse anti-VEGF, and Alexa Fluor 594 anti-mouse. bFGF was stained with biotinylated anti-bFGF, and streptavidin Alexa Fluor 488.

Multicomponent nanopatterning is important for generating high-density, high-sensitivity arrays that could be very useful in fields such as proteomics, diagnostics, and biosensors. Such patterns could also be used to generate complex surfaces to investigate cell behavior and to control cell shape, for instance surfaces that present multiple integrin-binding proteins and growth factors to direct stem cell lineage or surfaces that present a gradient of a bioactive molecule.42-48 To investigate this capability, multiplexed protein arrays with nanometer-sized spacing and features were generated in close proximity by direct writing of different types of proteins including immunoglobulins, enzymes, and growth factors (Figures 2-6, 2-7, 2-8).

Fluorescent staining of the patterns and the corresponding AFM images showcase the nanoscale resolution patterning and alignment capabilities of this technique. Three different IgGs, human, chicken, and mouse, were patterned as triangles, circles, and squares, respectively, in an alternating configuration with the individual IgG shapes spaced 2 µm apart (Figure 2-6).

Multiprotein nanoarrays of HRP with chicken IgG (Figure 2-7) and VEGF with bFGF (Figure 2-

8) were also written. Each feature was composed of ~100 nm wide lines and the overall pattern in these arrays covered a dimension of 5 x 5 µm2. For HRP and chicken IgG, the individual elements comprising the protein pattern shape had a center-to-center distance of 400 nm, while the HRP and IgG shapes were separated by a distance of 3 µm for better visualization of the individual elements in the arrays. For VEGF and bFGF, the center-to-center distance of the individual elements was set to 250 nm, while each VEGF and bFGF pattern was separated by a distance of 500 nm. The data shows that indeed nanoarrays are possible. Such patterns could be very useful in generating bioactive surfaces for investigating fundamental questions about protein-protein interactions, as well as studying the effect of geometry on cell behavior or the effect of multivalency in cell behavior.

Figure 2-6. Multiple IgG nanopatterns. a) Fluorescent micrographs showing fluorescent antibody-stained individual IgG patterns with human IgG (triangles, green channel), chicken IgG

(circles, red channel) and mouse IgG (squares, blue channel) and merged. b) AFM of corresponding patterns with arrows indicating line width of 80 nm. Scale bars = 25 µm.

Figure 2-7. HRP and IgG nanopatterns. a) Fluorescent micrographs of antibody-stained HRP

(squares, green channel) and chicken IgG (circles, red channel) and merged image. b) AFM of corresponding HRP and chicken IgG stained patterns with arrows indicating line width of 110 nm. Scale bars = 25 µm.

Figure 2-8. Growth factors nanopatterns. a) Fluorescent micrographs of antibody-stained bFGF (hourglass shapes, green channel) and VEGF (diamonds, red channel) and merged images. b) AFM of corresponding bFGF and VEGF patterns with arrows indicating line width of 100 nm.

Scale bars = 25 µm.

2.3 Conclusions

In conclusion, the use of a polystyrene backbone trehalose glycopolymer, PolyProtek, has been shown to allow the direct patterning of bioactive proteins by providing stabilization to high- energy radiation and vacuum conditions during EBL. This was demonstrated through the direct patterning of various proteins and generating complexed, multicomponent micro- and nanopatterns of proteins. Although protein patterns were demonstrated, it is readily envisioned that this approach can be extended to other biomolecules, such as DNA, carbohydrates, and so on.

Thus, we expect that this biomolecule patterning technique will find broad application in different disciplines and will be important in designing bioanalytical assays, biosensors, microreactors, and bioactive interfaces for cell culture.

2.4 Materials and Methods

Materials

Silicon wafers were obtained from Cemat Silicon S.A., Poland. Trehalose was purchased from The Healthy Essential Management Corporation (Houston, TX). Sheep anti-glucose oxidase antibody was obtained from Abcam. Human, chicken, and mouse IgGs and the fluorescent antibodies Alexa Fluor 488 goat anti-HRP, Alexa Fluor 488 donkey anti-human IgG,

Alexa Fluor 594 donkey anti-chicken IgG were purchased from Jackson ImmunoResearch.

Alexa Fluor 350 donkey anti-mouse IgG, Alexa Fluor 488 donkey anti-rabbit IgG, Alexa Fluor

488 goat anti-sheep IgG were bought from Invitrogen. 2-O-(4-vinylbenzylidene)-α,α-trehalose and PolyProtek were prepared by following our literature procedure.38 All the other chemicals were obtained from Sigma-Aldrich and Fisher Scientific unless otherwise stated. For generating multicomponent patterns, silicon chips with gold alignment marks were fabricated by standard photolithography, metal evaporation and lift-off techniques.49

Effect of Polymer and Protein Concentration

Silicon substrates were cleaned with freshly prepared piranha solution (3:1 H2SO4/H2O2,

Caution! Piranha solution reacts violently with organic materials). After washing with Milli-Q water and drying under a stream of air, the cleaned substrates were spin-coated (500 rpm, 5 sec;

1000 rpm, 5 sec; 2000 rpm, 20 sec; or 4000 rpm, 10 sec) with a 30 µL solution containing

PolyProtek and HRP. To investigate the effect of protein concentration, 25 µL of PolyProtek (0.5 wt% in H2O) was mixed with 5 µL of protein solution (ranging from 0.60 to 600 µM in D-PBS) leading to final protein concentrations ranging from 0.1 to 100 µM. PolyProtek-HRP patterns were generated using a JC Nabity e-beam lithographic system (Nanometer Pattern Generation

System, version 9.0) modified from a JEOL JSM-6610 scanning electron microscope. An

44 accelerating voltage of 30 kV was used, with a beam current of ~15 pA, a spot size of 30-40 nm.

Square patterns (10 µm) were written at a dose range from 5 to 80 µC/cm2. After electron beam exposure, any non-crosslinked polymer on the chips was washed away with D-PBS containing 1 mM of Tween-20 (PBS-Tween); the surfaces were then stained with Alexa Fluor 488 goat anti-

HRP antibody (10 µg/mL in PBS-Tween).

To investigate the effect of polymer concentration, 25 µL of PolyProtek (0.3-2 wt% in

H2O) was mixed with 5 µL of protein solution (60 µM in D-PBS) leading to a final protein concentration of 10 µM. Patterns were written at doses ranging from 3 to 96 µC/cm2 and stained as described above.

Patterning of Multiple Proteins and Immunostaining

Silicon substrates with gold alignment marks were piranha cleaned and spin-coated with a 30 µL solution of PolyProtek (25 µL, 0.5 wt% in H2O) and protein (5 µL of protein 30 µM in

D-PBS) as described above. After the first pattern had been written with human IgG, the substrates were washed with wash buffer followed by a very brief rinsing with Milli-Q water.

After alignment to the first layer, a complementary pattern was written with chicken IgG. The third layer was written with mouse IgG. The generated patterns were visualized by immunofluorescence staining. The substrates were sequentially incubated with fluorescently labeled donkey anti-human, donkey anti-chicken and donkey anti-mouse IgGs (30 min, 10

µg/mL in PBS-Tween). In between the incubation steps, the substrates were washed with wash buffer.

HRP and chicken IgG multi-patterned substrates were prepared similarly as described above. In brief, the alignment substrates were spin-coated with a solution containing 25 µL of

0.5 wt% PolyProtek in H2O and 5 µL of 60 µM HRP or IgG in D-PBS. The first layer patterned

45 was HRP followed by chicken IgG with washes in between the patterning and spin-coating steps.

For visualization by immunofluorescence, the patterns were sequentially stained with Alexa

Fluor 488 anti-HRP and Alexa Fluor 594 anti-chicken IgGs (30 min, 10 µg/mL in PBS-Tween) with washes in between the incubation steps. VEGF and bFGF multi-patterned substrates were prepared similarly by first spin-coating and patterning VEGF followed by bFGF. The spin- coating solutions contained 25 µL of 0.5 wt% PolyProtek in H2O and 5 µL of a 2 mg/mL VEGF or bFGF solution in D-PBS. After patterning, the substrates were sequentially incubated with biotinylated anti-bFGF, mouse anti-VEGF, SAv-conjugated Alexa Fluor 488 and Alexa Fluor

594 anti-mouse IgG (30 min, 10 µg/mL in PBS-Tween) and washed in between incubation steps.

Fluorescence Microscopy

The immunostained patterns were visualized by fluorescence microscopy using a Zeiss

Axiovert 200 fluorescent microscope equipped with an AxioCam MRm monochrome camera, and pictures were acquired and processed using AxioVision LE 4.6 software. NIH ImageJ software was used to calculate the S/N ratio as (signal - background)/(standard deviation of background).

Atomic Force Microscopy

Height characterization of patterns was imaged on a Bruker Dimension Icon AFM using

Peak Force tapping mode with ScanAsyst Air probes. AFM imaging was performed on a scan size of 25.0 µm, with a scan rate of 0.7 Hz and 512 samples per line.

Film Thickness Measurements

Film thicknesses from spin-coating solutions of 5 µL HRP (60 µM in D-PBS) and 25 µL

PolyProtek (0.3, 0.5, or 1 wt%) were measured. In addition, 0.5 wt% of PolyProtek and 0.1-100

µM HRP were evaluated. Ellipsometry was performed using a Gaertner LSE ellipsometer equipped with a 633 nm HeNe laser fired at a 70° incidence angle. The silicon oxide on the piranha-cleaned silicon wafer was measured and fitted using the refractive index of Palik (n1 =

0.54264, k1 = 0.00) and silicon as substrate (n1 = 3.589, k1 = 0.016). The measurement was repeated on the same wafer after spin-coating the HRP and PolyProtek solution. The subsequent protein and polymer layer was fitted using values for the previously obtained silicon oxide thickness and an additional Cauchy layer model (n1 = 1.45, k1 = 0.01). A minimum of 15 measurements were performed at three different locations and the values were then averaged.

Acknowledgements

This work was funded by the National Science Foundation (CHE-1112550). E.B. thanks

The Netherlands Organization for Scientific Research and Marie Curie Cofund Action for the financial support (Rubicon Grant 680-50-1101). J.L. thanks the NIH for Biotechnology Training

Fellowship (5T32 GM67555-7). The scanning electron microscope was purchased with funds from the California NanoSystems Institute.

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CHAPTER 3

DIRECT WRITE PROTEIN PATTERNS FOR MULTIPLEXED CYTOKINE DETECTION FROM LIVE CELLS USING ELECTRON BEAM LITHOGRAPHY

This chapter is an edited version of a paper published as: Lau, U. Y.; Saxer, S. S.; Lee, J.; Bat, E.; Maynard, H. D., “Direct Write Protein Patterns for Multiplexed Cytokine Detection from Live Cells Using Electron Beam Lithography,” ACS Nano 2015, Article ASAP.

3.1 Introduction

Extracellular signaling molecules are secreted by cells in response to a variety of factors, including temperature, addition of drugs, chemicals, and other changes in the surrounding environment. Therefore, knowing what types of molecules and the amounts secreted would provide important insight and information on the functional state of the cell. For example, interleukin-6 (IL-6) and tumor necrosis factor alpha (TNFα) are common cell signaling proteins

(cytokines) that influence many aspects of the immune and inflammatory response. These molecules are also important biomarkers; elevated IL-6 or TNFα serum levels are associated with various diseases, such as rheumatoid arthritis and prostate cancer.1,2 Thus, detection of these molecules secreted from cells is an important undertaking.

Currently available and commonly used methods for cytokine detection include intracellular cytokine cytometry (ICC) and enzyme-linked immunospot (ELISPOT) assays.3 ICC involves inhibiting stimulated cells from secreting cytokines, and then allowing labeled anti- cytokine antibodies to permeate into the cells. This is coupled with flow cytometry analysis. In the ELISPOT assay, cells are plated directly into wells coated with anti-cytokine antibodies, and the released cytokines are visualized by an enzyme-labeled detection antibody, similar to standard ELISA procedures. An important contribution towards improving biosensor technology aims to miniaturize sample volumes and for inline detection from live cells, where micro- and nanofabrication approaches become highly desired. Benefits from miniaturization are better sensitivities and high-throughput analyses. Examples of developing technologies for the detection of secreted cytokines from living cells include a microfluidics platform for immobilizing antibodies,4,5 photolithography to fabricate gold electrodes for modification with aptamers,6 and nanoimprint lithography to generate structures for cytokine biosensing.7

There are a variety of patterning techniques that have been used for biomolecule immobilization with micro- and nanoscale features.8-10 Common techniques include ink-jet printing,11 stamping techniques such as microcontact printing12,13 and nanoimprint lithography,14,15 scanning probe microscopy techniques such as dip-pen nanolithography

(DPN),16,17 polymer pen lithography (PPL),18 and electron beam lithography (EBL).19-21

Expectedly, each technique has advantages and limitations. Stamping techniques allow for the rapid generation of patterns, but require template fabrication and preparing dense patterns with different biomolecules can be difficult. DPN and PPL enable direct write methods with nanoscale spacing between features, but are limited in that most often they are serial processes.

EBL is also a serial process and suffers from significantly longer exposure times; however it allows for the generation of user-designed patterns with high resolution and nanoscale alignment capability, allowing for multiplexing and high density patterns.22

Direct writing of proteins by EBL presents a challenge due to the nature of the harsh processing conditions. EBL requires high vacuum and involves high energy irradiation with electrons for patterning, which are not ideal for maintaining protein bioactivity. Recently, we have reported that polymers with trehalose side chains (PolyProtek) are able to stabilize various proteins against environmental stresses such as heat and lyophilization.23-25 We have also shown the utilization of PolyProtek as a novel aqueous-based resist material that enables the direct writing of proteins by EBL (covered in Chapter 2).26 PolyProtek was shown to stabilize proteins under high vacuum and direct exposure to high energy irradiation. Another material, silk, was also used as a positive or negative resist for protein patterning with EBL as shown by Kim et al.27 Herein, we demonstrate the ability to use PolyProtek for direct write EBL of IL-6 or TNFα antibodies for capture of these important biomolecules secreted from stimulated macrophages.

The patterns were visualized via dark field microscopy exploiting the surface plasmon resonance of silver enhanced gold nanoparticle secondary antibodies.

3.2 Results and Discussion

The amount of non-specific adsorption is an important consideration for any biosensor application because background adsorption increases noise and decreases signal thus reducing sensitivity.28 Surfaces were therefore passivated to prevent non-specific adsorption and ensure specific recognition. This was achieved by the single-step surface modification of grafting poly(ethylene glycol) (PEG)-silane onto silicon surfaces. This passivation strategy allows for ultrathin and stable protein-repellent PEG monolayers.29 The surface immunoassay described in this section employs gold nanoparticle-labeled analytes for detection. Gold nanoparticles generate a localized surface plasmon resonance signal that can be simply visualized using a dark field optical microscope. Reducing the non-specific adsorption of gold nanoparticles and primary antibodies helps eliminate background noise and thus improve signal and sensitivity. The amount of non-specifically adsorbed neutravidin-conjugated gold nanoparticles was evaluated for plain silicon substrates or silicon coated with PEG-silane of different molecular weights (Figure 3-1).

All surfaces were incubated with 30 nm neutravidin-conjugated gold nanoparticle solutions for

18 hours. Compared to the piranha-cleaned silicon substrates (Figure 3-1a), the PEG 400 coating also had considerable fouling (Figure 3-1b) while PEG 2000 showed a slight reduction in protein adsorption (Figure 3-1c). A PEG-silane of molecular weight 5 kDa was required to significantly reduce non-specific adsorption to barely detectable levels (Figure 3-1d). Specifically, a 95% reduction in protein adsorption was observed compared to the bare silicon substrate. Therefore, the 5 kDa PEG-silane coating was utilized in all subsequent experiments.

Figure 3-1. Non-specific adsorption of neutravidin-conjugated 30 nm gold nanoparticles to treated silicon substrates: a) bare silicon, b) PEG 400-silane, c) PEG 2000-silane, and d) PEG

5000-silane.

The process of fabricating antibody patterns by direct electron beam lithography is depicted in Figure 3-2. PEG-silane coated silicon substrates were spin-coated with an aqueous solution containing the antibody, 0.5% wt/vol PolyProtek, and 1 mM ascorbic acid. The resulting thickness for PEG-silane was 26.8 ± 1.1 Å, and the spin-coated antibody layer was 205.3 ± 1.9

Å, as measured by ellipsometry. When irradiated by electrons, the trehalose polymer cross-links to the PEG-coated silicon substrate likely by a mechanism similarly observed for other polymers.30,31 The cross-linking results in a trehalose-based hydrogel, which helps to stabilize and immobilize the protein. Ascorbic acid is a known radical scavenger,32 and therefore can help protect proteins from e-beam radiation damage.26 We verified this in our system by patterning without or with 1 and 2 mM ascorbic acid and found that 1 mM ascorbic acid provided the highest signal by dark field microscopy (Figure 3-3). Hereafter, 1 mM ascorbic acid was added to the resist/protein solutions. For multiplexed antibody patterns, the chips were fabricated by re-

56 subjecting the substrate to another round of e-beam processing of spin-coating with a different antibody, aligning, and writing steps.

Figure 3-2. Direct electron beam patterning process for generating antibody patterns. An aqueous solution containing PolyProtek, ascorbic acid, and target antibody is spin-coated on a

PEG-silane coated substrate and subsequently patterned by electron beam lithography. Non- crosslinked areas are washed away. Multicomponent antibody patterns can be generated by spin- coating a new layer with a different antibody, alignment and subsequent e-beam patterning and rinsing.

Figure 3-3. Effect of ascorbic acid on anti-TNFα patterning and immunoassay signals. Anti-

TNFα patterned with area dose of 25 µC/cm2 with 0.5% wt/vol PolyProtek and ascorbic acid concentration of a) 0 mM, b) 1 mM, and c) 2 mM. Scale bars = 35 µm.

The direct electron beam writing and assay of antibodies was first evaluated using human immunoglobulin G (IgG) to optimize various parameters and components of the immunoassay system. Silicon wafers were spin-coated with a solution containing 0.5% wt/vol PolyProtek, 1 mM ascorbic acid, and 1 mg/mL human IgG. The human IgG coated surfaces were patterned in an “A1/Circle” shape (Figure 3-4c) with an area dose of 25 µC/cm2 and after processing, the chips were sequentially incubated with biotinylated anti-human IgG and neutravidin-conjugated

30 nm gold nanoparticles, followed by silver enhancement. Dark field microscopy images were taken from a reference pattern without neutravidin-conjugated gold nanoparticles (Figure 3-4a), with bound gold nanoparticles (Figure 3-4b) and the silver enhanced gold nanoparticles pattern

(Figure 3-4c). The localized surface plasmon resonance (LSPR) signal was observed for the gold nanoparticle sample; however silver enhancement greatly increased signal and thus sensitivity, and this process was utilized for all further studies.

Figure 3-4. Dark field micrographs for the sandwich assay of human IgG patterns. IgG patterns were incubated with biotin anti-human IgG, neutravidin-conjugated gold nanoparticles, and

58 silver enhanced. Patterns are shown a) before gold nanoparticle incubation, b) after gold nanoparticle incubation, and c) after silver enhancement. Scale bars = 35 µm.

Cytokines IL-6 and TNFα were chosen for multiplexed detection by the surface immunoassay because these secreted cell signaling molecules are commonly associated with macrophage activation. Cultured macrophages readily secrete the signaling proteins to regulate the immune response upon stimulation. RAW 264.7 mouse macrophages were stimulated with lipopolysaccharide (LPS), a component of the outer membrane in Gram-negative bacteria, to secrete IL-6, TNFα, and many other cytokines into the cell culture media. An enzyme-linked immunosorbent assay (ELISA) was performed to detect the presence of IL-6 and TNFα in media

(Figure 3-5). Importantly, the results show that there are undetectable levels of cytokines from unstimulated macrophages and that the secondary antibodies have high specificity for the target antibody and no cross-reactivity. These properties are necessary for translating the components towards a multiplex surface immunoassay system.

Figure 3-5. ELISA detection of IL-6 and TNFα from LPS-stimulated RAW 264.7 macrophages

(5 × 105 cells). IL-6 × TNFα represents anti-IL-6 capture antibody and biotinylated anti-TNFα antibody, and TNFα × IL-6 represents anti-TNFα capture antibody and biotinylated anti-IL-6 antibody. Media represents the amount of detected cytokines from unstimulated macrophages.

Electron beam lithography was first used for direct writing of anti-IL-6 and anti-TNFα antibody patterns on separately patterned chips. The antibody patterns were sequentially incubated with undiluted cell culture media from RAW 264.7 macrophages that were stimulated with LPS for 24 hours. Then, either biotinylated anti-IL-6 or biotinylated anti-TNFα was added, followed by 30 nm neutravidin-conjugated gold nanoparticles and the silver enhancement step.

The signal depended on the concentration of antibody used during spin-coating. The concentration of antibody tested ranged from 1 to 5 µM, with 5 µM resulting in the highest observed signal (Figure 3-6). Not surprisingly, with increased antibody concentrations, there was an increase in signal, which is likely related to a higher density of bioactive antibodies following electron beam processing. Both TNFα and IL-6 capture was observed by dark field microscopy with silver-enhanced gold nanoparticles bound to micro sized patterns of defined texts and shapes (Figure 3-7a, b). The gold nanoparticles were visualized on submicron patterns with scanning electron microscopy, where the gold antibodies were observed as dots on 500x500 nm- sized squares (Figure 3-7c). Furthermore, nanopatterns were fabricated. Anti-TNFα nanopatterns with line widths of 150-180 nm were observed by atomic force microscopy (AFM, Figure 3-7e, f). The line patterns were fabricated in concentric shapes that were visualized by dark field microscopy as squares and circles (Figure 3-7d) demonstrating that nanopatterns of active anti-

TNFα were achieved. These results confirm that the immunoassay following the direct write of antibodies by EBL can be used for the detection of cytokines in complex cell culture media.

These patterns of shapes, letters, and numbers demonstrate the capability to generate arbitrary patterns with high resolution.

Figure 3-6. The effect of anti-TNFα spin-coating concentration on immunoassay signals.

Concentrations of anti-TNFα used for spin-coating at a) 1 µM, b) 2 µM, and c) 5 µM. Scale bars

= 35 µm.

Figure 3-7. Cytokine detection with a bound silver enhanced gold nanoparticles immunoassay on surface immobilized micro- and nanopatterns. Two cytokines, a) TNFα and b) IL-6, were detected from cell media of LPS-stimulated RAW 264.7 macrophages, scale bars = 35 µm. c) electron micrograph of anti-TNFα submicron patterns, scale bar = 1 µm. Anti-TNFα nanopatterns showing d) dark field micrograph cytokine detection immunoassay of circles and

61 squares with nanometer line widths, scale bar = 20 µm, e) AFM of one of the circle patterns with

150 nm line widths, and f) AFM of one of the square patterns with 180 nm line widths indicated by the arrows.

Sensitivity to cytokine concentrations and detection limit are both important considerations for immunoassays. The levels of TNFα and IL-6 in LPS-stimulated macrophage culture media were quantified by ELISA and used as a benchmark. From 5 × 105 cultured macrophages, the amount of TNFα and IL-6 detected was 43 ng/mL and 53 ng/mL, respectively.

Patterns of anti-TNFα and anti-IL-6 were incubated in varying dilutions ranging from 200 ng/mL to 0 of TNFα or IL-6 from cell media of stimulated macrophages and further developed. Figure

3-8 and 3-9 shows the sensitivity of the immunoassay to TNFα and IL-6, respectively. Signals were strongly observed at TNFα and IL-6 concentrations of 200 ng/mL and 40 ng/mL, while detection was possible above 5 pg/mL for TNFα and 50 pg/mL for IL-6. This is comparable to the detection limit of ELISA, which from the manufacturer’s kit (eBioscience) has a standard curve range of 4-500 pg/mL.

Figure 3-8. Detection sensitivity of anti-TNFα patterns to varying concentrations of TNFα in media: a) 200 ng/mL, b) 40 ng/mL, c) 500 pg/mL, d) 50 pg/mL, e) 5 pg/mL, and f) 0. Scale bars

= 20 µm.

Figure 3-9. Detection sensitivity of anti-IL-6 patterns to varying concentrations of IL-6 in media: a) 200 ng/mL, b) 50 ng/mL, c) 500 pg/mL, d) 50 pg/mL, e) 5 pg/mL, and f) 0. Scale bars

= 20 µm.

Multiplexed antibody patterns to detect both IL-6 and TNFα were then investigated. First, anti-TNFα patterns were generated and the process of spin-coating with anti-TNFα, writing, and rinsing on the same substrate was repeated four times. The signal-to-noise of the patterns was comparable between the four processing and exposure cycles (Figure 3-10) suggesting that the process is amenable to multiplex patterning. Taking advantage of the alignment capabilities of electron beam lithography, anti-IL-6 and anti-TNFα patterns of “TNFα” and “IL-6” were generated in close proximity. Following the direct writing of anti-TNFα, another round of patterning was performed with anti-IL-6 to achieve multicomponent antibody patterns. The detection of both IL-6 and TNFα was observed by visualization of their associated patterns

(Figure 3-11a). The surface immunoassay also demonstrates high specificity towards both cytokines, with slight cross-reactivity against IL-6. The corresponding patterns were visible when stained for only TNFα (Figure 3-11b) or for only IL-6 (Figure 3-11c).

Figure 3-10. Repeated processing and exposure cycles of anti-TNFα patterned on the same substrate. The same substrate was subjected to four cycles of spin-coating with anti-TNFα antibody, e-beam patterning, and rinsing. The chip was then developed with the immunoassay.

Dark field micrographs show the pattern from the a) first cycle, b) second cycle, c) third cycle, and d) fourth cycle. The signal-to-noise of the square in the first cycle was 20.9 and in the fourth cycle 23.4 calculated as (signal from the box – signal of the background) / (standard deviation of the background). Scale bars = 35 µm.

Figure 3-11. Dark field micrographs of multiple antibody patterning and cytokine detection of

IL-6 and TNFα. The generated patterns were stained to detect for a) both IL-6 and TNFα, or selectively for: b) TNFα, and c) IL-6. Scale bars = 35 µm.

Finally, the multiplexed patterns were used to monitor IL-6 and TNFα secretion over time. Multicomponent IL-6 and TNFα patterns were directly incubated with RAW 264.7 macrophages. Upon addition of LPS to the cultured macrophages, the patterned chips (with

“TNFα”/B2square shape for TNFα and “IL-6”/A1circle shape for IL-6) were then removed from cell cultures at various time points over the course of 24 hours. Subsequent development of the immunoassay showed an expected increase in LSPR signals and thus developing visualization of the patterns incubated from 2 to 24 hours (Figure 3-12). This steady increase in IL-6 and TNFα secretion over 24 hours is in agreement with levels reported in literature.33

Figure 3-12. Multiplexed detection of IL-6 and TNFα from stimulated RAW 264.7 macrophages over time. Chips were directly incubated in LPS-stimulated macrophages at time points: a) 2 hr, b) 12 hr, and c) 24 hr. Scale bars = 35 µm. 65

The detection of both cytokines, as well as the capability to selectively detect one specific signaling protein from cellular milieu, is essential for a multiplexed immunoassay system. Using a trehalose glycopolymer, the antibodies retained their binding activity even after subjection to repeated electron beam processing conditions, such as vacuum and rinsing cycles, exemplifying the use of PolyProtek as a stabilizing resist. Furthermore, low amounts of non-specific adsorption were observed on the e-beam generated patterns. Specifically, the amount of the gold nanoparticles bound to the opposing cytokine pattern was greatly reduced (Figure 3-11b, c). The direct write assay was capable of detection as low as cytokine concentrations of 5 pg/mL and could detect increasing concentrations of the secreted proteins over time from live cells. The combination of all these properties demonstrates that multiplexed patterns can be successfully patterned with this technique, and that the detection of two different cytokines in relevant cell culture conditions on these patterns is possible. Electron beam lithography is a serial process, but there are several new techniques in development. For example, stencil masks or character projection technology allow for generation of complex shapes with one exposure, and multiple electron beams for parallel processes are being investigated.34,35 The process we describe herein should also be applicable to these technologies.

3.3 Conclusions

We have successfully demonstrated a cytokine detection method by directly patterning antibodies onto surfaces utilizing PolyProtek and EBL. The patterned IL-6 and TNFα antibodies retained their recognition properties, allowing for binding to the respective cytokines released from LPS-stimulated macrophages. Capture of the cytokines was successfully detected via a sandwich immunoassay visualized from LSPR of silver-enhanced gold nanoparticles bound to

66 the patterns. Multiplexed cytokine detection was also shown to specifically visualize IL-6 and

TNFα concentrations in relevant cell culture conditions. These results demonstrate the potential of this patterning technique for developing biosensors and diagnostic assays.

3.4 Materials and Methods

Materials

5 kDa mPEG-silane was purchased from Creative PEGWorks. Silicon wafers were purchased from Cemat Silicon S.A., Poland. Human IgG and Alexa Fluor 488 donkey anti- human IgG were purchased from Jackson ImmunoResearch. Rat anti-mouse IL-6 and biotinylated rat anti-mouse IL-6 IgGs were purchased from eBioscience. Rabbit anti-mouse

TNFα IgG was purchased from Thermo Fisher Scientific, and biotinylated rabbit anti-mouse

TNFα IgG was from Invitrogen. Antibodies for patterning were concentrated with Centriprep

MWCO 100 kDa prior to use. Mouse IL-6 and mouse TNF-alpha ELISA Ready-SET-Go!® reagent sets were purchased from eBioscience. Neutravidin-conjugated 30 nm gold nanoparticles was purchased from Nanopartz. LI Silver Enhancement Kit was purchased from Molecular

Probes. Lipopolysaccharides from Escherichia coli (055:B5) were from Sigma-Aldrich. RAW

264.7 murine macrophage cells were kindly provided by Professor Tian Xia from UCLA.

Synthesis of Trehalose Glycopolymer (PolyProtek)

The synthesis of a styrenyl ether-based trehalose glycopolymer was modified from a previously reported procedure.24 Styrenyl ether trehalose monomer (375.2 mg, 8.18 x 10-1 mmol)

-2 and AIBN (3.13 mg, 1.91 x 10 mmol) were dissolved in H2O (2.73 mL) and DMF (1.36 mL), respectively. Both solutions were added to a reaction flask and subjected to five cycles of freeze- pump-thawing. The polymerization was started by immersing the flask in a 75 °C oil bath. After

8.33 h, the polymerization was stopped exposing the solution to oxygen and cooling with liquid nitrogen. Residual monomer was removed by dialysis against H2O (MWCO 3,500 g/mol) for 3 days and lyophilized to obtain a white powder, with number average molecular weight Mn (GPC)

= 15.7 kDa, and molecular weight dispersity Ð = 3.25.

PEG-silane Coating of Si Chips

Silicon substrates were cleaned by immersing into freshly prepared piranha solution (3:1

H2SO4 to 30% H2O2) and heated at 70 °C for 15 min. The chips were extensively rinsed with

Milli-Q water and dried under a stream of filtered air. The cleaned chips were immediately incubated in a 1% wt/vol mPEG-silane solution in anhydrous toluene for at least 24 hours. The

PEG-silane coated chips were then rinsed with methanol, followed by a large excess of Milli-Q water, and then dried under a stream of air. The substrates were immediately used for subsequent patterning experiments.

Film Thickness Measurements

Film thicknesses from the PEG-silane layer and spin-coated solutions of antibody layers were measured using a Gaertner LSE ellipsometer equipped with a 633 nm HeNe laser fired at a

70° incidence angle. The silicon oxide on the piranha-cleaned silicon wafer was measured and fitted using the refractive index of Palik (n1 = 0.54264, k1 = 0.00) and silicon as substrate (n1 =

3.589, k1 = 0.016). The measurement was repeated on the same sample after PEG-silane coating and spin-coating the protein and PolyProtek solution. The subsequent protein and polymer layer was fitted using values for the previously obtained silicon oxide thickness and an additional

Cauchy layer model (n1 = 1.45, k1 = 0.01). A minimum of 15 measurements were performed at three different locations and the values were then averaged.

Electron Beam Lithography

Silicon substrates were spin-coated using Spin Coater Model ACE-200 (Dong-Ah).

Aqueous solutions were spin-coated at 500 rpm for 5 sec, ramped to 1000 rpm for 5 sec, then ramped to 2000 rpm for 20 sec, and finally to 4000 rpm for 10 sec. PEG-silane coated silicon substrates were first spin-coated with Milli-Q H2O. Then, the substrates were spin-coated with a solution comprised of anti-IL-6 or anti-TNFα antibody, 0.5% wt/vol styrenyl ether-based trehalose glycopolymer, and 1 mM L-ascorbic acid in H2O. Patterns for electron beam lithography were designed in DesignCAD Express 16 software, and were generated using JC

Nabity Lithography System (Nanometer Pattern Generation System, Ver. 9.0) modified from a

JEOL JSM-6610 scanning electron microscope. An accelerating voltage of 30 kV, a spot size of

34 nm, and a beam current of 15 pA were used (dosage 25 µC/cm2). Following electron beam irradiation, any non-crosslinked polymer was rinsed away with wash buffer (0.05% Tween-20 in

D-PBS). Alignment silicon wafers were fabricated via standard photolithography, metal evaporation and lift-off techniques as previously described.19 To generate multicomponent protein patterns, the second antibody was spin-coated onto the same substrate. The chips were aligned by the prefabricated gold features, and patterned in close proximity to the first antibody.

Non-crosslinked polymer was removed by rinsing with wash buffer.

Atomic Force Microscopy

AFM characterization of patterns was performed on a Bruker Dimension Icon AFM using

Peak Force tapping mode with ScanAsyst Air probes. AFM imaging was performed on a scan size of 25 µm with a scan rate of 0.7 Hz and 512 samples per line.

Stimulation of RAW 264.7 Macrophages

RAW 264.7 murine macrophages were cultured in RPMI 1640 medium supplemented with 10% fetal bovine serum (FBS) and 1% penicillin-streptomycin. The cells were seeded in 6-

5 well plates at a density of 5 × 10 cells per well and incubated at 37 °C and 5% CO2. After 24 hours, the culture media was replaced with 10 µg/mL LPS in the supplemented RPMI 1640 media. After incubated for 24 hours, the cell culture media was collected and centrifuged at

1,500 rpm for 5 min at 4 °C. Aliquots of the supernatant were stored at -80 °C until use.

Surface Immunoassay

Following electron beam lithography, the antibody-patterned surfaces were incubated in cytokine-containing media for 2 hours. The chips were then rinsed with wash buffer for 5 min, followed by incubation with biotinylated anti-IL-6 or anti-TNFα (5 µg/mL in D-PBS), or both, for 2 hours. After rinsing with washing buffer, neutravidin-conjugated 30 nm gold nanoparticles

(1:100 in D-PBS) were allowed to bind to the patterns. The chips were rinsed again, and the gold nanoparticles were silver-enhanced, following the procedure as described by the manufacturer, for 5 min prior to a final rinsing step. The developed substrates and patterns were then imaged under aqueous conditions (D-PBS) using an Olympus BX51 microscope equipped with

QImaging Retiga-2000R camera under dark field.

Measurement of Cytokine Detection by ELISA

Anti-IL-6 or anti-TNFα antibodies (2 µg/mL in D-PBS) were incubated in 96-well plates for 16 hours at 23 °C. The plate was washed between each step with wash buffer. The plate was then blocked with 1% BSA in D-PBS for 2 hours at 23 °C. Supernatant from RAW 264.7 cell culture media was added to the wells and incubated for 2 hours. Biotin anti-IL-6 or biotin anti-

TNFα conjugate antibodies (0.25 µg/mL in 1% BSA D-PBS) were added and incubated for 2

70 hours, followed by incubation with streptavidin-HRP (1:200 in 1% BSA D-PBS) for 20 min. The plate was developed with 1-Step™ Ultra TMB solution (Pierce Biotechnology, Rockford) for 7 min prior to adding 1 M H2SO4 as the stop solution. The absorbance signals were measured at

450 nm and the background at 630 nm was subtracted.

Cytokine Detection Sensitivity and Time Studies

Concentrated solutions of IL-6 and TNFα from LPS-simulated macrophage cell culture media were quantified using ELISA. Serial dilutions of the cell culture media in D-PBS were prepared with the cytokine concentrations at 200 ng/mL, 50 ng/mL, 500 pg/mL, 50 pg/mL, and 5 pg/mL. The dilutions were used to incubate onto anti-IL-6 and anti-TNFα patterned surfaces.

Subsequent processing of the chips with biotinylated antibodies, neutravidin-conjugated gold nanoparticles and the silver enhancement steps were performed and imaged under dark field as described above.

For detection of IL-6 and TNFα secretion over time, multi-antibody patterns were first fabricated and stored in D-PBS until use. RAW 264.7 macrophages were seeded in 6-well plates at a density of 5 × 105 cells per well and incubated for 24 hours. Then, the culture media was replaced with 10 µg/mL LPS in the working media. Prepared chips were incubated in the wells following LPS addition at the time points: 2, 12, 18, and 24 h at 37 °C for 30 min. After rinsing, the immunoassay was continued as described above.

Conflict of Interest

The authors declare that a provisional patent application has been filed on the use of

PolyProtek as a resist for EBL.

Acknowledgements

This work was supported by funds from the Center for Scalable and Integrated

NanoManufacturing (SINAM, CMMI-0751621). The authors thank Professor Yu Huang

(UCLA) for the use of her optical microscope. U.Y.L. thanks the NIH Chemistry-Biology

Interface Training Program for support (T32 GM 008496) for funding. S.S. thanks the Swiss

National Science Foundation for the fellowship (PBEZP2-133211). E.B. thanks The Netherlands

Organization for Scientific Research and Marie Curie Cofund Action for the financial support

(Rubicon Grant 680-50-1101).

3.5 References

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CHAPTER 4

ENHANCED STABILITY OF GRANULOCYTE COLONY- STIMULATING FACTOR USING TREHALOSE GLYCOPOLYMERS

The chapter contains portions of an edited version of a paper published as: Lee, J.; Lin, E.-W.; Lau, U. Y.; Hedrick, J. L.; Bat, E.; Maynard, H. D., “Trehalose Glycopolymers as Excipients for Protein Stabilization,” Biomacromolecules 2013, 14, 2561-2569.

4.1 Introduction

Granulocyte colony-stimulating factor (G-CSF) is a growth factor that helps regulate the development of neutrophils by regulating the production of neutrophils within the bone marrow and stimulating neutrophil progenitor proliferation, differentiation, and end-cell functional activation.1,2 Therapeutically, G-CSF is used to stimulate bone marrow precursors to form neutrophils. In 2001, a recombinant methionyl human G-CSF, marketed as Neupogen®, was approved for clinical use to fight infection in patients undergoing chemotherapy.3,4 This 18.8 kDa protein is produced by recombinant DNA technology in E. coli and contains 175 amino acids.5 The use of G-CSF has later been approved for aplastic anemia, severe congenital neutropenia, bone marrow transplantation procedures, myelodysplastic syndromes, and for support of patients with acquired immune deficiency syndrome (AIDS). Cancer patients undergoing chemotherapy are at a particularly high risk to develop low neutrophil levels, leading to an increased risk of febrile neutropenia accompanied by the possibility of sepsis and eventual organ failure.

However, G-CSF (Neupogen®) has a relatively short serum half-life between 3.5 and 3.8 hours.3 Due to the rapid clearance from the body, a daily dosing regimen is necessary for treatment. PEGylation, or the covalent attachment of poly(ethylene glycol) (PEG) to a protein of interest, is a well-known technique to improve biological half-life of proteins in vivo.6

Conjugation of PEG increases the half-life by decreasing kidney filtration of the higher molecular weight conjugate, as well as sterically hindering the approach of proteases and antibodies. The covalent conjugate of G-CSF and PEG, sold by Amgen as Neulasta®, demonstrates superior pharmacokinetics compared to the un-PEGylated form and is widely used in the clinic with a combined revenue over 4.4 billion dollars each year.7-9 The PEGylated G-

CSF protein has a prolonged circulation serum half-life of 42 hours and only requires a single administration per chemotherapy cycle. However, the attachment of PEG does not increase the drug stability to environmental stresses. Neulasta® must be stored at refrigerated temperatures, and freeze-thaw, neutral pH and room temperature conditions must be avoided to prevent protein unfolding and loss of activity.10,11

Previous work by our group (Maynard and co-workers) with trehalose glycopolymers showed that glycopolymer protein conjugates and trehalose glycopolymers as an additive significantly increases the resistance of proteins to denaturation during lyophilization and heat stresses relative to the native protein.12,13 Glycopolymer conjugates of lysozyme were prepared and exhibited enhanced stability compared to wild-type lysozyme or the addition of equivalent concentrations of trehalose, PEG, or glycopolymers excipients.12 The activities of other enzymes, including horseradish peroxidase (HRP) and β-galactosidase (β-gal), were also retained against environmental stresses with the glycopolymers as excipients.13 PEGylation of protein therapeutics increases the molecular weight, which reduces kidney clearance and increases circulation lifetime.14,15 Similar to PEGylation, conjugation of trehalose polymers is expected to sterically shield a protein of interest from proteolysis or clearance. Kidney filtration has been shown to be a major mechanism regulating G-CSF bioavailability in vivo, and PEGylation of G-

CSF greatly reduced kidney clearance because of its increased size.16 Therefore, conjugates of trehalose glycopolymer and G-CSF should to be significantly more stable against environmental stresses, with the additional benefit of enhanced pharmacokinetic properties. The glycopolymer

G-CSF conjugate would enable a heat stable, bioactive protein therapeutic with increased in vivo half-life.

Neulasta® contains a 20 kDa linear aldehyde end functionalized monomethoxy-PEG attached via reductive amination to the N-terminal methionine residue of the G-CSF protein.

This site-specific conjugation preserved the highest activity following modification, yielding N- terminal PEGylated conjugates with 68% activity relative to the native G-CSF.17 We aimed to synthesize a trehalose glycopolymer G-CSF conjugate using a similar strategy in order to directly compare to the PEGylated therapeutic and to minimize the decrease in bioactivity of G-

CSF.

The work presented in this chapter demonstrates that the trehalose polymer can be applied to enhance the therapeutic potential of the clinically used protein, G-CSF, and shows promising results that prompt future in vitro and in vivo work. First, recombinant G-CSF in E. coli was isolated through construction of the plasmid and subsequent expression and purification steps. Various trehalose polymers were synthesized and screened to evaluate their cytotoxicity in various cell lines, and their ability to stabilize G-CSF against heat stress was assessed. Finally, trehalose polymer G-CSF conjugates were prepared and characterized, and their stabilization properties were evaluated.

4.2 Results and Discussion

Soluble expression and purification of G-CSF was achieved by fusing G-CSF to maltose binding protein (MBP). To help with purification, a polyhistidine-tag was appended to the N- terminus of MBP, and an enterokinase (EK) recognition site (DDDDK) was inserted between

MBP and G-CSF (His6-MBP-GCSF). The designed fusion protein sequence was synthesized and inserted into Saci/BamHI sites of a pMAL-p2E plasmid. The resulting plasmid was then transformed into BL21(DE3) competent E. coli cells. Batch cultures of the transformed E. coli

79 were grown at 30 °C and induced with 1 mM isopropyl β-D-1-thiogalactopyranoside (IPTG) and incubated for an additional 12 hours for optimal overexpression of His6-MBP-GCSF. The cells were collected, lysed, and centrifuged for the collection of the supernatant containing the overexpressed fusion protein.

Purification of the His6-MBP-GCSF fusion protein from the rest of the cell lysate was performed on a nickel(II)-nitrilotriacetic acid (Ni-NTA) column. After loading the crude lysate

(Figure 4-1, lane 2) onto the Ni-NTA column and washing steps, the protein was eluted and the resulting fraction contained the His6-MBP-GCSF protein (Figure 4-1, lane 3); this process removed most of the non-specific proteins. Other than the desired fusion protein, His-tagged

MBP without the attached G-CSF was also included in the fraction. Endotoxin was then removed using an endotoxin removal spin column (Pierce). The level of endotoxins of the purified fusion protein was 0.10 EU/mL.

Figure 4-1. Expression and purification of His6-MBP-GCSF fusion protein following a Ni-NTA column. SDS-PAGE analysis of protein content during initial purification process; lane 1, molecular weight ladder; lane 2, crude lysate after 1 mM IPTG induction; lane 3, purified fraction after Ni-NTA column showing His6-MBP-GCSF (60.8 kDa) and His6-MBP (42 kDa).

The conditions for protease cleavage of the His6-MBP-GCSF fusion protein by EK were optimized (Figure 4-2). Varying concentrations of EK ranging from 0 to 2 enzyme units were evaluated for maximal efficiency to release G-CSF. The expected cleavage products were observed by SDS-PAGE and Western blot analysis showing an increase in the His6-MBP protein and the desired G-CSF protein with increasing EK concentrations. An EK concentration of 0.5 units was selected for large-scale reaction and purification conditions.

Figure 4-2. Enterokinase digestion of His6-MBP-GCSF fusion protein. a) SDS-PAGE analysis; lane 1, molecular weight ladder; lane 2, His6-MBP-GCSF fusion protein without any added EK; lane 3, 0.01 unit of EK; lane 4, 0.1 unit of EK; lane 5, 0.5 unit of EK; lane 6, 1 unit of EK; lane

7, 2 units of EK. b) Western blot analysis; lane 1, 0 units of EK; lane 2, 0.001 unit of EK; lane 3,

0.01 unit of EK; lane 4: 0.1 unit of EK.

After EK digestion of the His6-MBP-GCSF fusion protein, a second Ni-NTA column was then used to selectively bind the His-tagged proteins, allowing the cleaved G-CSF protein to freely elute from the column. Further purification was performed to remove any other contaminating proteins by fast protein liquid chromatography (FPLC). SDS-PAGE analysis

81 shows the protein content following the second Ni-NTA column (Figure 4-3, lane 1), and subsequent FPLC purification of G-CSF (Figure 4-3, lanes 3 and 4).

Figure 4-3. SDS-PAGE analysis of the purification of G-CSF from the His6-MBP-GCSF fusion protein. Lane 1, crude following second Ni-NTA column and prior to FPLC purification; lane 2, molecular weight ladder; lane 3 and 4, fractions of purified G-CSF from FPLC.

The purified G-CSF protein was characterized by MALDI, ELISA, and bioactivity in

NFS-60 cells. The MALDI spectra showed a sharp peak at m/z = 18777.4, confirming the expected G-CSF size of 18.8 kDa (Figure 4-4). The peak at m/z = 37575.7 likely indicates the presence of G-CSF dimer. The concentration of purified G-CSF was determined by ELISA.

Then, the bioactivity of the purified G-CSF protein was assayed by cell proliferation of mouse

NFS-60 myelogenous leukemia lymphoblast cells (Figure 4-5). The dose-response curve was sigmodal and the EC50 of the purified G-CSF protein was 0.23 ng/mL. The results suggest that the purified protein is slightly less active compared to commercially available G-CSF, which has

18 a reported EC50 of 0.1 ng/mL.

Figure 4-4. MALDI spectra of purified G-CSF protein.

1 0 0

)

(

h t

w 5 0

r G

0 0 .0 0 1 0 .0 1 0 .1 1 1 0 1 0 0 C o n c e n tr a tio n (n g /m l)

Figure 4-5. Bioactivity of purified G-CSF in NFS-60 cell proliferation assay.

Scheme 4-1. Structures of trehalose polymers (P1, P2, P3, and P1-benzaldehyde).

Trehalose polymers with polystyrene or polymethacrylate backbones were prepared by free radical polymerization using 2,2-azobis(2-methylpropionitrile) (AIBN) as an initiator as previous described.13 A polystyrene backbone with an acetal linkage to trehalose (P1), polymethacrylate backbone with an acetal linkage to trehalose (P2), and polystyrene backbone with an ether linkage to trehalose (P3) were used (Scheme 4-1). An aldehyde-functionalized P1,

P1-benzaldehyde, was prepared for protein conjugation (shown in Scheme 4-1 and 4-5).

Additionally, a degradable trehalose polymer containing hydrolytically degradable esters in the backbone and trehalose side chains was synthesized. First, reversible addition-fragmentation chain transfer (RAFT) polymerization was performed with a trithiocarbonate chain transfer agent

(CTA), but-3-enyl methacrylate (bMA), 5,6-benzo-2-methylene-1,3-dioxepane (BMDO), and

AIBN to yield P4 (Scheme 4-2) with the respective 1H NMR spectrum shown in Figure 4-6. P4 and protected thiolated trehalose were then coupled via thiol-ene chemistry to prepare P5,

1 followed by deprotection with K2CO3 to yield P6 (p(BMDO-co-trehalose), Scheme 4-3). The H

NMR spectra of the acetylated polymer, P5, and the deprotected trehalose polymer, P6, are shown in Figure 4-7 and Figure 4-8, respectively.

Scheme 4-2. RAFT polymerization of BMDO and bMA.

Figure 4-6. 1H NMR spectrum of P4.

Scheme 4-3. Synthesis of p(BMDO-co-trehalose).

Figure 4-7. 1H NMR spectrum of P5.

Figure 4-8. 1H NMR spectrum of p(BMDO-co-trehalose).

Degradation of p(BMDO-co-trehalose) was performed under basic conditions by incubation in a 5% KOH solution. Complete degradation of the polymer was observed within 3 days as analyzed by gel permeation chromatography (GPC) (Figure 4-9). Degradation into smaller fragments would facilitate the rapid clearance of the polymer and protein-polymer conjugates from the human body.19 The degraded products were then neutralized, dialyzed in water to remove salts and tested for cell cytotoxicity.

Figure 4-9. GPC analysis of p(BMDO-co-trehalose) before and after degradation with 5% KOH.

Other degradation conditions tested for p(BMDO-co-trehalose) included cell culture media (DMEM + 10% FBS), cell-conditioned media, and enzymatic degradation using esterase from porcine liver. No apparent degradation was observed in any of these conditions. The p(BMDO-co-trehalose) was incubated in cell cultured media for up to 6 weeks with no observed degradation. Addition of esterase (2% v/v) from a stock concentration of 16.5 mg/mL and

88 incubation at 37 °C for up to 5 days also showed no degradation effects. The trehalose may have helped protect the polymer against degradation by sterically blocking the enzyme from cleaving the polymer backbone.

To examine the cell compatibility of the trehalose glycopolymers (P1, P2, and P3), NIH

3T3 mouse fibroblasts, RAW 264.7 macrophages, human dermal fibroblasts (HDF), and human umbilical vein endothelial cells (HUVEC) were separately cultured in the presence of each polymer. The results were compared to control wells containing media only. The cells were cultured in 48-well plates at a density of 5 × 103 cells per well. After 24 hours, the culture media was replaced with the working medium containing each polymer up to a concentration of 8 mg/mL. After 48 hours, cell viability was analyzed by LIVE/DEAD reagent. Live and dead cells were counted, and the percent cell viability was calculated by dividing the number of live cells by the total number of cells. All experiments were conducted with four repetitions and averaged.

The results for 1-8 mg/mL for P1, P2, P3, 20 kDa PEG, and trehalose are shown in Figure 4-10.

In each case, the viability of the cells in the presence of the trehalose polymers was the same as no additive, trehalose, and the nontoxic polymer PEG. The cell counts were similar in all cases as well, except for P2, where fewer cells were observed at the 8 mg/mL concentration. This result is important for applications such as stabilization of cells, where lower cell counts and decreases in proliferation would be a disadvantage. The data confirmed that the trehalose glycopolymers are non-cytotoxic to all four tested cell lines.

Figure 4-10. Cytotoxicity assay of P1-P3, 20 kDa PEG, and trehalose with a) NIH 3T3, b) RAW

264.7 macrophages, c) HDF, and d) HUVEC cells, p > 0.05 for all polymers relative to the control. The conditions were repeated four times, and the results are provided as the average with standard deviation relative to the control with no additive.

Additional cytotoxicity assays of the degraded and non-degraded products of p(BMDO- co-trehalose) was carried out in HDF cells by LIVE/DEAD assay and in NFS-60 cells by cell proliferation (Figure 4-11). At 1 mg/mL of degraded p(BMDO-co-trehalose), cell viability was reduced to 74% in HDF cells, while the non-degraded polymer and benzyl trehalose was shown to be non-cytotoxic (Figure 4-11a). Since the benzyl trehalose is non-cytotoxic, the results indicate that the degraded BMDO group, and not the trehalose monomer unit, is most likely to be associated with the observed cytotoxicity. The effects of P1 and p(BMDO-co-trehalose) on NFS-

60 cell proliferation and cell viability was also evaluated at concentrations up to 1 mg/mL

(Figure 4-11b, c). At the highest concentration, the percent cell proliferation relative to no

90 additive control, was reduced to 84% for P1, 60% for non-degraded p(BMDO-co-trehalose), and

38% for degraded p(BMDO-co-trehalose) (Figure 4-11b). When evaluated by LIVE/DEAD assay, P1 exhibited no cytotoxicity (100% cell viability relative to no excipient), p(BMDO-co- trehalose) was 95%, and the degraded p(BMDO-co-trehalose) dropped to 90% at a concentration of 1 mg/mL (Figure 4-11c). Consistent with the cytotoxicity in HDF cells, degraded p(BMDO- co-trehalose) had significantly greater adverse effect on cells compared to the non-degraded polymer, which suggests that the degraded products are cytotoxic. Growth inhibition due to high concentrations of excipients was evident from the data as well. Decreased cell proliferation with no change in percent cell viability indicates that the polymers are inhibiting NFS-60 cell growth.

However, cell growth inhibition was not observed in the adherent cell lines tested above.

Furthermore, the combination of inhibition and cytotoxicity, as shown with p(BMDO-co- trehalose) and the degraded products, resulted in a significant decrease in NFS-60 cell proliferation at higher excipient concentrations.

Figure 4-11. Cytotoxicity assay of p(BMDO-co-trehalose) before and after degradation and benzyl trehalose in a) HDF cells by LIVE/DEAD assay (n = 4), and of the non-degraded, degraded products and P1 in NFS-60 cells based on b) cell proliferation (n = 6), and c) cell viability by LIVE/DEAD assay (n = 4). Results are shown as the average with standard deviation

92 and normalized to 100% to the condition of no excipients. ** = p < 0.01, *** = p < 0.005 relative to no excipients.

Next, the trehalose glycopolymers were evaluated for their ability to stabilize the purified

G-CSF protein to heat stress. G-CSF was heated at 40 °C for 30 minutes with and without excipients and the bioactivity was assayed by NFS-60 cell proliferation. The excipients included

PEG, trehalose, p(BMDO-co-trehalose), and P1, and were screened with weight equivalents from 1 to 500 relative to G-CSF. The resulting G-CSF bioactivities with and without excipients are shown in Figure 4-12. The activity of G-CSF was reduced to about 30% when heated at 40

°C. The addition of PEG did not stabilize G-CSF against heat stress as the activity levels (37% ±

3.8) were similar to the negative control (Figure 4-12a). In the presence of trehalose, 77% ± 7.6 activity was retained at 500 wt. eq. relative to G-CSF, which showed moderate stabilization

(Figure 4-12b). Moderate stabilization of G-CSF was also observed with the addition of p(BMDO-co-trehalose) showing 66% ± 4.6 and 51% ± 5.9 of the original activity with 10 and

500 wt. eq., respectively (Figure 4-12c). P1, especially at 500 wt. eq., showed full retention of activity (Figure 4-12d). In all cases with the trehalose glycopolymers even at the lowest concentration of 1 wt. eq., stabilization with statistical significance was observed compared to the negative control. Additionally, the excipients were added without heating the protein to determine any effects the additive had on the protein and activity. With PEG, trehalose, and p(BMDO-co-trehalose), no significant increase or a decrease in activity was observed. However, the addition of P1 increased G-CSF activity to over 100% relative to the positive control. This result in enhancing activity was observed in previous studies with other proteins, and suggests that the polymer may help with enhancing protein/substrate binding or stabilization of the complex as does trehalose.20

Figure 4-12. Bioactivity of G-CSF without any additive or with 1, 10, 100, or 500 weight equivalents of excipient to protein without heating (untreated) and with heating (treated) to 40 °C for 30 minutes. Excipients shown are a) 20 kDa PEG, b) trehalose, c) p(BMDO-co-trehalose), and d) P1. Data shown as the average (n = 6) and standard deviation. *** = p < 0.005 relative to heated G-CSF control.

The excipient stabilization data and cytotoxicity of the polymers together indicates that

P1 performed the best. Although the p(BMDO-co-trehalose) as excipients helped stabilized G-

CSF against heat stress to some extent, the degradation products demonstrated cytotoxicity at concentrations of 1 mg/mL. Because of the cytotoxicity concerns, the use of the BMDO and trehalose copolymer for protein therapeutics would be less viable. In contrast, the trehalose glycopolymer, P1, stabilized G-CSF against heat stress with full retention of bioactivity and without cytotoxicity in various cell lines up to 8 mg/mL. These results are promising for using

P1, and thus this polymer was investigated further as the target polymer for conjugation with G-

CSF.

Scheme 4-4. Synthesis of benzaldehyde end-functionalized CTA.

Conjugation of G-CSF and P1 was performed via the reductive amination between the N- terminal methionine residue of G-CSF and an aldehyde-functionalized P1 trehalose glycopolymer.21 To prepare the aldehyde end-group trehalose polymer, a benzaldehyde end- functionalized CTA was first synthesized (Scheme 4-4). The reaction involves a one-step EDC coupling of the benzaldehyde and a trithiocarbonate acid.22 Figure 4-13 and Figure 4-14 shows the 1H and 13C NMR spectra, respectively.

1 Figure 4-13. H NMR spectrum of benzaldehyde CTA (CDCl3).

13 Figure 4-14. C NMR spectrum of benzaldehyde CTA (CDCl3).

Scheme 4-5. RAFT polymerization of trehalose monomer with benzaldehyde CTA (P1- benzaldehyde).

P1-benzaldehyde

The synthesized benzaldehyde CTA was used for RAFT polymerization of styrenyl acetal trehalose monomer (Scheme 4-5). The resulting polymer was analyzed by 1H NMR spectroscopy to confirm the presence of an aldehyde peak (Figure 4-15), and then used for conjugation to purified G-CSF protein.

Figure 4-15. 1H NMR spectrum of benzaldehyde end-functionalized trehalose polymer from

RAFT (D6DMSO).

Scheme 4-6. Preparation of G-CSF-P1 conjugate by reductive amination.

P1-benzaldehyde was conjugated to the N-terminal methionine of G-CSF by reductive amination (Scheme 4-6). To test the efficiency of the conjugation, reactions with molar ratios of

10, 50, and 100 of the polymer to G-CSF protein were incubated for 48 hours and then evaluated by silver stained SDS-PAGE analysis (Figure 4-16). The reaction temperature was also tested at both 4 °C (Figure 4-16, lanes 1-3) and 30 °C (Figure 4-16, lanes 5-7). The shift towards higher molecular weight and the band smearing indicates the formation of the G-CSF-P1 conjugate. As expected, increased G-CSF modification occurred at an elevated incubation temperature (30 °C) and with higher equivalents of P1-benzaldehyde. The conjugate was also analyzed by Western blot to confirm that the higher molecular weight band contained G-CSF (Figure 4-17).

Figure 4-16. Silver stained SDS-PAGE analysis of G-CSF and P1-benzaldehyde conjugation with various reaction conditions after 48 hours. Lane 1, 10:1 molar ratio of polymer:protein at 4

°C; lane 2, 50:1 at 4 °C; lane 3, 100:1 at 4 °C; lane 4, molecular weight ladder; lane 5, 10:1 at 30

°C; lane 6, 50:1 at 30 °C; lane 7, 100:1 at 30 °C.

Figure 4-17. Western blot analysis of G-CSF-P1 conjugate.

The stability of the G-CSF-P1 conjugate compared to G-CSF was evaluated following multiple lyophilization cycles (Figure 4-18). After 1 cycle of lyophilization, G-CSF lost 28% of its activity compared to only a 9% activity loss with the G-CSF-P1 conjugate. After five lyophilization cycles, G-CSF retained 56% of its original activity, while the G-CSF-P1 conjugate retained 77% of its original activity. One concern is that there is a possibility that residual free

P1 could still be present in the conjugate and not fully purified, which could also explain the enhanced stability of the G-CSF-P1 conjugate. The trehalose polymer significantly increased the stability of G-CSF to repeated vacuum, drying, and rehydration steps, and greatly reduced the amount of activity loss following each lyophilization cycle.

Figure 4-18. Bioactivity in NFS-60 cell proliferation of G-CSF and G-CSF-P1 conjugate after 5 cycles of lyophilization. G-CSF and G-CSF-P1 conjugate are normalized to untreated samples as

100% bioactivity. n = 6, *** = p < 0.005 between the activities of G-CSF and G-CSF-P1 conjugate.

Next, the heat stability of the G-CSF-P1 conjugate was evaluated. The conjugate was heated starting at 35 °C and ramped 5 °C every 30 minutes with time points sampled to measure bioactivity (Figure 4-19). The protein was heated until full bioactivity was lost to determine the extent of heat stabilization. In all sampled time points, the activity of the G-CSF-P1 conjugate was higher than G-CSF showing that the conjugate exhibited improved heat stability. The activity of native G-CSF dropped to under 10% of its original activity after heat ramping to 45

°C, while the G-CSF-P1 conjugate was able to retain significantly more activity compared to the native G-CSF before decreasing to less than 10% of original activity at 55 °C.

100

Figure 4-19. Bioactivity of G-CSF and G-CSF-P1 conjugate after heat ramping from 35-70 °C, increasing 5 °C every 30 minutes. Activities are normalized to untreated samples as 100% bioactivity, with data shown as the average (n = 6) and standard deviation. *** = p < 0.005 between the activities of G-CSF and G-CSF-P1 conjugate.

The G-CSF-P1 conjugate exhibited improved stability against multiple lyophilization cycles and higher temperatures while maintaining bioactivity as shown in the NFS-60 cell proliferation assay. The bioactivity of the conjugate was significantly higher after five freeze- drying cycles compared to native G-CSF after exposed to only one cycle of lyophilization.

Furthermore, the G-CSF-P1 conjugate maintained bioactivity to higher temperatures compared to the native protein. These results demonstrate the potential of the G-CSF conjugate for future studies.

101

4.3 Conclusions

Trehalose glycopolymers were screened with the goal of stabilizing G-CSF and enhancing the protein’s pharmacokinetic properties. The trehalose glycopolymers included different backbones and a degradable trehalose polymer, p(BMDO-co-trehloase). Cell compatibility of the polymers were evaluated in mouse and human cell lines and P1, P2, and P3 were demonstrated to be non-cytotoxic up to 8 mg/mL. However, the degraded products of p(BMDO-co-trehalose) exhibited cytotoxicity at a lower concentration of 1 mg/mL. Stability screening with the different glycopolymers on G-CSF showed that styrenyl acetal linked trehalose polymer, P1, best stabilized G-CSF against heat compared to the other excipients. The combined results from the polymer cytotoxicity and excipient stability data indicated that P1 would be the best polymer candidate to pursue for protein conjugation. Therefore, P1 with a benzaldehyde functionality was synthesized and conjugated to G-CSF. The resulting G-CSF-P1 conjugate retained higher bioactivity compared to native G-CSF when subjected to heat and lyophilization stressors. It is anticipated that the increased molecular weight due to attachment of the trehalose glycopolymer would similarly increase the in vivo circulation time. Similar to PEG, the attached polymer adds steric bulk to the protein therapeutic and is also expected to reduce the immunological response and degradation of the protein by enzymes. Future work will aim to investigate the pharmacokinetic properties of the G-CSF-P1 conjugate in vivo.

102

4.4 Materials and Methods

Materials

All chemicals and reagents were purchased from Sigma-Aldrich or Fisher Scientific and used as received unless otherwise indicated. SDS-PAGE analysis was performed using Bio-Rad

Any kD Mini-PROTEAN TGX precast protein gels. Enterokinase was purchased from

GenScript. Human G-CSF DuoSET ELISA kit was purchased from R&D Systems. LIVE/DEAD

Viability/Cytotoxicity assay kit and CellTiter-Blue cell viability assay were purchased from

Invitrogen and Promega, respectively. Butenyl methacrylate (bMA) was synthesized as previously described.23 BMDO was synthesized as previously described.24

Constructing Plasmid for His6-MBP-GCSF

The G-CSF gene was modified by appending to the N-terminus with a polyhistidine and maltose binding protein (MBP) tag. An enterokinase recognition site was inserted between MBP and G-CSF to enable digestion and release of G-CSF. The sequence was synthesized and inserted into SacI/BamHI sites of pMAL-p2E (provided by UCLA DOE-Protein Expression

Technology Center). The gene sequence between the SacI (GAGCTC) and BamHI (GGATCC) sites encoding His6-MBP-GCSF is: GAGCTCG CACCACCACCACCACCAC GGAGGAGGA

GATGACGATGACAAGATGACCCCCCTGGGCCCTGCCAGCTCCCTGCCCCAGAGCTTC

CTGCTCAAGTGCTTAGAGCAAGTGAGGAAGATCCAGGGCGATGGCGCAGCGCTCCA

GGAGAAGCTGTGTGCCACCTACAAGCTGTGCCACCCCGAGGAGCTGGTGCTGCTCG

GACACTCTCTGGGCATCCCCTGGGCTCCCCTGAGCAGCTGCCCCAGCCAGGCCCTGC

AGCTGGCAGGCTGCTTGAGCCAACTCCATAGCGGCCTTTTCCTCTACCAGGGGCTCC

TGCAGGCCCTGGAAGGGATCTCCCCCGAGTTGGGTCCCACCTTGGACACACTGCAGC

TGGACGTCGCCGACTTTGCCACCACCATCTGGCAGCAGATGGAAGAACTGGGAATG

103

GCCCCTGCCCTGCAGCCCACCCAGGGTGCCATGCCGGCCTTCGCCTCTGCTTTCCAG

CGCCGGGCAGGAGGGGTCCTGGTTGCCTCCCATCTGCAGAGCTTCCTGGAGGTGTCG

TACCGCGTTCTACGCCACCTTGCCCAGCCCTGATAA GGATCC.

Transformation and Growth of Bacteria

Competent E. coli BL21(DE3) cells were transformed with the constructed plasmid. The plasmid and competent E. coli cells were incubated on ice for 30 min, followed by heat shock in a 42 °C water bath for 30 sec, then transferred onto ice for 2 min. LB broth was added and incubated at 37 °C, 250 rpm for 1 h. Transformed cells were selected on LB agar plates containing ampicillin (50 µg/mL). 1 L of LB medium with 50 µg/mL ampicillin was inoculated with a colony of transformed BL21(DE3) cells at 30 °C, 200 rpm until the optical density at 600 nm (OD600) reached 0.5. IPTG was added to a final concentration of 1 mM and the cultures were grown overnight at 30 °C, 200 rpm. The following day, cultures were centrifuged at 4,000 rpm,

20 min at 4 °C. The supernatant was removed and pellets were stored at -80 °C until lysis.

Purification of His6-MBP-GCSF

Cell pellets were resuspended in lysis buffer (pH 8.0, 0.1 M Tris-HCl and 50 mM NaCl with protease inhibitor tablet EDTA-free). The cells were lysed using a homogenizer

(Emulsiflex) at 15,000 psi for 3 cycles and then centrifuged at 15,000 rpm for 45 min at 4 °C.

The supernatant was filtered through a 0.45 µm cellulose acetate membrane. The resulting solution was subjected through a Ni-NTA column and eluted using the appropriate buffer containing 200 mM imidazole. After elution, endotoxins were removed using an endotoxin removal resin spin column following the manufacturer’s protocol. The His-tagged proteins were

104 concentrated and buffer exchanged using Centriprep 30000 MWCO to pH 5.0, 25 mM sodium acetate buffer.

Purification of G-CSF from His6-MBP-GCSF Fusion Protein

Enterokinase digestion was performed in EK buffer (50 mM Tris-HCl, pH 8.0, 1 mM

CaCl2, 0.1% v/v Tween-20). His6-MBP-GCSF was diluted 1:1 with EK buffer and 20 µL samples were mixed with varying amounts of EK from 0 to 2 units. The samples were incubated at 21 °C for 18 h. Optimal enzyme concentration was determined by SDS-PAGE analysis of the samples, and the large scale enzyme digestion was performed at the optimal EK concentration.

The digested protein solution was run through a Ni-NTA column to collect the initial flow through and to remove any polyhistidine-tagged proteins from the mixture. The collected solution was then concentrated and buffer exchanged using Centriprep (MWCO 10000) to pH

5.0, 25 mM sodium acetate buffer. The concentrate was filtered through a 0.20 micron PTFE membrane and purified by FPLC on a Bio-Rad BioLogic DuoFlow chromatography system equipped with a GE Healthcare Life Sciences size exclusion column (Superdex 75 10/300 GL) in pH 5.0, 25 mM sodium acetate buffer at a flow rate of 0.5 mL/min. The fractions corresponding to G-CSF were collected, concentrated and exchanged into pH 4.0, 10 mM acetic acid buffer, and stored at 4 °C until use.

Concentration Determination of G-CSF (ELISA)

Mouse anti-human G-CSF capture antibody (2 µg/mL in D-PBS) was incubated in 96- well plates for 16 h at 23 °C. The plate was washed between each step with wash buffer. The plate was then blocked with 1% BSA in D-PBS for 2 h at 23 °C. Recombinant G-CSF standards and samples were added to the wells and incubated for 2 h. Biotinylated goat anti-human G-CSF

105 detection antibody (0.25 µg/mL in 1% BSA D-PBS) was added and incubated for 2 h, followed by incubation with streptavidin-HRP (1:200 in 1% BSA D-PBS) for 20 min. The plate was developed with 1-Step™ Ultra TMB solution (Pierce Biotechnology, Rockford) for 4 min prior to adding 1 M H2SO4 as the stop solution. The absorbance signals were measured at 450 nm and the background at 630 nm was subtracted.

G-CSF Bioactivity Evaluation

Activity of purified G-CSF was assayed in a NFS-60 mouse myelogenous leukemia lymphoblast cell line. NFS-60 cells were cultured in RPMI-1640 media supplemented with 10%

FBS and 2 ng/mL interleukin-3 (IL-3). Prior to treating NFS-60 cells with the G-CSF samples,

NFS-60 cells were prepared in RPMI-1640 media with 10% FBS without additional growth factors. The cells were plated in 96-well plates at a density of 20,000 cells/well and a final concentration of G-CSF ranging from 0 to 10 ng/mL. Following 48 h incubation at 37 °C/5%

CO2, CellTiter-Blue viability assay was performed to measure cell proliferation. All experimental groups were normalized to the control of non-treated G-CSF, with a total of six replicates.

Synthesis of p(BMDO-co-trehalose)

Trithiocarbonate CTA (6 mg, 0.02 mmol), bMA (96 mg, 0.76 mmol), BMDO (124 mg,

0.76 mmol), and AIBN (0.63 mg, 0.004 mmol) were added to a dry Schlenk tube. Toluene (1.3 mL) was added to dissolve and the tube was subjected to 4 freeze-pump thaw cycles (down to

100 mTorr) and heated to 80 ºC. After 13.5 h, the polymerization was quenched in liquid N2 and conversion was assessed by 1H NMR. Toluene was removed in vacuo and the crude oil was

1 diluted with CH2Cl2 and precipitated 3x into cold hexanes to yield P4 (86.1 mg). H NMR (500

106

MHz, CDCl3): δ 9.96, 7.92, 7.89, 7.87, 7.25, 7.16, 5.81, 5.14, 5.10, 5.08, 4.06, 3.96, 3.27, 2.62,

1 2.36, 2.13, 1.86, 1.77, 1.45, 1.25, 1.03, 0.97, 0.87, 0.80. Mn ( H NMR) = 4300, Mn (GPC) =

2900, and Ð = 1.76.

P4 (15 mg, 94 µmol allyl groups), thiolated trehalose (305 mg, 467 µmol), DMPA (12 mg, 47 µmol) and THF (0.9 mL) were mixed in a vial with screw cap. The solution was degassed for 10 min and then irradiated with 365 nm light from a handheld UV lamp. After 4 h, the vial was opened and the solution precipitated into 15 mL cold MeOH to yield P5 (54.5 mg). 1H NMR

(500 MHz, CDCl3): δ 9.96, 7.93, 7.24, 7.17, 5.38, 5.24, 5.02, 4.87, 4.16, 3.90, 2.66, 2.58, 2.08,

1 2.04, 2.00, 1.98, 1.95, 1.93, 1.93, 1.92, 1.92, 1.65, 1.59, 1.25, 1.00, 0.78. Mn ( H NMR) = 22900,

Mn (GPC) = 20900, and Ð = 1.22.

P5 (24.6 mg, 0.9 µmol) was dissolved in CHCl3:MeOH 1:1 (2 mL). K2CO3 (26.5 mg,

0.19 mmol, 1 eq. per hydroxyl) was added and stirred at room temperature. After 1.5 h, the solution was diluted with CHCl3, and the precipitate was isolated, diluted with H2O, and dialyzed against 3.5 kDa MWCO in 50% MeOH:H2O, switching to 100% H2O. The purified solution was lyophilized to remove water and yield P6. 1H NMR (500 MHz, D2O+CD3CN): 7.27, 7.19, 5.07,

1 5.04, 3.94, 3.78, 3.72, 3.65, 3.50, 3.31, 3.25, 2.90, 2.57, 1.91, 1.61, 1.15, 0.98, 0.75. Mn ( H

NMR) = 14200, Mn (GPC) = 3300, and Ð = 1.29.

Degradation of p(BMDO-co-trehalose)

p(BMDO-co-trehalose) was dissolved in a 5% KOH solution and incubated over the course of 5 days at 23 °C. Samples were taken at 1, 3, and 5 days and lyophilized. After drying, the samples were dissolved in GPC mobile phase (20% MeCN, 0.3 M NaNO3, 20 mM phosphate buffer), neutralized with concentrated HCl, filtered through a 0.20 µm filter, and analyzed by

GPC.

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Synthesis of Benzaldehyde End-Functionalized Chain Transfer Agent

To the flame-dried flask, 2-(ethyltrithiocarbonate)propionic acid (500 mg, 2.38 mmol) and 4-hydroxybenzaldehyde (377.38 mg, 3.09 mmol) were added and dissolved in DCM (20 mL). The reaction flask was cooled with an ice bath, and EDC (911.38 mg, 4.75 mmol) and

DMAP (58.08 mg, 4.75 × 10-1 mmol) were added. The reaction was stirred at 0 °C for 30 min and stirred at 25 °C for another 2 h. The reaction was washed with H2O three times and the organic layer was collected, dried over MgSO4 and purified by silica gel column chromatography

(EtOAc:Hex = 1:1) to yield 466.7 mg of yellow product (62% yield). 1H NMR (500 MHz in

CDCl3) δ: 9.99 (s, 1H), 7.92-7.90 (d, J = 9.47 Hz, 2H), 7.29-2.27 (d, J = 8.84 Hz, 2H), 5.01-4.97

(q, J = 7.18, 7.40 Hz, 1H), 3.42-3.35 (m, 2H), 1.75-1.73 (d, J = 7.41 Hz, 3H), 1.38-1.35 (t, J =

13 857.62 Hz, 3H), C NMR (500 MHz in CDCl3) δ: 221.7, 190.9, 169.3, 155.2, 134.2, 131.3,

122.2, 47.8, 31.8, 16.3, 13.0. IR: ν = 2974, 2925, 2740, 1755, 1698, 1597, 1501, 1449, 1425,

1376, 1298, 1266, 1202, 1151, 1420, 1066, 1032, 1013, 970, 896, 857, 821 cm-1. HRMS (ESI)

+ calculated for C13H14S3O3Na ([M + Na] ) 337.0003, found 337.0012.

RAFT Polymerization of Trehalose Monomer with CTA (P1-benzaldehyde)

CTA (2.35 mg, 7.47×10-3 mmol), styrenyl acetal trehalose monomer (92.0 mg, 2.02×10-1 mmol), and AIBN (0.49 mg, 2.99×10-3 mmol) were dissolved in 0.25 mL of DMF. The solution underwent four cycles of freeze-pump-thaw and polymerization was initiated by immersing the flask into 80 °C oil bath. The polymerization was stopped at 89% conversion by 1H NMR and

1 purified by dialyzing against H2O (MWCO 3,500 g/mol) to yield P1-benzaldehyde. H NMR

(500 MHz in CDCl3) δ: 9.92, 7.59, 7.15, 6.52, 5.44, 5.17, 4.92, 4.79, 4.37, 4.08, 3.95, 3.75, 3.67,

3.55, 3.46, 3.14, 1.49. Mn = 9,900 g/mol (by GPC), Ð = 1.10.

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Cytotoxicity of Trehalose Polymers

The cell compatibility of the trehalose glycopolymers to NIH 3T3 mouse embryonic fibroblast cells (NIH 3T3, ATCC), RAW 264.7 murine macrophages (RAW 264.7, from

Professor Genhong Cheng), and human dermal fibroblasts (HDFs, Promocell GmbH) were evaluated using a LIVE/DEAD viability/cytotoxicity assay (Invitrogen). Controls were 20 kDa

PEG, trehalose, and buffer only. Cytotoxicity of p(BMDO-co-trehalose), KOH degraded p(BMDO-co-trehalose), and benzyl trehalose was analyzed in HDF cells. NIH 3T3 and RAW

264.7 cells were cultured in Dulbecco’s Modified Eagle Medium (DMEM; Gibco) supplemented with 10% fetal bovine serum (FBS) and 1% penicillin-streptomycin. HDF cells were cultured in fibroblast growth medium (Promocell) containing 2% fetal calf serum (FCS), 1 ng/mL basic fibroblast growth factor (bFGF), 5 µg/mL insulin, and 1% penicillin-streptomycin. HUVECs were cultured in endothelial cell growth medium (Promocell) containing 2% FCS with supplements recommended by the supplier. The cells were seeded in 48-well plates (BD Falcon) at a density of 5 × 103 cells per well. After 24 h, culture media were replaced with 200 µL of the working medium containing known polymer concentrations of 0.1, 0.5, 1, 4, and 8 mg/mL. After incubation for 48 h, the cells were gently washed twice with pre-warmed D-PBS, and stained with the LIVE/DEAD reagent (2 µM calcein AM and 4 µM ethidium homodimer-1). Fluorescent images of each well were captured on an Axiovert 200 microscope with an AxioCam MRm camera and FluoArc mercury lamp. The number of live and dead cells was counted, and percent cell viability was calculated by dividing the number of live cells by the total number of cells. All experiments were conducted with four repetitions. The percent viability was calculated as 100 ×

(number of live cells/total number of cells). The data is provided by normalizing each set to the control without any additives.

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Effects of p(BMDO-co-trehalose), KOH-treated p(BMDO-co-trehalose), and P1 on NFS-

60 cells were evaluated by cell proliferation and LIVE/DEAD assay. The polymers were dissolved in working media containing growth factors (RPMI-1640) + 10% FBS + 2 ng/mL IL-

3) and sterile filtered. NFS-60 cells were plated in 96-well plates at a cell density of 20,000 cells/well. The polymers were diluted and added to the wells at a final concentration of 0, 0.01,

0.1, 0.5, and 1 mg/mL. The plates were incubated at 37 °C/5% CO2. After 48 h incubation,

CellTiter-Blue viability assay was performed to measure cell proliferation. LIVE/DEAD assay was performed as described above.

Preparation of G-CSF-P1 Conjugate

P1-benzaldehyde (24.5 mg, 2.47 µmol) was added to purified G-CSF (200 µg, 0.01

µmol), followed by adding 30 µL of a stock solution of 24 mg/mL NaBH3CN in pH 5.0, 100 mM sodium acetate buffer. The reaction was incubated at 4 °C, with 300 rpm shaking for 48 h.

The conjugate was purified by FPLC and concentrated by Centriprep MWCO 10000 and buffer exchanged with pH 4.0, 10 mM sodium acetate buffer.

Western Blot Analysis of G-CSF and G-CSF-P1 Conjugate

Following standard SDS-PAGE protocols, proteins from the SDS-PAGE gel was transferred to the nitrocellulose membrane for 60 min at 90 V. The membrane was blocked overnight at 4 °C using blocking solution (5% non-fat milk in TBST). The membrane was then incubated with the primary antibody, rabbit anti-G-CSF, diluted 1:1000 in blocking buffer for 4 h. The membrane was washed with TBST 3 times for 10 min each, followed by incubation with the secondary antibody, anti-rabbit HRP IgG diluted 1:2000 in blocking buffer for 2 h, and

110 washed in TBST 3 times for 10 min each. The membrane was developed with SuperSignal West

Pico Chemiluminescent Substrate and imaged using FluorChem FC2 imaging system.

Lyophilization Studies of G-CSF-P1 Conjugate

G-CSF-P1 conjugate and G-CSF protein were diluted in pH 4.0 20 mM sodium acetate buffer to a concentration of 715 ng/mL with a total volume of 100 µL. The samples were frozen with liquid N2, lyophilized, and reconstituted with 100 µL of sterile water. Lyophilization and reconstitution cycles were repeated for a total of 1, 3, or 5 cycles. Lyophilized samples were kept at 4 °C until plating. All samples were reconstituted with 100 µL sterile H2O and further diluted with working media (RPMI-1640 + 10% FBS). NFS-60 cells were plated in a 96-well plate at a cell density of 20,000 cells/well with the final G-CSF concentration at 0.5 ng/mL. The plate was incubated at 37 °C/5% CO2. After 48 h, cell proliferation was assayed by CellTiter-Blue reagent.

Heat Ramping Stability of G-CSF-P1 Conjugate

G-CSF and G-CSF-P1 conjugate were diluted in 10 mM acetic acid buffer and sterile filtered to a concentration of 500 ng/mL. The samples were heat ramped starting at 35 °C and increased 5 °C every 30 min up to a final temperature of 80 °C with 50 µL samples taken at 30 min intervals. NFS-60 cells were plated in a 96-well plate at a cell density of 20,000 cells/well.

The time point samples were diluted with working media (RPMI-1640 + 10% FBS) and added to the 96-well plate at a final concentration of 0.25 ng/mL. The plate was incubated at 37 °C/5%

CO2. After 48 h, cell proliferation was assayed by CellTiter-Blue reagent.

111

4.5 References

(1) Duhrsen, U.; Villeval, J. L.; Boyd, J.; Kannourakis, G.; Morstyn, G.; Metcalf, D. Effects of Recombinant Human Granulocyte Colony-Stimulating Factor on Hematopoietic Progenitor Cells in Cancer Patients. Blood 1988, 72, 2074-2081.

(2) Souza, L. M.; Boone, T. C.; Gabrilove, J.; Lai, P. H.; Zsebo, K. M.; Murdock, D. C.; Chazin, V. R.; Bruszewski, J.; Lu, H.; Chen, K. K.; et al. Recombinant Human Granulocyte Colony-Stimulating Factor: Effects on Normal and Leukemic Myeloid Cells. Science 1986, 232, 61-65.

(3) Molineux, G. The Design and Development of Pegfilgrastim (PEG-rmetHuG- CSF, Neulasta). Curr. Pharm. Des. 2004, 10, 1235-1244.

(4) Morstyn, G.; Foote, M. A.; Walker, T.; Molineux, G. Filgrastim (r-metHuG-CSF) in the 21st Century: SD/01. Acta Haematol. 2001, 105, 151-155.

(5) Hill, C. P.; Osslund, T. D.; Eisenberg, D. The Structure of Granulocyte-Colony- Stimulating Factor and its Relationship to Other Growth Factors. Proc. Natl. Acad. Sci. U.S.A. 1993, 90, 5167-5171.

(6) Harris, J. M.; Chess, R. B. Effect of Pegylation on Pharmaceuticals. Nat. Rev. Drug Discovery 2003, 2, 214-221.

(7) Vogel, C. L.; Wojtukiewicz, M. Z.; Carroll, R. R.; Tjulandin, S. A.; Barajas- Figueroa, L. J.; Wiens, B. L.; Neumann, T. A.; Schwartzberg, L. S. First and Subsequent Cycle Use of Pegfilgrastim Prevents Febrile Neutropenia in Patients with Breast Cancer: A Multicenter, Double-Blind, Placebo-Controlled Phase III Study. J. Clin. Oncol. 2005, 23, 1178- 1184.

(8) Green, M. D.; Koelbl, H.; Baselga, J.; Galid, A.; Guillem, V.; Gascon, P.; Siena, S.; Lalisang, R. I.; Samonigg, H.; Clemens, M. R.; Zani, V.; Liang, B. C.; Renwick, J.; Piccart, M. J.; International Pegfilgrastim 749 Study, G. A Randomized Double-Blind Multicenter Phase III Study of Fixed-Dose Single-Administration Pegfilgrastim Versus Daily Filgrastim in Patients Receiving Myelosuppressive Chemotherapy. Ann. Oncol. 2003, 14, 29-35.

(9) Yang, B. B.; Savin, M. A.; Green, M. Prevention of Chemotherapy-Induced Neutropenia with Pegfilgrastim: Pharmacokinetics and Patient Outcomes. Chemotherapy 2012, 58, 387-398.

(10) Piedmonte, D. M.; Treuheit, M. J. Formulation of Neulasta (pegfilgrastim). Adv. Drug Deliv. Rev. 2008, 60, 50-58.

(11) Ricci, M. S.; Sarkar, C. A.; Fallon, E. M.; Lauffenburger, D. A.; Brems, D. N. pH Dependence of Structural Stability of Interleukin-2 and Granulocyte Colony-Stimulating Factor. Protein Sci. 2003, 12, 1030-1038.

112

(12) Mancini, R. J.; Lee, J.; Maynard, H. D. Trehalose Glycopolymers for Stabilization of Protein Conjugates to Environmental Stressors. J. Am. Chem. Soc. 2012, 134, 8474-8479.

(13) Lee, J.; Lin, E. W.; Lau, U. Y.; Hedrick, J. L.; Bat, E.; Maynard, H. D. Trehalose Glycopolymers as Excipients for Protein Stabilization. Biomacromolecules 2013, 14, 2561-2569.

(14) Jain, A.; Jain, S. K. PEGylation: An Approach for Drug Delivery. A Review. Crit. Rev. Ther. Drug Carrier Syst. 2008, 25, 403-447.

(15) Duncan, R. The Dawning Era of Polymer Therapeutics. Nat. Rev. Drug Discovery 2003, 2, 347-360.

(16) Yang, B. B.; Kido, A. Pharmacokinetics and Pharmacodynamics of Pegfilgrastim. Clinical Pharmacokinetics 2011, 50, 295-306.

(17) Kinstler, O. B. T. O., CA, US), Gabriel, Nancy Elise (Thousand Oaks, CA, US), Farrar, Christine E. (Newbury Park, CA, US), Deprince, Randolph B. (Raleigh, NC, US); Amgen Inc. (Thousand Oaks, CA, US): United States, 2010.

(18) Bai, Y.; Ann, D. K.; Shen, W. C. Recombinant Granulocyte Colony-Stimulating Factor-Transferrin Fusion Protein as an Oral Myelopoietic Agent. Proc. Natl. Acad. Sci. U.S.A. 2005, 102, 7292-7296.

(19) Yamaoka, T.; Tabata, Y.; Ikada, Y. Distribution and Tissue Uptake of Poly(ethylene glycol) with Different Molecular Weights After Intravenous Administration to Mice. J. Pharm. Sci. 1994, 83, 601-606.

(20) Paz-Alfaro, K. J.; Ruiz-Granados, Y. G.; Uribe-Carvajal, S.; Sampedro, J. G. Trehalose-Mediated Thermal Stabilization of Glucose Oxidase from Aspergillus Niger. J. Biotechnol. 2009, 141, 130-136.

(21) Kinstler, O. B.; Brems, D. N.; Lauren, S. L.; Paige, A. G.; Hamburger, J. B.; Treuheit, M. J. Characterization and Stability of N-terminally PEGylated rhG-CSF. Pharm. Res. 1996, 13, 996-1002.

(22) Wood, M. R.; Duncalf, D. J.; Rannard, S. P.; Perrier, S. Selective One-Pot Synthesis of Trithiocarbonates, Xanthates, and Dithiocarbamates for Use in RAFT/MADIX Living Radical Polymerizations. Org. Lett. 2006, 8, 553-556.

(23) Campos, L. M.; Killops, K. L.; Sakai, R.; Paulusse, J. M. J.; Damiron, D.; Drockenmuller, E.; Messmore, B. W.; Hawker, C. J. Development of Thermal and Photochemical Strategies for Thiol-ene Click Polymer Functionalization. Macromolecules 2008, 41, 7063-7070.

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(24) Bailey, W. J.; Ni, Z.; Wu, S. R. Free-Radical Ring-Opening Polymerization of 4,7-Dimethyl-2-Methylene-1,3-Dioxepane and 5,6-Benzo-2-Methylene-1,3-Dioxepane. Macromolecules 1982, 15, 711-714.

CHAPTER 5

TREHALOSE GLYCOPOLYMERS FOR CELL STABILIZATION AND PRESERVATION

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5.1 Introduction

Biological cells and tissues are in high demand for clinical applications, such as transplantations, and hold tremendous promise in therapies and for the treatment of diseases.

Demand for clinically relevant cells requires efficient storage and transportation for transplantation, cell banking, pooling, future use, or ex vivo manipulation purposes.1,2 Specific examples include transplantation of pancreatic islet cells for the treatment of type 1 diabetes and blood components for patient infusions. Biological cells also suffer from similar challenges as proteins regarding stability and viability. Cryopreservation by freezing is a widely applied stress for the long-term storage of cells. DMSO is the most often used cryopreservative. These cryoprotectants work by penetrating the cell membrane and reducing the formation of ice crystals during the cooling process, and thus limiting cell damage.3 However, cryoprotectants typically replace up to 30% of the freezing medium, resulting in significant drawbacks, such as cytotoxicity. In humans/animals, DMSO has been associated with adverse health effects including nausea, dyspnea, and cardiac arrhythmia and even case fatalities.4 Therefore, there could be significant benefits toward the development of alternative preservation agents.

One significant area that would greatly benefit from improved stabilization techniques is in the storage of blood components. The demand for blood components is high since more than

41,000 blood donations happen every day with a total of 30 million blood components transfused every year in the United States.5 The storage of platelets for transfusion is currently limited to five days due to efficacy and safety reasons.6 Due to the high demand of platelets, this short shelf life severely compromises platelet banking and creates shortages. There have been approaches toward increasing the stability and functionality of stored platelets, but these methods involve cryopreservation and lyophilization, the use of alternative additive solutions, and the use of

115 excipients.7-10 Platelet additive solutions, such as PlasmaLyte, T-Sol, and Intersol, are used to replace the role of plasma and provide some buffering capacity, osmotic neutrality, and optimize energy metabolism.11 The formulations of different platelet additive solutions mainly vary in the composition of sodium acetate, sodium citrate, phosphate, glucose, and various salts. Platelets can be cryopreserved using DMSO, but storage with DMSO is not ideal due to its associated adverse health effects and requirement for removal before transfusion.12 Freeze-drying and recovery of platelets has been achieved with platelets loaded with trehalose.13 Excipients that have also been evaluated for platelet preservation include epigallocatechin-3-O-gallate, and uridine diphosphate-galactose.9,14

Investigation into the properties of trehalose and trehalose glycopolymers aims to address these challenges by improving the stability of proteins and biological cells. Trehalose has been used for long-term storage in a variety of cells, including hematopoetic stem cells, red blood cells, and platelets, through loading high amounts of trehalose into the cells.13,15,16 The use of trehalose glycopolymers for cell stabilization was evaluated and presented in this chapter, and we hypothesized that the glycopolymers can minimize damage to cells exposed to stresses, such as cryopreservation. We aimed to expand the usage of the trehalose glycopolymers towards stabilizing cells and expected that lower equivalents of the material would be sufficient to confer stabilization. The polymers could be potentially be utilized as a more efficient and clinically safer alternative for cell stabilization and preservation agents.

Herein, trehalose glycopolymers were used for the purpose of enhancing the stability of cells and three different approaches were evaluated. (1) Covalent attachment of the trehalose to cells. Analogs of N-acetyl-D-mannosamine (ManNAc) with specific functional groups have been demonstrated to be converted through the sialic acid biosynthetic pathway and incorporated into

116 cell surface glycoconjugates when cultured with cells.17,18 This results in the expression of orthogonal functional groups on cell surfaces depending on the ManNAc analog. A ManNAc analog with a ketone functionality was prepared and expressed on cells for conjugation of the trehalose polymers via oxime click chemistry. (2) The polymers were simply added as excipients. (3) Internalization of trehalose glycopolymers by cell permeabilization with streptolysin O (SLO) was investigated. SLO is a thiol-activated, bacterial membrane-damaging protein that has been used to open pores in cell membranes for delivery of proteins up to 100 kDa molecular weight into the cytosol.19 The amount of trehalose glycopolymer uptake in HDF cells was analyzed by fluorescence and by the anthrone assay. Stabilization of HDF cells against cryopreservation and the effects on platelets and E. coli were also evaluated.

5.2 Results and Discussion

Cell surfaces can be modified using analogs of ManNAc, which are internalized, converted and expressed on cell surface glycans.20 N-levulinoyl mannosamine (ManLev) was synthesized following literature procedure.21 ManLev was then incubated with mouse NIH 3T3 fibroblast cells resulting in the expression of accessible ketones on cell surfaces. To verify this modification, the cells were treated with aminooxy-biotin, followed by Alexa Fluor 488 conjugated streptavidin for fluorescent visualization (Figure 5-1). Without the presence of

ManLev, no fluorescence signal was observed (Figure 5-1a). With the addition of ManLev, fluorescent signals was observed (Figure 5-1b), confirming that the cell surface was modified with the ketone functionality.

117

Figure 5-1. Fluorescent micrographs of NIH 3T3 cells incubated with aminooxy-biotin and

Alexa Fluor 488 streptavidin, a) without the addition of ManLev, and b) with ManLev.

Next, an aminooxy end functionalized fluorescent trehalose polymer, Atto590-AO-P3, was prepared (Scheme 5-1). The BOC-protected trithiocarbonate (BOC-AO-P3) was transformed into a free thiol via aminolysis using ethanolamine. The fluorophore, Atto590-maleimide, was added to react with the free thiol (Atto590-BOC-AO-P3). Then, deprotection of the BOC group using 10% TFA was performed to obtain the aminooxy functional group (Atto590-AO-P3).

118

Scheme 5-1. Synthesis of aminooxy-functionalized fluorophore-labeled P3 (Atto590-AO-P3).

The Atto590-AO-P3 was then used to conjugate to the cell surface and verify the

ManLev cell surface modification. After ManLev treatment of cells, Atto590-AO-P3 was incubated with cells for 4 hours and imaged by confocal microscopy (Figure 5-2). Non-specific uptake was observed in cells not treated with ManLev (Figure 5-2b) as indicated by the presence of a fluorescence signal in the cytosol of the cells. However, the fluorescent signals in ManLev- treated cells were higher (Figure 5-2c), which suggested that the ketone-modified cells helped with the localization of the functionalized trehalose polymers towards the cells. These results also suggested that non-specific uptake of the trehalose polymer is a possibility for stabilization that should be further investigated.

119

Figure 5-2. Confocal fluorescent images of NIH 3T3 cells showing DAPI stained nuclei (blue channel) and trehalose polymer (red channel), a) control cells without ManLev or Atto590-AO-

P3, b) Atto590-AO-P3 in untreated cells, and c) Atto590-AO-P3 in ManLev-treated cells.

To evaluate the effectiveness of extracellular trehalose polymer on the stability of cells, two trehalose polymers with different backbones, P1 and P3 (see Chapter 4 for details on these polymers), were screened. Jurkat cells were heated to 50 °C with the addition of P1 or P3 at a concentration up to 1 mg/mL (Figure 5-3). Initially, P3 as an excipient slightly stabilized the cells against heat (80% cell proliferation) compared to the negative control (64% cell proliferation) as shown in Figure 5-3a. However, longer stress times showed no significant difference between all conditions (Figure 5-3b).

120

Figure 5-3. Cell proliferation of Jurkat cells with the addition of P1 and P3 at 0.01, 0.1, and 1 mg/mL following heat treatment at 50 °C for a) 2 min, and b) 3 min. Data shown as the average

(n = 6) and standard deviation. *** = p < 0.005 relative to the heated control.

Freezing and cryopreservation stress was also evaluated with P3 in the presence or absence of DMSO in the freezing media (Figure 5-4). Without DMSO in the freezing media, no cell viability was observed in any condition. After the addition of DMSO, cell proliferation without P3 increased to 55.6% ± 2.1, while addition of P3 at 100 µg/mL showed slightly better cell viability of 65.7% ± 3.1. The data indicates that P3 as a supplement with DMSO could help with improving cell viabilities, but likely will not fully replace DMSO as a cryopreservative agent.

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Figure 5-4. Cell proliferation of Jurkat cells with the addition of P3 at 1, 10, and 100 µg/mL with and without DMSO. Data shown as the average (n = 6) with standard deviation. * = p <

0.05, and *** = p < 0.005 relative to DMSO only.

Stabilization utilizing the trehalose glycopolymer as an excipient with other cell types, including platelets and bacteria, was also examined. The trehalose glycopolymer was added as an excipient and evaluated for platelets stored at 4 °C or 23 °C, as well as bacteria under heat stress

(50 °C). Platelet viability was assessed by measuring the release of lactate dehydrogenase

(LDH).22 This enzyme marker was evaluated for examining the possible preservation of using trehalose or trehalose glycopolymer for the cold storage of platelets. Plasma rich platelets (PRP) isolated from porcine blood were stored with 5 mg/mL trehalose or P3 at 4 °C or at 23 °C (room temperature) with LDH activities measured at specific time points up to seven days (Figure 5-5).

After storage at 4 °C for seven days, the measured LDH activity with no excipients (355 ± 13 mU/mL), with trehalose (373 ± 37 mU/mL), and with P3 (375 ± 7 mU/mL), showed no

122 significant difference upon the addition of trehalose or P3 (Figure 5-5a). However, at room temperature storage (Figure 5-5b), a significant increase in LDH activity was observed, and thus lower platelet viability after 48 hours. After 96 hours, LDH activity was significant higher in the samples with trehalose (963 ± 30 mU/mL) and P3 (1044 ± 9 mU/mL) compared to the control

(807 ± 29 mU/mL), indicating that the excipients enhanced the release of LDH. The LDH activity in the room temperature samples after 48 hours already had higher activities than 4 °C storage after seven days, which was unexpected because it has been reported that platelets remain viable when kept at room temperature conditions (around 22 °C).23,24 There are many other factors that may influence the viability of stored platelets. These include the regulation of pH and gases through the use of specialized storage containers, as well as agitation conditions, which contribute greatly to platelet viability.25,26 Here, PRPs were stored without proper gas regulation, which could explain the unexpected results of increased LDH release and that proper storage conditions also have a large effect on the viability of platelets.

Figure 5-5. LDH activity of porcine platelets with and without excipients, trehalose or P3, stored at a) refrigerated temperature (4 °C), and b) room temperature (23 °C). Data shown as the average (n = 3) with standard deviation. * = p < 0.05, and *** = p < 0.005 relative to PRP only.

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The stabilization of E. coli against heating conditions was evaluated with the addition of trehalose or P3. As a measure of stabilization, cell proliferation was used to determine bacteria survival. E. coli was heated for 30 minutes at 50 °C with trehalose and P3 as excipients

(concentrations ranging from 0.1 to 10 mg/mL) and then growth was monitored by measuring the OD600 (Figure 5-6). Samples heated in the presence of P3 (0.1 mg/mL OD600 = 0.625, 1 mg/mL OD600 = 0.585, 10 mg/mL OD600 = 0.467) had lower OD600 measurements compared to with trehalose (0.1 mg/mL OD600 = 0.655, 1 mg/mL OD600 = 0.668, 10 mg/mL OD600 = 0.636).

Growth was inhibited or the trehalose glycopolymer negatively affected the bacteria during heating compared to trehalose alone. It is important to note that trehalose and P3 were added extracellularly as excipients and that cytosolic trehalose is known to be more effective in cell stabilization.27,28

Figure 5-6. Growth curves of bacteria after heated with trehalose or P3 (0.1 to 10 mg/mL) at 50

°C for 30 minutes.

Attempts to stabilize various cell types with extracellular protection using trehalose glycopolymers against heat stress and replacing DMSO in cryopreservation were unsuccessful.

124

The results suggest that extracellular trehalose may not be sufficient to confer stabilization.

Uptake of trehalose glycopolymers into cells may help with stabilization, as it has been reported in the literature.29 To test this, a bacterial pore-forming agent was used to open pores in cell membranes for loading trehalose glycopolymers intracellularly.

Permeabilization of cells by SLO was optimized by staining using Trypan blue solution.

Cells incubated with activated SLO allows formed pores in their cellular membranes, as indicated by the uptake of Trypan blue (Figure 5-7). SLO readily oxidizes by contact with air and can be reactivated using cysteine or DTT. SLO was incubated with DTT for 2 hours at 37 °C for activation of the enzyme. Cells incubated with non-activated SLO did not allow for uptake of the Trypan blue dye (Figure 5-7c). The concentration of SLO was optimized to determine the right conditions to allow at least 60% of the total cells to be permeabilized (Figure 5-8). The activity of the pore forming bacteria toxin varied depending on the cell type, as shown by the comparison between 3T3 and HDF cells. Furthermore, optimization for each cell type was crucial as higher SLO concentrations would result in cytotoxicity effects on the cells.

Figure 5-7. Phase contrast micrographs of cells stained with Trypan blue solution with a) no

SLO added, b) activated SLO (2,000 ng/mL), and c) non-activated SLO (4,000 ng/mL).

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1 0 0 3T 3

n H D F

o 8 0

i l

i 6 0

a e

4 0

e P

2 0 %

0 5 0 0 0 0 2 5 0 0 0 1 2 5 0 0 1 2

S L O c o n c e n tr a tio n (n g /m L )

Figure 5-8. Percent permeabilization of NIH 3T3 and HDF cells by SLO.

Next, the ability to load trehalose glycopolymers into mammalian cells treated with SLO was examined. To visualize the uptake of the trehalose polymers into the cells, a fluorescent trehalose glycopolymer was synthesized. P3 was polymerized by free radical polymerization in the presence of a rhodamine monomer to yield a fluorescent P3 polymer (P3-Rho, Scheme 5-2).

The respective 1H NMR spectrum is shown in Figure 5-9.

Scheme 5-2. Synthesis of fluorescent P3 by free radical polymerization (P3-Rho).

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Figure 5-9. 1H NMR spectrum of P3-Rho.

Figure 5-10 shows the uptake of P3-Rho in HDF cells with and without permeabilization using SLO. At higher concentrations of the trehalose glycopolymer (500 µg/mL), there was comparable uptake of the polymer into cells regardless of whether or not the cells were permeabilized. The difference in uptake was more noticeable at lower P3 loading concentrations

(50 µg/mL) where more polymer uptake was observed in SLO-permeabilized cells. This is to be expected since high concentrations of polymer are known to permeate into the cytosols of cells.30,31

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Figure 5-10. Uptake of P3-Rho in HDF cells with and without permeabilization by SLO. Phase contrast and fluorescence channels are overlaid as a merged image.

The anthrone method, a colorimetric test for quantification of carbohydrates, was used as an alternative method to quantitate the amount of trehalose glycopolymer loaded into the cells. In this test, sulfuric acid is used to dehydrate the sugar and the resulting furfural compound reacts with the anthrone reagent to produce a colored product. Standard curves of trehalose and P3 of known concentrations were generated by the anthrone assay (Figure 5-11). Lower signals for P3 compared to trehalose at the same concentrations was expected since the trehalose side chains in the polymer is the only group that can react with the anthrone reagent. This explains that a larger amount of P3 is necessary to quantitate the amount of P3 in the samples.

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Figure 5-11. Anthrone assay standard curve generated using trehalose (red circles) and P3 (green squares).

After permeabilizing HDF cells with SLO, trehalose and P3 were separately loaded into cells with initial concentrations ranging from 0 to 10 mg/mL. Intracellular concentrations of trehalose and P3 was evaluated following the anthrone method and non-porated and porated

HDF cells were compared (Figure 5-12). Results of intracellular trehalose concentrations (Figure

5-12a) show that trehalose does not non-specifically enter cells, and permeabilized cells show increased uptake of trehalose as expected. Figure 5-12b shows the intracellular concentration of

P3. P3 was detected in permeabilized cells at a loading concentration of 1 mg/mL, while non- specific uptake was observed at 10 mg/mL for non-permeabilized cells. Consistent with the fluorescent P3 uptake studies, higher concentrations of P3 led to the accumulation of the polymer in cells. However, intracellular concentrations of P3 were generally higher with permeabilized cells compared to non-permeabilized cells in the conditions tested. The discrepancy of the detected polymer concentration between the fluorescent P3 uptake studies and the anthrone assay are likely due to the different conditions used and the sensitivity of the methods. The anthrone

129 assay is less sensitive as it requires higher concentrations for detection. Furthermore, the polymer uptake studies were evaluated on adhered cells compared to detached HDF cells, so the cell densities are different. Regardless of the difference, the results between the two methods are consistent with each other in the general trends observed for P3 uptake.

Figure 5-12. Intracellular concentration as determined by anthrone assay in HDF cells when loaded at a range of 0 to 10 mg/mL of a) trehalose and b) P3.

The effects of extracellular and intracellular P3 on cryopreservation of HDF cells were evaluated. HDFs were permeabilized with SLO and loaded with P3 concentrations ranging from

0 to 10 mg/mL, which were then cryopreserved in preservation media containing DMSO with and without P3 (Figure 5-13). The results show the cell viabilities from LIVE/DEAD staining of

HDF cells without extracellular P3 (Figure 5-13a), and with 2 mg/mL extracellular P3 (Figure 5-

13b). Due to the variability of the results, no significant difference or enhancement in cryopreservation stability was observed with increased P3 loading or a combination of intracellular and extracellular trehalose glycopolymers. Many have reported that trehalose loadings as high as 0.2 M intracellularly and 0.5 M extracellularly were required to aid in cryopreservation.29,32,33 This suggests that a possible reason is that the intracellular P3 concentration may not be high enough to confer stabilization against cryopreservation.

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Figure 5-13. Percent cell viability of cryopreserved HDF cells with or without intracellular P3 and in preservation media (cryo-SFM) a) without extracellular P3 and b) with extracellular 2 mg/mL P3.

One particular issue was the compromise between the amount of loading into the cells and cytotoxicity due to the SLO porating agent. The effects of increased SLO concentration and thus increased permeabilization of HDF cells on viability after cryopreservation were evaluated

(Figure 5-14). Visualized by LIVE/DEAD staining, increased SLO concentration on the cells led to decreased viability and fewer cells overall following cryopreservation. This was expected as

SLO is a known bacterial toxin and has been shown to be cytotoxic to a variety of cell lines.34,35

Increased permeabilization of HDF cells would result in a higher capacity for P3 loading, however, a compromise had to be made to balance loading and the cytotoxic effects.

Cytotoxicity from the trehalose glycopolymer is not likely because it was already determined that the trehalose glycopolymers are non-cytotoxic to four different cells lines, including HDF cells, up to a concentration of 8 mg/mL (see Chapter 4). In this case, utilizing SLO may not be an ideal approach for intracellular uptake of the trehalose glycopolymers as there are limitations in the amount that can be loaded. SLO has not been used for the purpose of loading trehalose or other protective agents for enhancing cell preservation. Other techniques that have been

131 evaluated, such as loading with a genetically engineered pore forming toxin29 or electroporation,36 would be potential alternatives to test with the trehalose glycopolymers.

Figure 5-14. Effects of SLO concentration ranging from 0 to 750 ng/mL on cell viability of cryopreserved HDF cells.

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5.3 Conclusions

Loading trehalose glycopolymers intracellularly into mammalian cells was investigated for the purpose of enhancing stability against cryopreservation and eliminating the need for toxic preservation agents. Cell permeabilization by the bacteria toxin, streptolysin O, was utilized and optimized to balance between cytosolic loading and toxicity of the cells. A fluorescent trehalose polymer, P3-Rho, was synthesized and uptake was shown in HDF cells. This result was supported with the colorimetric anthrone assay to specifically measure the amount of trehalose inside the cells. Cryopreservation studies with intra- and extra-cellularly loaded trehalose glycopolymer showed no significant improvements in cell viability at the tested concentrations.

Possible reasons include that the system of loading trehalose has not been previously evaluated with SLO and the cytotoxic effects of SLO were an issue. This can lead to insufficient intracellular loading into the cells. The trehalose glycopolymers as excipients were also evaluated to stabilize Jurkat cells, platelets and E. coli cells. The addition of the trehalose polymers was not sufficient to provide stabilization, and suggests that other factors, such as higher internal loading, need to be considered. Other techniques, such as microinjection or electroporation, which would enhance and ensure the cytosolic uptake of trehalose glycopolymer are potential approaches for future work.

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5.4 Materials and Methods

Materials

ManLev was synthesized following literature procedures.21 Lactate Dehydrogenase

Activity Assay Kit was purchased from Sigma-Aldrich. Streptolysin O from Streptococcus pyogenes was purchased from Sigma-Aldrich. RIPA lysis and extraction buffer was purchased from ThermoFisher Scientific. Cryo-SFM was obtained from Promocell, LIVE/DEAD

Viability/Cytotoxicity Kit was purchased from Invitrogen. All other chemicals and reagents were purchased from Sigma-Aldrich or Fisher Scientific unless noted otherwise.

Cell Surface Modification

Mouse embryonic fibroblast cells (NIH 3T3, ATCC) were cultured in Dulbecco’s

Modified Eagle Medium (DMEM, Gibco) supplemented with 10% FBS and 1% penicillin- streptomycin. Cells were seeded in 48-well plates at a density of 4.5 × 103 cells per well. The cells were supplemented with a sterile-filtered 100 mM solution of ManLev in PBS to a final

ManLev concentration of 10 mM. Following incubation with ManLev for 72 h at 37 °C, 5%

CO2, the cells were washed two times with ligation buffer (PBS, pH 6.5 5% FBS). The cells were then incubated with 2.5 mM aminooxy-biotin in ligation buffer for 4 h at 37 °C. The cells were washed twice with a labeling buffer (PBS pH 7.4, 0.1% FBS, 0.1% NaN3), followed by incubation with Alexa Fluor 488-streptavidin for 20 min in the dark. The cells were washed twice and fluorescent images were captured on an Axiovert 200 microscope with AxioCam

MRm camera and FluoArc mercury lamp.

Fluorophore (Atto590) Labeling of Aminooxy P3 (Atto590-AO-P3)

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BOC-AO-P3 was prepared using a BOC-protected aminooxy chain transfer agent with an acetylated trehalose monomer using RAFT polymerization and subsequent deprotection of the acetate groups using NaOMe. BOC-AO-P3 (1.06 mg, 1.23 × 10-4 mmol) was dissolved in

-3 degrassed H2O (0.4 mL). Ethanol diamine (0.15 µL in 0.1 mL H2O, 2.49 × 10 mmol) was added and the reaction was stirred for 20 min at 40 °C under Ar (g). The reaction was cooled

-4 down to 25 °C and Atto590-maleimide (0.5 mg in 0.2 mL H2O, 6.15 × 10 mmol) was added.

Then the reaction was stirred at 25 °C for 12 h under Ar (g) in the dark. The mixture was dialyzed against H2O (MWCO 3,500 g/mol) and lyophilized until use.

Cell Surface Modification with Atto590-AO-P3

Mouse NIH 3T3 cells were seeded on autoclaved coverslips in 24-well plates at a density of 1.35 × 104 cells per well. When appropriate, the cells were supplemented with a sterile-filtered

100 mM solution of ManLev in PBS to a final concentration of 10 mM. Following incubation with ManLev for 72 h at 37 °C, the cells were washed three times with ligation buffer. Cells were then incubated with Atto590-AO-P3 in ligation buffer for 4 h at 37 °C. After being washed three times with buffer, the cells were fixed in 4% formaldehyde in PBS for 15 min at 24 °C. The cells were washed and permeabilized with 0.2% Triton-X in PBS for 10 min at 24 °C. The cells on coverslips were mounted using Vectashield mounting medium with DAPI. Samples were analyzed and visualized using fluorescent confocal microscopy.

Cell Proliferation Studies

Jurkat cells were cultured in RPMI-1640 supplemented with 10% FBS and 1% penicillin- streptomycin. In a 2 mL Eppendorf tube, Jurkat cells were added at a density of 5 × 105 cells/mL and 0, 10, 100, or 1000 µg/mL of P1 or P3 in a total volume of 200 µL. The samples were heated

135 at 50 °C for 2 or 3 min, followed by placing the vial on ice for cooling. The cells were plated in a

96-well plate at a 1:4 dilution with working media, and incubated at 37 °C. After 72 h, cell proliferation was measured using CellTiter Blue assay.

For cryopreservation studies, Jurkat cells were prepared at a cell density of 1 × 106 cells/mL with 0, 1, 10, and 100 µg/mL P3 in both conditions with and without 5% DMSO. The cells with P3 were added to cryovials and cooled to -80 °C at a rate of -1 °C/min and transferred to a liquid N2 tank for storage. The cells were thawed at 37 °C and plated in 96-well plate at a 1:4 dilution of the sample to working media and incubated at 37 °C. After 72 h, cell proliferation was measured using CellTiter Blue assay.

Free Radical Polymerization of Styrenyl Ether Trehalose Monomer with Methacryloxyethyl Rhodamine (P3-Rho)

AIBN (0.44 mg, 2.85 µmol) and styrenyl ether trehalose monomer (48.67 mg, 0.11 mmol) were dissolved in a 1:2 mixture of DMF/H2O. The oxygen was removed by five cycles of freeze-pump-thaw and the reaction was stirred at 80 °C for 30 h. In a separate flask, methacryloxyethyl rhodamine (0.186 mg, 0.27 µmol) was dissolved in DMF and subjected to three cycles of freeze-pump-thaw and added to the polymerization flask. The reaction continued at 80 °C for 7 h. Reaction was stopped by immersing into liquid nitrogen, dialyzed against H2O for 3 days (MWCO 3,500 g/mol), and lyophilized to obtain a pink powder. Mn (GPC) = 3.2 kDa and Ð = 1.10.

Streptolysin O Permeabilization

SLO was activated by incubation in a solution of 10 mM DTT in a buffer of Hank’s

Balanced Salt Solution (HBSS) and 20 mM HEPES (H-HBSS). A 20 mM DTT solution in H-

HBSS was added to an equal volume of SLO to bring to a final concentration of 10 mM DTT

136 and 20,000 ng/mL SLO. The solution was incubated in a 37 °C water bath for 2 h prior for activation. NIH 3T3 cells or HDF cells were trypsinized and washed 3 times with HBSS. The cells were diluted to 1 × 106 cells/mL (3T3) or 5 × 105 cells/mL (HDF) in H-HBSS. Percent permeabilization of cells was determined via Trypan blue staining. A sample of 90 µL of cells and 10 µL of activated SLO of different concentrations were incubated at 37 °C for 15 min prior to addition of equal volume of Trypan blue solution. The cells were visualized using a hemocytometer and counted for permeabilized cells (blue-stained cells), and non-permeabilized cells (no Trypan blue uptake).

P3-Rho Uptake in HDF Cells

HDFs were plated in 24-well plates (500 µL of 700,000 cells/mL) and incubated for 24 h to adhere to plate. The cells were washed 3 times with HBSS (no calcium, no magnesium) and then added H-HBSS (non-porated condition) or activated SLO (500 ng/mL) in H-HBSS (porated condition). The plate was incubated at 37 °C for 15 min. A solution of P3-Rho was prepared in

H-HBSS and added to each cell condition to a final concentration of 0, 50, or 500 µg/mL, and incubated at room temperature for 20 min. The wells were washed 2 times with HBSS and then imaged.

Intracellular Trehalose and P3 Concentration Determination

The intracellular concentration of trehalose and P3 was evaluated by the anthrone method. HDF cells were trypsinized, washed 3 times with HBSS, and adjusted to 5 x 105 cells/mL in H-HBSS. The cells were incubated with H-HBSS or with activated SLO for 15 min at 37 °C, followed by incubation with trehalose (0 or 10 mg/mL), or P3 solution (0, 1, or 10 mg/mL). The cells were incubated for 45 min, followed by addition of ice cold 20 mM CaCl2

137

HBSS for 20 min. Each sample was centrifuged at 2,500 x g for 5 min, washed two times with cold buffer. The cells were lysed with minimal RIPA buffer and shook for 15 min, followed by pelleting at 14,000 x g for 15 min. The supernatant was collected for the anthrone assay. In a 96- well plate, 50 µL of the supernatant was added into a well. A 2 mg/mL solution of anthrone reagent in concentrated sulfuric acid was prepared, 100 µL was added to each well, prior to heating at 95 °C for 10 min, and then cooled to room temperature for the color to develop. The absorbance was read at 630 nm. The samples were analyzed to a standard curve of trehalose or

P3 to determine the intracellular concentration.

Cryopreservation and Cell Viability Assay

Following P3 loading into HDF cells, the cells were resuspended in cryopreservation media (cryo-SFM) or with cryo-SFM with 2 mg/mL P3 (extracellular P3). The solutions were transferred to cryovials and cooled -1 °C/min at -80 °C for 6.5 h. Afterwards, the cryovials were transferred to a liquid nitrogen tank for storage until plating. Following storage, the cryovials were thawed in a 37 °C water bath and diluted into 5 mL of working media. 500 µL of each condition was plated in a 24-well plate and incubated at 37 °C/5% CO2 for 24 h prior to cell viability assay.

Fluorescence Microscopy

Cell viability was analyzed by the LIVE/DEAD reagent (2 µM calcein AM and 4 µM ethidium homodimer-1). The cells were visualized by fluorescence microscopy using a Zeiss

Axiovert 200 fluorescent microscope equipped with an AxioCam MRm monochrome camera, and pictures were acquired and processed using AxioVision LE 4.6 software. The number of live

138 and dead cells was counted, and percent cell viability was calculated by dividing the number of live cells by the total number of cells.

Isolation of Porcine Platelets

Porcine blood was kindly provided by Sandra Duarte-Vogel, DVM from UCLA DLAM.

The blood was collected into sodium citrate tubes. To isolate the plasma rich platelets (PRP), the porcine blood was centrifuged at 160 × g for 20 min, and the top portion was collected as the

PRP.

Lactate Dehydrogenase Assay for Platelet Viability

Storage of PRP was compared at conditions of 4 °C and 23 °C, with trehalose, P3 trehalose polymer, or no additive. The final concentration of the excipients was 5 mg/mL. PRP was kept in vented-cap 50-mL tubes and on a shaker table at 70 rpm at 4 °C or 23 °C for up to 7 days and analyzed at 0, 8, 24, 48, 96, and 168 h. To assay for LDH activity, 100 µL of PRP was diluted with 400 µL of cold LDH assay buffer. The solution was centrifuged at 10,000 × g for 15 min at 4 °C, and the supernatant was collected for analysis. LDH activity was measured in a 96- well plate format following the manufacturer’s protocol. Briefly, the collected supernatant was mixed with LDH Substrate Mix and absorbance at 450 nm was measured every 5 min, and LDH activity was calculated by (amount NADH generated × dilution factor) / (reaction time × volume).

Bacteria Heat Studies

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A colony of ampicillin-resistant E. coli was grown in LB media containing 50 µg/mL ampicillin in a vented tube. At an OD600 of 0.5, the bacteria was diluted 1:1 in LB media containing excipients, trehalose or P3, at a final concentration of 0.1, 1, or 10 mg/mL. The samples were heated in 50 °C water bath for 30 min. Following heat stress, 500 µL of the solution was added to 20 mL of fresh LB media and incubated at 37 °C and 200 rpm. Cell growth was monitored by measuring the absorbance at 600 nm.

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